Nanotechnology for Nucleic Acid Delivery: Methods and Protocols [2nd ed.] 978-1-4939-9091-7, 978-1-4939-9092-4

Table of contents :
Front Matter ....Pages i-xv
Sequence-Defined Cationic Lipo-Oligomers Containing Unsaturated Fatty Acids for Transfection (Sören Reinhard, Ernst Wagner)....Pages 1-25
Synthesis of Bioreducible Polycations with Controlled Topologies (Ye-Zi You, Jun-Jie Yan, Fei Yu, Zhi-Qiang Yu, David Oupicky)....Pages 27-38
Histidine-Rich Cationic Cell-Penetrating Peptides for Plasmid DNA and siRNA Delivery (Antoine Kichler, A. James Mason, Arnaud Marquette, Burkhard Bechinger)....Pages 39-59
Covalent Fluorophore Labeling of Oligonucleotides and Generation of Other Oligonucleotide Bioconjugates (Cornelia Lorenzer, Johannes Winkler)....Pages 61-72
Synthetic Oligonucleotides in SPECT/CT In Vivo Imaging: Chemical Modifications, In111 Complex Formation, Incorporation into Drug Delivery Systems (Maxim Antopolsky)....Pages 73-82
Synthesis of Polyethylenimine-Based Nanocarriers for Systemic Tumor Targeting of Nucleic Acids (Wolfgang Rödl, Alexander Taschauer, David Schaffert, Ernst Wagner, Manfred Ogris)....Pages 83-99
Cationic Photopolymerized Polydiacetylenic (PDA) Micelles for siRNA Delivery (Manon Ripoll, Patrick Neuberg, Jean-Serge Remy, Antoine Kichler)....Pages 101-122
Lipids for Nucleic Acid Delivery: Cationic or Neutral Lipoplexes, Synthesis, and Particle Formation (Michel Bessodes, Helene Dhotel, Nathalie Mignet)....Pages 123-139
Preparation, Characterization, and In Vitro Evaluation of Lipidoid–Polymer Hybrid Nanoparticles for siRNA Delivery to the Cytosol (Kaushik Thanki, Xianghui Zeng, Camilla Foged)....Pages 141-152
Layer-by-Layer Assembled Nanoparticles for siRNA Delivery (Michaela Guter, Miriam Breunig)....Pages 153-160
Layer-By-Layer Film Engineering for Sequential Gene Delivery (Lingxiao Xie, Yi Zou, Sean Carroll, Maria Muniz, Guangzhao Mao)....Pages 161-176
Surface- and Hydrogel-Mediated Delivery of Nucleic Acid Nanoparticles (Angela K. Pannier, Tyler Kozisek, Tatiana Segura)....Pages 177-197
In Situ AFM Analysis Investigating Disassembly of DNA Nanoparticles and Nanofilms (Yi Zou, Lei Wan, Jenifer Blacklock, Lingxiao Xie, Sean Carroll, David Oupicky et al.)....Pages 199-209
Lyophilization of Synthetic Gene Carriers (Julia Christina Kasper, Sarah Hedtrich, Wolfgang Friess)....Pages 211-225
Firefly Luciferase-Based Reporter Gene Assay for Investigating Nanoparticle-Mediated Nucleic Acid Delivery (Katharina Müller, Manfred Ogris, Haider Sami)....Pages 227-239
Enhancing Nucleic Acid Delivery with Ultrasound and Microbubbles (Heleen Dewitte, Silke Roovers, Stefaan C. De Smedt, Ine Lentacker)....Pages 241-251
Magnetic and Acoustically Active Microbubbles Loaded with Nucleic Acids for Gene Delivery (Dialechti Vlaskou, Olga Mykhaylyk, Christian Plank)....Pages 253-290
Lactate Dehydrogenase Assay for Assessment of Polycation Cytotoxicity (Ladan Parhamifar, Helene Andersen, S. Moein Moghimi)....Pages 291-299
Combined Fluorimetric Caspase-3/7 Assay and Bradford Protein Determination for Assessment of Polycation-Mediated Cytotoxicity (Anna K. Larsen, Arnaldur Hall, Henrik Lundsgart, S. Moein Moghimi)....Pages 301-311
Determination of Polycation-Mediated Perturbation of Mitochondrial Respiration in Intact Cells by High-Resolution Respirometry (Oxygraph-2k, OROBOROS) (Arnaldur Hall, S. Moein Moghimi)....Pages 313-322
Evaluating the Regulation of Cytokine Levels After siRNA Treatment in Antigen-Specific Target Cell Populations via Intracellular Staining (Rima Kandil, Daniel Feldmann, Yuran Xie, Olivia M. Merkel)....Pages 323-331
Anti-PEG IgM Production via a PEGylated Nanocarrier System for Nucleic Acid Delivery (Amr S. Abu Lila, Tatsuhiro Ishida)....Pages 333-346
Near-Infrared Optical Imaging of Nucleic Acid Nanocarriers In Vivo (Claire Rome, Julien Gravier, Marie Morille, Gilles Divita, Anne-Laure Bolcato-Bellemin, Véronique Josserand et al.)....Pages 347-363
Flow Cytometry-Based Cell Type-Specific Assessment of Target Regulation by Pulmonary siRNA Delivery (Olivia M. Merkel, Leigh M. Marsh, Holger Garn, Thomas Kissel)....Pages 365-375
Microbubbles for Nucleic Acid Delivery in Liver Using Mild Sonoporation (Nathalie Mignet, Corinne Marie, Anthony Delalande, Simona Manta, Michel-Francis Bureau, Gilles Renault et al.)....Pages 377-387
Lipopeptide Delivery of siRNA to the Central Nervous System (Mark D. Zabel, Luke Mollnow, Heather Bender)....Pages 389-403
Back Matter ....Pages 405-406

Citation preview

Methods in Molecular Biology 1943

Manfred Ogris Haider Sami Editors

Nanotechnology for Nucleic Acid Delivery Methods and Protocols Second Edition

METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences, University of Hertfordshire, Hatfield, Hertfordshire AL10 9AB, UK

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

Nanotechnology for Nucleic Acid Delivery Methods and Protocols Second Edition

Edited by

Manfred Ogris Laboratory of MacroMolecular Cancer Therapeutics (MMCT), Department of Pharmaceutical Chemistry, Center of Pharmaceutical Sciences, Faculty of Life Sciences, University of Vienna, Vienna, Austria Center for NanoScience (CeNS), Ludwig-Maximilians-University, Munich, Germany

Haider Sami Laboratory of MacroMolecular Cancer Therapeutics (MMCT), Department of Pharmaceutical Chemistry, Center of Pharmaceutical Sciences, Faculty of Life Sciences, University of Vienna, Vienna, Austria

Editors Manfred Ogris Laboratory of MacroMolecular Cancer Therapeutics (MMCT) Department of Pharmaceutical Chemistry Center of Pharmaceutical Sciences Faculty of Life Sciences University of Vienna Vienna, Austria

Haider Sami Laboratory of MacroMolecular Cancer Therapeutics (MMCT) Department of Pharmaceutical Chemistry Center of Pharmaceutical Sciences Faculty of Life Sciences University of Vienna Vienna, Austria

Center for NanoScience (CeNS) Ludwig-Maximilians-University Munich, Germany

ISSN 1064-3745 ISSN 1940-6029 (electronic) ISBN 978-1-4939-9091-7 ISBN 978-1-4939-9092-4 (eBook) https://doi.org/10.1007/978-1-4939-9092-4 Library of Congress Control Number: 2019931016 © Springer Science+Business Media, LLC, part of Springer Nature 2019 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, express 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 Press imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 233 Spring Street, New York, NY 10013, U.S.A.

Preface Nanotechnology-enabled nucleic acid delivery is a rapidly evolving field and offers a multitude of possibilities for addressing the challenges faced by nucleic acid therapies. Since the release of the first edition of this volume in 2013, several new technologies have emerged, and existing protocols have further improved. Within this edition, we aim at updating the existing chapters and at the same time adding new topics to offer an up-to-date view on technologies used for nucleic acid delivery. We have organized the contents of the current edition according to the sequence in which technologies are employed to develop formulations for nucleic acid delivery, starting with chemical synthesis of carriers, nanoformulations, their characterization in vitro and in vivo, and their application in preclinical models. The first three chapters deal with carrier synthesis. In Chap. 1, So¨ren Reinhard and colleagues provide a new chapter on the synthesis of defined cationic lipid-oligomers for plasmid and siRNA transfection. David Oupicky continues with biodegradable and disulfide-crosslinked polymers based on oligo- and polycations in Chap. 2. Antoine Kichler and colleagues describe synthesis and application of cationic, histidine-rich peptides for nucleic acid delivery (Chap. 3). Direct covalent modification of oligonucleotides is described in the next two chapters. Johannes Winker and colleagues, in Chap. 4, present a method on the generation of oligonucleotide bioconjugates by attaching fluorophores to the 30 or 50 end via click chemistry. Maxim Antopolski provides detailed protocol on the covalent attachment of radionuclides for SPECT/CT in vivo imaging via end-term attached chelators in Chap. 5. Wolfgang Ro¨dl et al. update the synthesis protocols for peptide-polyethylenimine conjugates in Chap. 6. The new Chap. 7 by Antoine Kichler and colleagues deals with a micellar delivery system for siRNA delivery, which can be stabilized by photopolymerization. Nathalie Mignet and co-workers present a protocol for the synthesis of cationic and neutral lipids and their particle formation with nucleic acids in Chap. 8. Lipidoids are a new class of transfection reagents. In Chap. 9, Kaushik Thanki et al. describe synthesis of a lipidoid-polymer-based formulation for siRNA delivery to cytosol. Coating of nanoparticles with successive layers of polymer and nucleic acid allows formation of a well-defined, size-controlled delivery formulation. In Chap. 10, Miriam Breunig and colleagues describe layer-by-layer technique for siRNA delivery and also include targeting ligands. Guangzhao Mao and co-workers apply layer-by-layer technique using biodegradable polymers for coating surfaces for sequential gene delivery (Chap. 11). Tatiana Segura and colleagues report a technology where polyplexes are incorporated into hydrogels for surface coating, which then allows for transfection of cells grown on top of the layer (Chap. 12). Analysis of nanoparticles by atomic force microscopy (AFM) gives insights not only into particle size but also surface topology. Guanzhao Mao and colleagues describe the application of AFM techniques for studying disassembly of DNA nanoparticles and nanofilms in Chap. 13. For further preclinical and clinical development, gene delivery formulations with long-term storage stability are desirable and offer ease of usage. Wolfgang Friess and colleagues in Chap. 14 present a lyophilization protocol for optimal freezing with cryoprotectant and freeze-drying of polyplexes. In Chap. 15, our lab presents a robust and cost-effective protocol for the measurement of luciferase reporter gene expression for investigating nanoparticle-mediated delivery of nucleic acids. Local activation of nanoformulations and enhancement of nucleic acid delivery in vivo can be achieved by

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several methods to achieve stimuli-based delivery. Ine Lentacker and colleagues apply ultrasound-activated microbubbles to enhance local nucleic acid delivery (Chap. 16). Olga Mykhaylyk and colleagues describe in Chap. 17 the synthesis and in vitro evaluation of magnetic nanoparticles and microbubbles for nucleic acid delivery, which can be targeted with magnetic force and activated by ultrasound. Biocompatibility and toxicity are decisive parameters for nanoformulations and the final fate of nucleic acid cargo, both on the cellular level and within an organism. Both the carrier system and the delivered nucleic acid itself can trigger side effects, which can also influence the therapeutic effect of nucleic acids. In three chapters (Chaps. 18–20), Moein Moghimi and colleagues provide detailed protocols for a quantitative evaluation of polycationmediated cytotoxicity. Lactate dehydrogenase (LDH) is released after cell rupture, being an early event in necrosis and a late event in apoptosis. Chapter 18 gives a protocol for a fast and accurate assay for LDH release, which can be applied prior to more detailed toxicological studies. Tracking onset of apoptosis gives insights into the mechanism of nanoparticle or polymer-induced toxicity. In Chap. 19, a fluorimetric assay for detection of caspase activity is reported. For polycations, a distinct mitochondrial toxicity has been observed, which is supposed to be due to perturbation of membrane integrity. In Chap. 20, Arnaldur Hall and Moein Moghimi present a protocol on measuring mitochondrial bioenergetics by highresolution respirometry. Both the innate and adaptive immune system can play a role in the immunological response when an organism is exposed to exogenous nucleic acids and carrier systems. The intrinsic effect of siRNA on cytokine induction in lung cells (after pulmonary delivery of siRNA) is analyzed in Chap. 21 by Olivia Merkel and colleagues, presenting a protocol for detecting intracellular cytokines after sorting of immune cells using antibody-coated magnetic beads. Although coating with polyethylene glycol (PEG) can improve biocompatibility and blood circulation of nanoparticles, antibodies against PEG can neutralize this effect. Tatsuro Ishida et al. in Chap. 22 provide a protocol for quantifying the anti-PEG induction in vivo to predict the blood clearance of PEGylated formulations after repeated administration. Investigation of the biodistribution of nucleic acid after in vivo administration is imperative for desirable therapeutic interventions. Near-infrared imaging using fluorophores allows real-time tracking of nucleic acid delivery in vivo. JeanLuc Coll and colleagues describe labeling of nanoparticles with appropriate near-infrared fluorophores and their detection in vivo (Chap. 23). The three final chapters include protocols for tissue-directed plasmid and siRNA delivery: In Chap. 24, Olivia Merkel and colleagues apply siRNA by intratracheal instillation into the lung and provide a protocol on identification of distinct cell types in the lung that take up siRNA. In Chap. 25, Nathalie Mignet et al. use cationic microbubbles in combination with ultrasound to allow efficient transgene expression in the liver using optimized small plasmids. In Chap. 26, Mark Zabel and colleagues describe preparation and application of lipopeptide formulation to achieve siRNA delivery to the central nervous system. Taken together, the second version of Nanotechnology for Nucleic Acid Delivery deals with not only already established methods but also emerging aspects in this field. A broad set of topics is covered ranging from chemical synthesis of macromolecules and bioconjugates; novel and established nanoformulations; characterization of these nanoformulations for biophysical, biological, and toxicological aspects; and also protocols dealing with application and imaging of such carrier systems in vivo. Vienna, Austria

Manfred Ogris Haider Sami

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

1 Sequence-Defined Cationic Lipo-Oligomers Containing Unsaturated Fatty Acids for Transfection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . So¨ren Reinhard and Ernst Wagner 2 Synthesis of Bioreducible Polycations with Controlled Topologies . . . . . . . . . . . . Ye-Zi You, Jun-Jie Yan, Fei Yu, Zhi-Qiang Yu, and David Oupicky 3 Histidine-Rich Cationic Cell-Penetrating Peptides for Plasmid DNA and siRNA Delivery. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Antoine Kichler, A. James Mason, Arnaud Marquette, and Burkhard Bechinger 4 Covalent Fluorophore Labeling of Oligonucleotides and Generation of Other Oligonucleotide Bioconjugates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cornelia Lorenzer and Johannes Winkler 5 Synthetic Oligonucleotides in SPECT/CT In Vivo Imaging: Chemical Modifications, In111 Complex Formation, Incorporation into Drug Delivery Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Maxim Antopolsky 6 Synthesis of Polyethylenimine-Based Nanocarriers for Systemic Tumor Targeting of Nucleic Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Wolfgang Ro¨dl, Alexander Taschauer, David Schaffert, and Ernst Wagner, and Manfred Ogris 7 Cationic Photopolymerized Polydiacetylenic (PDA) Micelles for siRNA Delivery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Manon Ripoll, Patrick Neuberg, Jean-Serge Remy, and Antoine Kichler 8 Lipids for Nucleic Acid Delivery: Cationic or Neutral Lipoplexes, Synthesis, and Particle Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Michel Bessodes, Helene Dhotel, and Nathalie Mignet 9 Preparation, Characterization, and In Vitro Evaluation of Lipidoid–Polymer Hybrid Nanoparticles for siRNA Delivery to the Cytosol . . . . . . . . . . . . . . . . . . . . Kaushik Thanki, Xianghui Zeng, and Camilla Foged 10 Layer-by-Layer Assembled Nanoparticles for siRNA Delivery . . . . . . . . . . . . . . . . Michaela Guter and Miriam Breunig 11 Layer-By-Layer Film Engineering for Sequential Gene Delivery . . . . . . . . . . . . . . Lingxiao Xie, Yi Zou, Sean Carroll, Maria Muniz, and Guangzhao Mao 12 Surface- and Hydrogel-Mediated Delivery of Nucleic Acid Nanoparticles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Angela K. Pannier, Tyler Kozisek, and Tatiana Segura

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In Situ AFM Analysis Investigating Disassembly of DNA Nanoparticles and Nanofilms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yi Zou, Lei Wan, Jenifer Blacklock, Lingxiao Xie, Sean Carroll, David Oupicky, and Guangzhao Mao Lyophilization of Synthetic Gene Carriers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Julia Christina Kasper, Sarah Hedtrich, and Wolfgang Friess Firefly Luciferase-Based Reporter Gene Assay for Investigating Nanoparticle-Mediated Nucleic Acid Delivery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ¨ ller, Manfred Ogris, and Haider Sami Katharina Mu Enhancing Nucleic Acid Delivery with Ultrasound and Microbubbles. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Heleen Dewitte, Silke Roovers, Stefaan C. De Smedt, and Ine Lentacker Magnetic and Acoustically Active Microbubbles Loaded with Nucleic Acids for Gene Delivery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Dialechti Vlaskou, Olga Mykhaylyk, and Christian Plank Lactate Dehydrogenase Assay for Assessment of Polycation Cytotoxicity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ladan Parhamifar, Helene Andersen, and S. Moein Moghimi Combined Fluorimetric Caspase-3/7 Assay and Bradford Protein Determination for Assessment of Polycation-Mediated Cytotoxicity. . . . . . . . . . . Anna K. Larsen, Arnaldur Hall, Henrik Lundsgart, and S. Moein Moghimi Determination of Polycation-Mediated Perturbation of Mitochondrial Respiration in Intact Cells by High-Resolution Respirometry (Oxygraph-2k, OROBOROS). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Arnaldur Hall and S. Moein Moghimi Evaluating the Regulation of Cytokine Levels After siRNA Treatment in Antigen-Specific Target Cell Populations via Intracellular Staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rima Kandil, Daniel Feldmann, Yuran Xie, and Olivia M. Merkel Anti-PEG IgM Production via a PEGylated Nanocarrier System for Nucleic Acid Delivery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Amr S. Abu Lila and Tatsuhiro Ishida Near-Infrared Optical Imaging of Nucleic Acid Nanocarriers In Vivo . . . . . . . . . Claire Rome, Julien Gravier, Marie Morille, Gilles Divita, Anne-Laure Bolcato-Bellemin, Ve´ronique Josserand, and Jean-Luc Coll Flow Cytometry-Based Cell Type-Specific Assessment of Target Regulation by Pulmonary siRNA Delivery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Olivia M. Merkel, Leigh M. Marsh, Holger Garn, and Thomas Kissel

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Microbubbles for Nucleic Acid Delivery in Liver Using Mild Sonoporation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377 Nathalie Mignet, Corinne Marie, Anthony Delalande, Simona Manta, Michel-Francis Bureau, Gilles Renault, Daniel Scherman, and Chantal Pichon Lipopeptide Delivery of siRNA to the Central Nervous System. . . . . . . . . . . . . . . 389 Mark D. Zabel, Luke Mollnow, and Heather Bender

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

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Contributors AMR S. ABU LILA  Department of Pharmacokinetics and Biopharmaceutics, Institute of Biomedical Sciences, Tokushima University, Tokushima, Japan; Department of Pharmaceutics and Industrial Pharmacy, Faculty of Pharmacy, Zagazig University, Zagazig, Egypt; Department of Pharmaceutics, College of Pharmacy, Hail University, Hail, Saudi Arabia HELENE ANDERSEN  Centre for Pharmaceutical Nanotechnology and Nanotoxicology, University of Copenhagen, Copenhagen, Denmark MAXIM ANTOPOLSKY  Teaching and Scientific Consulting, FGBPOU “Medical College”, Moscow, Russian Federation BURKHARD BECHINGER  Institut de Chimie, CNRS, UMR7177, Universite´ de Strasbourg, Strasbourg, France HEATHER BENDER  Department of Microbiology, Immunology and Pathology, Prion Research Center, College of Veterinary Medicine and Biomedical Sciences, Colorado State University, Fort Collins, CO, USA MICHEL BESSODES  Unite´ de Technologies Chimiques et Biologiques pour la Sante´, INSERM, U 1022, Paris, France; CNRS, UMR 8258, Paris, France; Faculte´ de Pharmacie, Sorbonne Paris Cite´, Universite´ Paris Descartes, Paris, France; Chimie ParisTech, PSL Research University, Paris, France JENIFER BLACKLOCK  Department of Biomedical Engineering, Wayne State University, Detroit, MI, USA ANNE-LAURE BOLCATO-BELLEMIN  Polyplus-Transfection, Illkirch, France MIRIAM BREUNIG  Department of Pharmaceutical Technology, University of Regensburg, Regensburg, Germany MICHEL-FRANCIS BUREAU  INSERM, U1022, Paris, France; CNRS, UMR8258, Paris, France; Faculte´ de Pharmacie, Sorbonne Paris Cite´, Universite´ Paris Descartes, Paris, France; Chimie ParisTech, PSL Research University, Paris, France SEAN CARROLL  Department of Chemical Engineering and Materials Science, Wayne State University, Detroit, MI, USA; Department of Biomedical Engineering, Wayne State University, Detroit, MI, USA JEAN-LUC COLL  INSERM U823, Equipe 5, Institut Albert Bonniot, Grenoble, France; Universite´ Joseph Fourier, Grenoble, France; INSERM UGA U1209, CNRS 5309, Institute For Advanced Biosciences, La Tronche, France STEFAAN C. DE SMEDT  Ghent Research Group on Nanomedicines, Laboratory of General Biochemistry and Physical Pharmacy, Faculty of Pharmaceutical Sciences, Ghent University, Ghent, Belgium ANTHONY DELALANDE  Centre de Biophysique Mole´culaire and Universite´ d’Orle´ans, CNRS-UPR 4301, Orle´ans, France HELEEN DEWITTE  Ghent Research Group on Nanomedicines, Laboratory of General Biochemistry and Physical Pharmacy, Faculty of Pharmaceutical Sciences, Ghent University, Ghent, Belgium HELENE DHOTEL  Unite´ de Technologies Chimiques et Biologiques pour la Sante´, INSERM, U 1022, Paris, France; CNRS, UMR 8258, Paris, France; Faculte´ de Pharmacie, Sorbonne

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Contributors

Paris Cite´, Universite´ Paris Descartes, Paris, France; Chimie ParisTech, PSL Research University, Paris, France GILLES DIVITA  CNRS UMR5237, Montpellier, France DANIEL FELDMANN  Department of Pharmaceutical Sciences, Wayne State University, Detroit, MI, USA CAMILLA FOGED  Department of Pharmacy, Faculty of Health and Medical Sciences, University of Copenhagen, Copenhagen, Denmark WOLFGANG FRIESS  Department of Pharmacy, Pharmaceutical Technology and Biopharmaceutics, Ludwig-Maximilians-Universit€ a t, Munich, Germany HOLGER GARN  Institute of Laboratory Medicine and Pathobiochemistry—Molecular Diagnostics, Philipps-Universit€ a t Marburg, Marburg, Germany; Sterna Biologicals GmbH & Co. KG, Marburg, Germany JULIEN GRAVIER  CEA Grenoble/LETI-DTBS, Grenoble, France MICHAELA GUTER  Department of Pharmaceutical Technology, University of Regensburg, Regensburg, Germany ARNALDUR HALL  Genome Integrity Unit, Danish Cancer Society Research Center, Copenhagen, Denmark SARAH HEDTRICH  Institute for Pharmacy, Pharmacology & Toxicology, Freie Universit€ at Berlin, Berlin, Germany TATSUHIRO ISHIDA  Department of Pharmacokinetics and Biopharmaceutics, Institute of Biomedical Sciences, Tokushima University, Tokushima, Japan VE´RONIQUE JOSSERAND  INSERM U823, Equipe 5, Institut Albert Bonniot, Grenoble, France; Universite´ Joseph Fourier, Grenoble, France; INSERM UGA U1209, CNRS 5309, Institute For Advanced Biosciences, La Tronche, France RIMA KANDIL  Department of Pharmaceutical Technology and Biopharmaceutics, LudwigMaximilians-Universit€ at Mu¨nchen, Munich, Germany JULIA CHRISTINA KASPER  Bioprocess and Pharmaceutical Development Biologicals, Boehringer Ingelheim Pharma GmbH and Co. KG, Biberach, Germany ANTOINE KICHLER  Laboratoire de Conception et Application de Mole´cules Bioactives, LabEx Medalis, Faculte´ de Pharmacie, CNRS, UMR7199, Universite´ de Strasbourg, Illkirch, France THOMAS KISSEL  Department of Pharmaceutics and Biopharmacy, Philipps-Universit€ at Marburg, Marburg, Germany TYLER KOZISEK  Department of Biological Systems Engineering, University of NebraskaLincoln, Lincoln, NE, USA ANNA K. LARSEN  Centre for Pharmaceutical Nanotechnology and Nanotoxicology, University of Copenhagen, Copenhagen, Denmark INE LENTACKER  Ghent Research Group on Nanomedicines, Laboratory of General Biochemistry and Physical Pharmacy, Faculty of Pharmaceutical Sciences, Ghent University, Ghent, Belgium CORNELIA LORENZER  Department of Pharmaceutical Chemistry, University of Vienna, Vienna, Austria HENRIK LUNDSGART  Centre for Pharmaceutical Nanotechnology and Nanotoxicology, University of Copenhagen, Copenhagen, Denmark SIMONA MANTA  INSERM, U1022, Paris, France; CNRS, UMR8258, Paris, France; Faculte´ de Pharmacie, Sorbonne Paris Cite´, Universite´ Paris Descartes, Paris, France; Chimie ParisTech, PSL Research University, Paris, France

Contributors

xiii

GUANGZHAO MAO  Department of Chemical Engineering and Materials Science, Wayne State University, Detroit, MI, USA CORINNE MARIE  INSERM, U1022, Paris, France; CNRS, UMR8258, Paris, France; Faculte´ de Pharmacie, Sorbonne Paris Cite´, Universite´ Paris Descartes, Paris, France; Chimie ParisTech, PSL Research University, Paris, France ARNAUD MARQUETTE  Institut de Chimie, CNRS, UMR7177, Universite´ de Strasbourg, Strasbourg, France LEIGH M. MARSH  Institute of Laboratory Medicine and Pathobiochemistry—Molecular Diagnostics, Philipps-Universit€ a t Marburg, Marburg, Germany; Ludwig Boltzmann Institute for Lung Vasculature Research, Graz, Austria A. JAMES MASON  Institute of Pharmaceutical Science, King’s College London, London, UK OLIVIA M. MERKEL  Department of Pharmaceutical Technology and Biopharmaceutics, Ludwig-Maximilians-Universit€ at Mu¨nchen, Munich, Germany; Department of Pharmaceutical Sciences, Wayne State University, Detroit, MI, USA; Department of Pharmacy, Ludwig-Maximilians-Universit€ a t Mu¨nchen, Munich, Germany; Department of Pharmaceutics and Biopharmacy, Philipps-Universit€ a t Marburg, Marburg, Germany NATHALIE MIGNET  Unite´ de Technologies Chimiques et Biologiques pour la Sante´, INSERM, U 1022, Paris, France; CNRS, UMR 8258, Paris, France; Faculte´ de Pharmacie, Sorbonne Paris Cite´, Universite´ Paris Descartes, Paris, France; Chimie ParisTech, PSL Research University, Paris, France S. MOEIN MOGHIMI  School of Pharmacy, Newcastle University, Newcastle upon Tyne, UK; Division of Stratified Medicine, Biomarkers and Therapeutics, Faculty of Medical Sciences, Institute of Cellular Medicine, Newcastle University, Newcastle upon Tyne, UK LUKE MOLLNOW  Department of Microbiology, Immunology and Pathology, Prion Research Center, College of Veterinary Medicine and Biomedical Sciences, Colorado State University, Fort Collins, CO, USA MARIE MORILLE  INSERM U646, Angers, France KATHARINA MU¨LLER  Laboratory of MacroMolecular Cancer Therapeutics (MMCT), Department of Pharmaceutical Chemistry, Center of Pharmaceutical Sciences, Faculty of Life Sciences, University of Vienna, Vienna, Austria MARIA MUNIZ  Department of Biomedical Engineering, Wayne State University, Detroit, MI, USA OLGA MYKHAYLYK  Institute of Molecular Immunology and Experimental Oncology, Klinikum Rechts der Isar, Technische Universit€ a t Mu¨nchen, Munich, Germany PATRICK NEUBERG  Laboratoire de Conception et Application de Mole´cules Bioactives, LabEx Medalis, Faculte´ de Pharmacie, CNRS, UMR7199, Universite´ de Strasbourg, Illkirch, France MANFRED OGRIS  Laboratory of MacroMolecular Cancer Therapeutics (MMCT), Department of Pharmaceutical Chemistry, Center of Pharmaceutical Sciences, Faculty of Life Sciences, University of Vienna, Vienna, Austria; Center for NanoScience (CeNS), Ludwig-Maximilians-University, Munich, Germany DAVID OUPICKY  Department of Pharmaceutical Sciences, Wayne State University, Detroit, MI, USA; University of Nebraska Medical Center, Omaha, NE, USA ANGELA K. PANNIER  Department of Biological Systems Engineering, University of NebraskaLincoln, Lincoln, NE, USA LADAN PARHAMIFAR  Centre for Pharmaceutical Nanotechnology and Nanotoxicology, University of Copenhagen, Copenhagen, Denmark

xiv

Contributors

CHANTAL PICHON  Centre de Biophysique Mole´culaire and Universite´ d’Orle´ans, CNRSUPR 4301, Orle´ans, France CHRISTIAN PLANK  Institute of Molecular Immunology and Experimental Oncology, Klinikum Rechts der Isar, Technische Universit€ a t Mu¨nchen, Munich, Germany SO¨REN REINHARD  Department of Pharmacy, Pharmaceutical Biotechnology, Center of Nanoscience (CeNS), Ludwig-Maximilians-Universit€ at Butenandtstr, Mu¨nchen, Germany JEAN-SERGE REMY  Laboratoire de Conception et Application de Mole´cules Bioactives, LabEx Medalis, Faculte´ de Pharmacie, CNRS, UMR7199, Universite´ de Strasbourg, Illkirch, France GILLES RENAULT  INSERM, U1016, Institut Cochin, Paris, France; CNRS, UMR8104, Paris, France; Sorbonne Paris Cite´, Universite´ Paris Descartes, Paris, France MANON RIPOLL  Laboratoire de Conception et Application de Mole´cules Bioactives, LabEx Medalis, Faculte´ de Pharmacie, CNRS, UMR7199, Universite´ de Strasbourg, Illkirch, France WOLFGANG RO¨DL  Pharmaceutical Biotechnology, Center for System Based Drug Research, Ludwig-Maximilians-University, Munich, Germany CLAIRE ROME  INSERM U823, Equipe 5, Institut Albert Bonniot, Grenoble, France; Universite´ Joseph Fourier, Grenoble, France; Grenoble Institute of Neuroscience, Grenoble, France SILKE ROOVERS  Ghent Research Group on Nanomedicines, Laboratory of General Biochemistry and Physical Pharmacy, Faculty of Pharmaceutical Sciences, Ghent University, Ghent, Belgium HAIDER SAMI  Laboratory of MacroMolecular Cancer Therapeutics (MMCT), Department of Pharmaceutical Chemistry, Center of Pharmaceutical Sciences, Faculty of Life Sciences, University of Vienna, Vienna, Austria DAVID SCHAFFERT  Department of Molecular Biology, Aarhus University, Aarhus, Denmark DANIEL SCHERMAN  INSERM, U1022, Paris, France; CNRS, UMR8258, Paris, France; Faculte´ de Pharmacie, Sorbonne Paris Cite´, Universite´ Paris Descartes, Paris, France; Chimie ParisTech, PSL Research University, Paris, France TATIANA SEGURA  Department of Biomedical Engineering, Duke University, Durham, NC, USA; Neurology and Dermatology, Duke University School of Medicine, Durham, NC, USA ALEXANDER TASCHAUER  Laboratory of MacroMolecular Cancer Therapeutics (MMCT), Department of Pharmaceutical Chemistry, Center of Pharmaceutical Sciences, Faculty of Life Sciences, University of Vienna, Vienna, Austria KAUSHIK THANKI  Department of Pharmacy, Faculty of Health and Medical Sciences, University of Copenhagen, Copenhagen, Denmark DIALECHTI VLASKOU  Institute of Molecular Immunology and Experimental Oncology, Klinikum Rechts der Isar, Technische Universit€ a t Mu¨nchen, Munich, Germany ERNST WAGNER  Department of Pharmacy, Pharmaceutical Biotechnology, Center of Nanoscience (CeNS), Ludwig-Maximilians-Universit€ at Butenandtstr, Mu¨nchen, Germany; Nanosystems Initiative Munich (NIM), Schellingstr, Mu¨nchen, Germany; Center for NanoScience (CeNS), Ludwig-Maximilians-University, Munich, Germany LEI WAN  Department of Chemical Engineering and Materials Science, Wayne State University, Detroit, MI, USA

Contributors

xv

JOHANNES WINKLER  Department of Pharmaceutical Chemistry, University of Vienna, Vienna, Austria; Department of Cardiology, Medical University of Vienna, Vienna, Austria LINGXIAO XIE  Department of Chemical Engineering and Materials Science, Wayne State University, Detroit, MI, USA YURAN XIE  Department of Pharmaceutical Sciences, Wayne State University, Detroit, MI, USA JUN-JIE YAN  CAS Key Lab of Soft Matter Chemistry, Department of Polymer Science and Engineering, University of Science and Technology of China, Hefei, China YE-ZI YOU  CAS Key Lab of Soft Matter Chemistry, Department of Polymer Science and Engineering, University of Science and Technology of China, Hefei, China FEI YU  University of Nebraska Medical Center, Omaha, NE, USA ZHI-QIANG YU  CAS Key Lab of Soft Matter Chemistry, Department of Polymer Science and Engineering, University of Science and Technology of China, Hefei, China MARK D. ZABEL  Department of Microbiology, Immunology and Pathology, Prion Research Center, College of Veterinary Medicine and Biomedical Sciences, Colorado State University, Fort Collins, CO, USA XIANGHUI ZENG  Department of Pharmacy, Faculty of Health and Medical Sciences, University of Copenhagen, Copenhagen, Denmark YI ZOU  Department of Chemical Engineering and Materials Science, Wayne State University, Detroit, MI, USA

Chapter 1 Sequence-Defined Cationic Lipo-Oligomers Containing Unsaturated Fatty Acids for Transfection So¨ren Reinhard and Ernst Wagner Abstract Sequence-defined cationic lipo-oligomers containing unsaturated fatty acids are potent nucleic acid carriers that are produced by solid-phase supported synthesis. However, the trifluoroacetic acid (TFA)-mediated removal of acid-labile protecting groups and cleavage from the resin can be accompanied by side products caused by an addition of TFA to the double bonds of unsaturated fatty acids. These TFA adducts are converted into hydroxylated derivatives under aqueous conditions. Here we describe an optimized cleavage protocol (precooling cleavage solution to 4
C, 20 min cleavage at 22
C), which minimizes TFA adduct formation, retains the unsaturated hydrocarbon chain character, and ensures high yields of the synthesis. Key words Nucleic acid delivery, Polyplexes, Sequence-defined, Solid phase synthesis, Oleic acid

1

Introduction Therapeutic nucleic acids like plasmid DNA (pDNA), microRNA, and small interfering RNA (siRNA) offer exciting opportunities for clinical applications [1–3]. While pDNA has its destination in the nucleus, RNA interference is initiated by cytosolic delivery of microRNA or synthetic siRNA [4, 5]. Efficient delivery of nucleic acids, however, is a major bottleneck for successful medical development [6, 7]. Nucleic acids are negatively charged and far larger than conventional drugs. In addition, they are rapidly cleared from the bloodstream and degraded. Therefore, various extracellular and intracellular barriers in the nucleic acid delivery pathway need to be addressed by formulation with dynamic and bioresponsive carriers [8–10]. Nucleic acids assemble into lipo-polyplexes upon mixing with cationic lipids or lipo-oligomers, driven by intermolecular electrostatic and hydrophobic interactions. Such cationic amphipathic carriers provide high polyplex stability and transfection efficacy [11–15]. Minor variations in the different subunits (polar head group, hydrophobic tails, linkages) can significantly influence the

Manfred Ogris and Haider Sami (eds.), Nanotechnology for Nucleic Acid Delivery: Methods and Protocols, Methods in Molecular Biology, vol. 1943, https://doi.org/10.1007/978-1-4939-9092-4_1, © Springer Science+Business Media, LLC, part of Springer Nature 2019

1

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So¨ren Reinhard and Ernst Wagner

bioactivity of cationic lipids [16, 17]. The cis-unsaturated fatty acid (FA) oleic acid was incorporated in well-known cationic lipids like DOPE, DOTMA, and DOSMA, which were first used to efficiently delivery DNA and mRNA in 1987 and 1989 respectively [18, 19]. Lipid membrane disruption or fusion events as mechanisms of endosomal escape play a crucial role for efficient release of nucleic acids into the cytosol. Cationic lipids containing cis-unsaturated FAs can destabilize membrane lipids by interacting with anionic phospholipids [20]. Cone-shaped cationic lipids, where the crosssectional area of the hydrophilic head groups is smaller than that of the hydrophobic tails, promote such fusion events [21–23]. Double bonds or other modifications of hydrophobic units broaden the conical shapes by increasing their steric demands and fluidity [24]. Defined chemical structures of nucleic acid carriers facilitate the establishment of structure–activity relationships and consequently their rational design and optimization [25, 26]. Solidphase assisted synthesis (SPS) is a convenient option to produce a variety of oligomers for screening by shuffling functional domains in a precisely defined way [27–35]. Polyethylenimine (PEI) was discovered in 1995 as a very potent cationic transfection polymer, which is commonly used in various topologies and modifications [36–38]. The aminoethylene motif contained in PEI has been incorporated into synthetic solid-phase compatible building blocks such as succinoyl-tetraethylene-pentamine (Stp) to introduce proton sponge activity in sequence-defined oligocations [39]. In combination with hydrophobic elements and disulfide-forming cysteines, potent nucleic acid carriers have been designed [40–42]. Especially incorporation of the cis-unsaturated C18 fatty acids oleic acid and linoleic acid as hydrophobic moieties has proven beneficial for nucleic acid delivery. The ability of such nanocarriers to lyse membranes is pH-dependent and most pronounced at pH 5.5, which benefits cell compatibility at physiological neutral pH and promotes efficient endosomal escape upon vesicular acidification [31, 40, 41]. The bioresponsive lytic activity is based on a synergy between the pH-responsive protonation of the oligoamino acid Stp, which increases the cationic character for endosomal membrane binding, and the diacyl moieties, which provide amphipathic character. Both properties seem to be required for cellular membrane disruption [43]. Cationic carriers containing lipids and amino acids can be produced by solid-phase-assisted synthesis or in solution and have successfully been applied for delivery of pDNA, siRNA and miRNA in vitro or in vivo [24, 30, 31, 35, 44–53]. If unsaturated fatty acids like oleic acid or linoleic acid are used in SPS, the trifluoroacetic acid (TFA)-mediated cleavage of acid-labile protecting groups and the oligomer from the resin is accompanied by side reactions caused by the addition of TFA to double bonds of alkenes [51, 54–62]. TFA esters of fatty acids are readily hydrolyzed in neutral or basic aqueous solution, generating hydroxylated

Defined Lipo-Oligomers Containing Unsaturated Fatty Acids

3

Fig. 1 Reaction mechanism of TFA addition to double bonds and subsequent TFA ester hydrolysis. Reproduced from ([51] Reinhard et al., ChemMedChem, 2017) with permission from Wiley

hydrocarbon chains (Fig. 1). Side products derived from TFA addition can drastically lower the yield of the synthesis or, if not purified properly, affect the properties of the nucleic acid carrier. The reaction kinetics of the addition of TFA to the monounsaturated oleic acid (C18:1) during TFA-mediated cleavage was investigated and the cleavage protocol was optimized regarding temperature and time (see Subheading 3.1.3, steps 9–12). By that method, the amount of side products was minimized, while complete cleavage of acid-labile protecting groups and liberation of the oligomer from the resin was assured [51]. T-shape lipo-oligomers (OleA-t, LinA-t) containing unsaturated oleic acid (C18:1) or linoleic acid (C18:2) and analogous structures containing the saturated or modified hydrophobic C18 moieties stearic acid (C18:0), and hydroxystearic acid were synthesized (Fig. 2). The lipo-oligomers OleA-t, OH-SteA-t, and SteA-t were evaluated regarding their nucleic acid delivery characteristics such as siRNA binding (see Note 1) and lytic potential (see Note 2 and Fig. 3a) of the oligomers and size (see Note 3), gene silencing efficacy (see Note 4 and Fig. 3b) and cytotoxicity (see Note 5) of the formed siRNA lipo-polyplexes. In conclusion, OleA-t showed particularly favorable pH dependency of endosomolytic activity, efficient gene silencing and excellent cell tolerability compared to its counterparts. Analogous studies with the linoleic acid (C18:2) containing lipo-oligomer LinA-t confirmed both the possibility of TFA adduct formation and the measures to avoid them. In this methodical chapter, the optimized synthesis of lipo-oligomers containing unsaturated fatty acids with focus on TFA-mediated cleavage of the oligomer and subsequent purification is reported in order to retain the unsaturated hydrocarbon chain character and to ensure high yields of the synthesis.

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Fig. 2 Sequence-defined lipo-oligomers with T-shape topology. Schematic overview of the structures with different modifications (C: cysteine, Y: tyrosine, K: lysine, Stp: succinoyl-tetraethylene-pentamine). Fmoc-Stp(Boc3)-OH is the protected form of the building block Stp, which can be used in SPS. The wavy lines represent amide linkages. Compare ([51] Reinhard et al., ChemMedChem, 2017)

2

Materials

2.1 Solid-Phase Assisted Synthesis (SPS)

2.1.1 Amino Acids and Other Building Blocks

Use solvents and reagents of high quality (e.g., peptide grade) for all experiments (see Note 6). The SPS can be carried out manually on an overhead shaker using microreactors (see Note 7) with polyethylene filters (Multisyntech GmbH, Witten Germany) or with an automated peptide synthesizer (Syro WaveTM, Biotage, Uppsala, Sweden). As solid support, 2-chlorotrityl chloride resin (Iris Biotech, Marktredewitz, Germany) is used. 1. Fmoc and Boc-protected α-amino acids (Iris Biotech, Marktredwitz, Germany). 2. Fmoc-Stp(Boc3)-OH (synthesis described in [63]). 3. Oleic acid, stearic acid, linoleic acid (Sigma-Aldrich, Munich, Germany).

Defined Lipo-Oligomers Containing Unsaturated Fatty Acids

5

Fig. 3 (a) Erythrocyte leakage triggered by lipo-oligomers at different concentrations and pH values. Negative control (hemoglobin release from PBS-treated erythrocytes) was set to 0%. Triton X-100 treatment served as positive control and was set to 100%. Data are presented as mean value (SD) out of quadruplicates. (b) Gene silencing by siRNA lipo-polyplexes in neuroblastoma cells (left) and human prostate cancer cells (right). Lipopolyplexes with 500 ng (37 pmol) eGFP-targeted siRNA (siGFP) per well respectively control siRNA (siCtrl) at N/P 12 and 20 were tested for eGFPLuc gene silencing in Neuro2AeGFPLuc and DU145-eGFPLuc cells. The luciferase activity of siRNA treated cells is presented related to buffer treated cells. HBG-treated cells were set to 100%. Data are presented as mean value (SD) out of triplicates. Adapted from ([51] Reinhard et al., ChemMedChem, 2017) with permission from Wiley 2.1.2 Reagents and Solvents

1. Dichloromethane (DCM). 2. N,N-dimethylformamide (DMF) (Iris Biotech, Marktredwitz, Germany). 3. N,N-diisopropylethylamine (DIPEA) (Iris Biotech, Marktredwitz, Germany). 4. 1-Hydroxybenzotriazole (HOBt) (Sigma-Aldrich, Munich, Germany).

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So¨ren Reinhard and Ernst Wagner

5. Benzotriazol-1-yl-oxy tripyrrolidinophosphonium hexafluorophosphate (Pybop®) (Multisyntech, Witten, Germany). 6. Fmoc-deprotection solution: 20% (v/v) piperidine–DMF. 7. Capping solution: (MeOH)–DIPEA.

80:15:5

(v/v/v)

DCM–methanol

8. Kaiser test solutions: 80% (w/v) phenol in EtOH; 5% (w/v) ninhydrin in EtOH; 20 μM KCN in pyridine (2 mL of 1 mM KCN (aq) in 98 mL of pyridine). 9. Dde-deprotection solution: 2% (v/v) hydrazine monohydrate in DMF. 10. DIPEA washing solution: 10% (v/v) DIPEA in DMF. 11. Cleavage cocktail: 94:2.5:2.5:1 (v/v/v/v) trifluoroacetic acid (TFA)–1,2-ethanedithiol (EDT)–triisopropylsilane (TIS)–dH2O. 12. Incubation solution for generation of OH-SteA-t: 95:5 (v/v) TFA–DCM. 13. Reducing agent: Tris(2-carboxyethyl)phosphine hydrochloride (TCEP) (Sigma-Aldrich, Munich, Germany). 14. Precipitating solution: 1:1 (v/v) methyl tert-butyl ether (MTBE)–n-hexane. 15. Ellman’s reagent stock solution: 4 mg/mL 5,50 -dithiobis(2-nitrobenzoic acid) (DTNB) in MeOH. 16. Cysteine stock solution: 50 mM L-cysteine in Ellman’s buffer. 2.1.3 Oligomer Purification

1. Chromaster HPLC-DAD (VWR Hitachi, Darmstadt, Germany) with Chromaster System Manager (Ver. 1.1 by Hitachi). 2. HPLC column: YMC-Pack C4, 250  10 mm, 5 μm, 12 nm (YMC, Kyoto, Japan). 3. HPLC mobile phases: A: 0.1% TFA in 40:60 MeOH–H2O B: 0.1% TFA in acetonitrile (ACN). 4. Anion exchange solution: 10 mM HCl in 30:70 (v/v) ACN–H2O.

2.1.4 Oligomer Analysis

1. AVANCE III HD 500 (500 MHz) NMR spectrometer with a 5 mm CPPBBO probe. 2. Deuterium oxide (D2O). 3. MestreNova (Ver. 9.0 by MestReLab Research). 4. Autoflex II MALDI mass spectrometer (Bruker Daltonics, Bremen, Germany). 5. MTP AnchorChip (Bruker Daltonics, Bremen, Germany).

Defined Lipo-Oligomers Containing Unsaturated Fatty Acids

7

6. sDHB (sum of 2,5-dihydroxybenzoic acid and 2-hydroxy-5methoxybenzoic acid) in 1/1 acetonitrile/water containing 0.1% (v/v) TFA. 7. GENESYS™ UV-VIS spectrophotometer (Thermo Scientific, Schwerte, Germany). 2.2 Polyplex Formation

1. Ready-to-use siRNA duplexes (Axolabs GmbH, Kulmbach, Germany) eGFP-targeted siRNA (siGFP) (sense: 50 -AuAucAuGGccGAcAAGcAdTsdT-30 ; antisense: 50 -UGCUUGUCGGCcAUGAuAUdTsdT-30 ; small letters: 20 methoxy; s: phosphorothioate) for silencing of eGFPLuc; control siRNA (siCtrl) (sense: 50 -AuGuAuuGGccuGuAuuAGdTsdT-30 ; antisense: 50 -uAAuAcAGGCcAAuAcAUdTsdT-30 ).

2.3 Biophysical Polyplex Characterization

1. Agarose NEEO Ultra (Carl Roth GmbH, Karlsruhe, Germany). 2. GelRed (VWR, Darmstadt, Germany). 3. Dark hood DH-40 with UV transilluminator (Biostep, Burkhardtsdorf, Germany). 4. Zetasizer Nano ZS with backscattering detection and folded capillary cells (Malvern Instruments, Worcestershire, UK).

2.4 Cell Culture and Biological Assays

1. Citrate-buffered human blood. 2. Triton X-100 2% (v/v) in PBS, pH 7.4, 6.5, 5.5. 3. Cell culture media and antibiotics (Invitrogen, Karlsruhe, Germany). 4. Trypsin (0.25%) and ethylenediamine tetraacetic acid (EDTA) (1 mM) (Invitrogen, Karlsruhe, Germany). 5. 75 cm2 cell culture flasks (TPP, Trasadingen, Switzerland). 6. Fetal calf serum (FCS) (Invitrogen, Karlsruhe, Germany). 7. Cell culture 5  lysis buffer (Promega, Mannheim, Germany). 8. D-luciferin sodium salt (Promega, Mannheim, Germany). 9. 10 mM luciferin solution: 48 mg of D-luciferin sodium salt in 470 μL of 1 M glycylglycine (Sigma-Aldrich, Munich, Germany). Fill up to 16 mL with dH2O, adjust to pH 8. 10. Luciferase assay reagent (LAR): 50.8 mg of DTT (SigmaAldrich, Munich, Germany) and 27.8 mg ATP (Sigma-Aldrich, Munich, Germany) in 2 mL of 1 M glycylglycine, 1 mL of 100 mM MgCl2, 20 μL of 500 mM EDTA and 0.5 mL of 42.6 mg/mL coenzyme A (Sigma-Aldrich, Munich, Germany). Fill up to 100 mL with dH2O, adjust to pH 8. 11. CellTiter-Glo® Kit (Promega, Mannheim, Germany).

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So¨ren Reinhard and Ernst Wagner

12. Microplate 96 well, V-bottom (Ratiolab, Dreieich, Germany). 13. Microplate 96 well (TPP, Trasadingen, Switzerland). 14. Duomax 1030 Germany).

orbital

shaker

(Heidolph,

Schwabach,

15. Spectrafluor Plus plate reader (Tecan, Gro¨dig, Austria). 16. Centro LB 960 plate reader luminometer (Berthold Technologies, Bad Wildbad, Germany). 2.5

Buffers

1. Ellman’s buffer: 0.2 M Na2HPO4, 1 mM EDTA in dH2O, pH 8.0. 2. TBE buffer: 89 mM TRIS, 89 mM boric acid, 2 mM ethylenediaminetetraacetic acid disodium salt (EDTA-Na2) in dH2O, pH 8.0. 3. HEPES buffer: 20 mM HEPES in dH2O, pH 7.4. 4. HBG buffer: 5% (w/v) glucose in 20 mM HEPES, pH 7.4. 5. Electrophoresis 6  loading buffer: 6 mL of glycerine, 1.2 mL of 0.5 M EDTA-Na2 solution, pH 8.0, 2.8 mL of dH2O, 20 mg of bromophenol blue. 6. PBS buffer: 0.2 g/L KCl, 0.2 g/L KH2PO4, 8 g/L NaCl, 1.15 g/L Ka2HPO4 in H2O, pH 7.4, 6.5 and 5.5. 7. Sodium citrate solution: 250 mM trisodium citrate dehydrate in dH2O. 8. Citrate-buffered PBS: 1/9 (v/v) sodium citrate solution in PBS buffer, pH 7.4.

3

Methods

3.1 Solid-Phase Assisted Synthesis (SPS)

SPS is an elegant approach to synthesize precisely defined oligomer structures. Defined structures facilitate the establishment of clear structure–activity relationships and consequently the rational design and optimization of nucleic acid carriers. Oligomers OleAt, OH-SteA-t, SteA-t, and LinA-t are synthesized by standard Fmoc-SPS following repetitive synthesis cycles (Fig. 4): 1. Coupling (90 min). 2. Washing (3 DMF, 3 DCM). 3. Kaiser Test. 4. Fmoc Deprotection (4  10 min with 20% (v/v) piperidine–DMF). 5. Washing (3 DMF, 3 DCM). 6. Kaiser Test.

Defined Lipo-Oligomers Containing Unsaturated Fatty Acids

9

Fig. 4 Standard procedure of a solid phase synthesis cycle. Reproduced and modified from ([63] Morys et al., Methods in Molecular Biology, 2016) with permission from Springer

A 2-chlorotrityl chloride resin is used as solid support. Fmoc protected amino acids and fatty acids are coupled with a fourfold excess in relation to the free amines using HOBt (fourfold excess) and PyBOP (fourfold excess) for activation. DIPEA is added as an auxiliary base with an eightfold excess in relation to the free amines. All coupling reagents are dissolved in 1/1 (v/v) DCM/DMF (10 mL/g resin) and mixed for preactivation before adding them to the resin. Couplings are carried out for 90 min at room temperature (RT) using an overhead shaker (see Note 8). Fmoc deprotection is accomplished by incubating the resin 4 times for 10 min with 20% (v/v) piperidine–DMF (10 mL/g resin). After each coupling and Fmoc deprotection step, the resin is washed three times with DMF and three times with DCM (10 mL/ g of resin). Sufficient coupling and Fmoc deprotection are verified by qualitative detection of free amines by Kaiser test [64]. In case of a positive result of the Kaiser test after coupling, the last coupling step is repeated. In case of a negative result after deprotection, the last deprotection step is redone. 3.1.1 Resin Loading

1. Place 0.75 g of 2-chlorotrityl chloride resin (1.2 mmol chloride) in a 10 mL syringe reactor. 2. Swell the resin by shaking for approximately 20 min in 5 mL of water-free DCM (see Note 9). 3. Discard the DCM. 4. Dissolve 0.75 eq of the first Fmoc-amino acid, Fmoc-Cys(Trt)OH in case of OleA-t, OH-SteA-t, SteA-t, and LinA-t, and 1.5 eq of DIPEA in 5 mL of dry DCM. 5. Add the solution to the resin and shake for 60 min. 6. Discard the solution and add 5 mL of the capping solution for at least 30 min to block remaining active chloride groups on the resin. 7. Discard the solution and wash three times with DMF and three times with DCM (5 mL each).

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So¨ren Reinhard and Ernst Wagner

8. Dry the resin under high vacuum and weight triplicate samples (between 5–10 mg each) into polystyrene microcentrifuge tubes. 9. Add 1 mL of Fmoc-deprotection solution to the samples and incubate for 90 min at RT in a vortexing unit (see Note 10). 10. Take 25 μL of the supernatant and dilute it by adding 975 μL DMF. 11. Vortex and calculate the resin loading based on the absorbance at λ ¼ 301 nm against a blank deprotection solution consisting of 25 μL Fmoc-deprotection solution and 975 μL DMF (see Notes 11 and 12). 12. Incubate the resin four times for 10 min with 5 mL of Fmocdeprotection solution. 13. Wash the resin three times with DMF and three times with DCM (5 mL each) and perform a Kaiser test. 14. In case of a positive Kaiser test result, dry the resin at high vacuum and store well sealed to exclude air and moisture. In case of a negative Kaiser test result, repeat step 12. 3.1.2 Kaiser Test

1. Transfer a few beads of resin into a polystyrene microcentrifuge tube and add 1 drop of each Kaiser test solution. 2. Vortex and spin down quickly. 3. Incubate at 100
C for 4 min. A positive reaction can occur within the first seconds of the incubation time. Free amines are indicated by blue color.

3.1.3 Synthesis of OleA-t, LinA-t, and SteA-t

1. Here the synthesis of t-shaped lipo-oligomers OleA-t, LinA-t, and SteA-t is described. OH-SteA-t is synthesized from OleAt by complete conversion of the unsaturated double bonds into TFA-esters and subsequent hydrolysis (see Note 13). Considering resin loading and desired synthesis scale size, take the required amount of preloaded L-Cys(Trt)-OH resin and transfer into the corresponding reactor. Here, a resin amount equivalent to 15 μmol is used for each synthesis. 2. Swell the resin for 30 min with 10 mL/g of DCM. 3. Synthesize the linear backbone of the oligomer (CY3Stp2K (Dde)Stp2Y3C) by sequential coupling and deprotection of three Fmoc-L-Tyr(tBu)-OH amino acids. Continue with two Fmoc-Stp(Boc3)-OH, Fmoc-L-Lys(Dde)-OH, two Fmoc-Stp (Boc3)-OH, three Fmoc-L-Tyr(tBu)-OH, and end with a Boc-L-Cys(Trt)-OH. 4. For Dde deprotection, freshly prepare the Dde-deprotection solution with 2% (v/v) hydrazine hydroxide solution in DMF and apply 5 mL/g of resin and vortex for 2 min. Repeat

Defined Lipo-Oligomers Containing Unsaturated Fatty Acids

11

30 deprotection cycles and exchange the Dde-deprotection solution after each cycle (see Note 14). 5. Wash the resin five times with DIPEA washing solution, three times with DMF and three times with DCM (10 mL/g of resin). 6. Continue with the synthesis by coupling a branching Fmoc-LLys(Fmoc)-OH using the standard coupling conditions. From this step on, the number of free amines is doubled (see Note 15). 7. After deprotection and Kaiser Test, attach the hydrophobic moieties (oleic acid in case of OleA-t, stearic acid in case of SteA-t and linoleic acid in case of LinA-t). No Fmocdeprotection is required after coupling and washing. 8. Dry the resins on high vacuum for approximately 30 min. 9. Prepare the cleavage cocktail and cool the solution to 4
C (see Note 16). 10. Precool the resin to 4
C and apply 10 mL/g of cleavage solution for 20 min at RT while shaking. 11. Immediately precipitate the oligomers by adding the cleavage solution dropwise into 50 mL of ice cold precipitating solution. 12. Centrifuge for 10 min (1789  g, 4
C), discard the supernatant and dry the precipitate under nitrogen (N2) stream. 13. Dissolve the obtained product in HPLC mobile phase A (0.1% TFA in 40:60 MeOH–H2O) and purify it by HPLC (see Note 17). 14. Pool the product containing fractions into a tared 15 mL tube, freeze with liquid N2 and lyophilize. 15. Perform an anion exchange to exchange TFA anions with Cl by three cycles of redissolving in anion exchange solution and subsequent lyophilizing. 16. Determine the product yield by balancing and analyze the samples by 1H-NMR and MALDI mass spectrometry (see Notes 18 and 19). 17. Store the obtained HCl salts of the oligomers (see Note 20) dry or as stock solutions tightly sealed under argon at 20
C (see Notes 21 and 22). 3.1.4 Synthesis of OH-SteA-t

1. Redissolve the precipitated OleA-t (after step 12 of Subheading 3.1.3) in 95/5 (v/v) TFA/DCM (10 mL/g) and incubate for 12 h under gentle shaking at room temperature to generate TFA-SteA-t. 2. Precipitate TFA-SteA-t by adding the solution dropwise into 50 mL of ice cold precipitating solution.

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3. Redissolve TFA-SteA-t in 20 mM HEPES, adjust the pH to 7.4 with 1 M NaOH and incubate for 12 h under gentle shaking at room temperature to generate OH-SteA-t by TFA ester hydrolysis. 4. Add a tenfold molar excess of reducing agent TCEP and stir for 30 min at room temperature (see Note 23). 5. Purify the product by HPLC and follow steps 14–17 of Subheading 3.1.3. 3.2

Ellman’s Assay

Ellman’s assay is a colorimetric method for determining low concentrations of mercaptans [65]. The availability of free thiol groups in cysteine-containing lipo-oligomers is important to ensure efficient disulfide cross-linking during polyplex formation. Premature oxidation or blocking by reactive intermediates during oligomer cleavage can cause a loss of free sulfhydryl groups. Therefore, sufficient amounts of scavengers (e.g., EDT) must be available in the cleavage cocktail and the purified oligomers should be stored well sealed to exclude air and moisture. 1. Prepare an Ellman’s reagent (DTNB) working solution by diluting Ellman’s reagent stock solution in Ellman’s buffer by 1:40 (50 μL of stock solution in 1950 μL Ellman’s buffer). 2. Prepare a 0.5 mM cysteine working solution by diluting cysteine stock solution in Ellman’s buffer by 1:100 (10 μL of stock solution in 990 μL Ellman’s buffer). 3. Prepare cysteine calibration solutions by diluting the cysteine working solution in Ellman’s buffer according to Table 1. 4. Calculate the maximum amount of free thiols based on molecular weight of the oligomers and prepare sample solutions containing 0.2–0.4 mM thiols in Ellman’s buffer. 5. Add 30 μL of the sample solutions and the cysteine calibration solutions to 170 μL of Ellman’s reagent working solution. 6. Incubate the samples for 15 min at 37
C while shaking. 7. Measure the absorption at λ ¼ 412 nm and prepare a calibration curve from the cysteine calibration solutions. Blank the UV-VIS spectrophotometer with Ellman’s buffer. 8. Calculate the concentration of thiols in the sample by using the calibration equation as a percentage of the calculated maximum amount (see Note 24).

3.3 Erythrocyte Leakage Assay

An erythrocyte leakage assay can be performed to assess the membranolytic potential of lipo-oligomers. The assay is based on the release of hemoglobin from erythrocytes during incubation with lipo-oligomers at different concentrations and pH values. Lysis of membranes can, on the one hand, cause toxic effects on cells, but on the other hand, improve endosomal escape of the formulations

Defined Lipo-Oligomers Containing Unsaturated Fatty Acids

13

Table 1 Preparation of a calibration curve for determination of free thiols by Ellman’s assay Ellman’s buffer [μL]

Cysteine working solution [μL]

c(SH) [μM]

100

0

0

80

20

0.1

60

40

0.2

40

60

0.3

20

80

0.4

0

100

0.5

and thereby increase the efficiency of the nucleic acid delivery system. Therefore, a pH-responsive lytic activity with low lysis at extracellular and cytosolic neutral pH values and increasing lytic activity at slightly acidic endosomal conditions is highly desirable. 1. Wash 0.5–1 mL of citrate-buffered blood with 20 mL citratebuffered PBS. 2. Centrifuge at 4
C, 447  g for 10 min and discard the supernatant. Repeat this step until the supernatant is clear (3–4 times). 3. Add 5 mL of PBS 7.4, count the cells and prepare three dilutions with PBS buffer at pH 7.4, 6.5 and 5.5 with 5  107 erythrocytes/mL each (see Note 25). 4. Prepare a 100 μM stock solution of the oligomer and dilute it with PBS buffer at pH 7.4, 6.5 and 5.5 to the double concentration of your intended test concentration. 5. Pipet quadruplicates with 75 μL erythrocyte suspension and 75 μL test substance in different concentrations at each pH value in a V-bottom 96 well plate. For each plate, prepare quadruplicates with 75 μL erythrocyte suspension and 75 μL PBS buffer as a negative control and 2% Triton X-100 solution as a positive control for each pH value. 6. Incubate the plates for 60 min at 37
C under constant shaking. 7. Centrifuge the plates at 4
C, 447  g for 10 min. 8. Transfer 100 μL of the supernatant in a 96 well plate and measure absorption at λ ¼ 405 nm (see Note 26). 9. Subtract the values of the PBS-treated negative control and calculate the lysis as a percentage of the Triton X-100-treated positive control. 3.4 Polyplex Formation

Lipo-polyplexes form by electrostatic and hydrophobic interactions upon mixing of positively charged lipo-oligomers with negatively charged nucleic acids. Nucleic acid binding and stability of the

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So¨ren Reinhard and Ernst Wagner

resulting lipo-polyplexes is highly dependent on the ratio between oligomers and nucleic acids. The N/P value is frequently used to describe the ratio between the oligomer and nucleic acid in the formulation (see Note 27). 1. Dilute 0.5 μg of siRNA in 10 μL of HBG for each desired N/P. 2. Dilute the calculated amount of oligomer in 10 μL of HBG for each desired N/P (see Note 28). 3. Add the oligomer solution to the siRNA solution and mix by rapid pipetting 10 times. 4. Incubate the solution for 40 min at room temperature to allow disulfide bond formation by air oxidation. Experiments should be carried out immediately after the incubation time is completed. 3.5 Biophysical Polyplex Characterization

3.5.1 Agarose Gel Shift Assay for siRNA Polyplexes

Biophysical properties of polyplexes are important regarding their stability and possible interactions with biological constituents, fluids and membranes, thereby influencing their efficiency and safety. Nucleic acid binding and polyplex stability can be screened by agarose gel retardation assays with or without serum. Negatively charged siRNA migrates in an electric field unless the nucleic acid is complexed by the cationic lipo-oligomers. Upon complexation, the electrophoretic mobility is reduced or even reversed due to loss of negative charge and increased size. The agarose gel shift assay is used to evaluate the ability of the lipo-oligomers to bind siRNA as well as to determine the minimum required N/P ratio for complete nucleic acid binding. Polyplexes can be exposed to 90% full serum at 37
C for several hours before electrophoresis, which can be an indicative assay for extracellular stability. Polyplex size, polydispersity, and ζ-potential are important biophysical properties and can be assessed by dynamic light scattering (DLS). 1. Fix a 15 cm  15 cm UV-transparent gel tray in a gel casting unit. 2. Weigh 4.5 g of agarose powder (2.5% gel, w/v) into a flask and add 180 mL of TBE buffer. 3. Dissolve agarose by boiling until a clear solution is obtained. 4. Allow the solution to cool down to ~50
C, then add 180 μL of 1000  concentrated GelRed™ for nucleic acid staining. 5. Pour the agarose solution into the gel casting unit. Eliminate bubbles with a pipette tip before fixating a well comb. Allow the gel to solidify (usually within 30 min). 6. Prepare polyplexes containing 0.5 μg of siRNA in 20 μL HBG at different N/Ps as described in Subheading 3.4. Optionally, add full serum to a final concentration of 90% after polyplex incubation is finished and incubate at 37
C for the desired time before performing the gel shift assay.

Defined Lipo-Oligomers Containing Unsaturated Fatty Acids

15

7. Prepare all relevant controls for the assay. If serum incubation is performed, use free siRNA (0.5 μg of siRNA in 20 μL HBG), 9/1 (v/v) full serum/HBG (20 μL) and free siRNA (0.5 μg of siRNA in 9/1 full serum/HBG, 20 μL) as a control. If the polyplexes are incubated without serum, use free siRNA (0.5 μg of siRNA in 20 μL HBG) as a control. 8. After polyplex incubation is completed, add 4 μL of 6  loading buffer to all samples and controls. 9. Remove the well comb from the gel and transfer the gel to the electrophoresis unit. 10. Fill the chamber with TBE buffer until the gel is fully covered and all pockets are filled. 11. Load the samples into the sample pockets and apply a voltage of 100 V for 40 min. 12. Observe the gel under UV light exposure. 3.5.2 Size Measurement via DLS and Measurement of ζ-Potential

1. Prepare oligomer solution of the desired N/P in 40 μL of HBG. 2. Prepare a solution of 2 μg siRNA in 40 μL of HBG. 3. Prepare polyplexes as described in Subheading 3.4. 4. After 40 min of incubation, transfer the polyplex solution into a folded capillary cell. 5. For size and polydispersity measurements, set the method of the zetasizer to three measurements with 15 subruns (see Note 29). 6. For the determination of the ζ-potential, add 720 μL HEPES 20 mM and measure each sample three times with 10–15 subruns at 25
C.

3.6 Cell Culture and Treatment

Different cell lines are used and cultured in the appropriate media (Table 2). Cells stably transfected with the eGFPLuc gene (Neuro2A/eGFPLuc cells and DU-145/eGFPLuc cells [66]) are used for siRNA silencing assays. For cell viability assays, also wild-type cells (Neuro2A cells and DU-145 cells) can be used. The cells are cultured in ventilated flasks inside incubators at 37
C with 5% CO2 in a humidified atmosphere. Cell lines are grown to 80–90% confluency and split when necessary. All experiments are performed in triplicates. 1. Seed 5  103 cells per well in 100 μL of media using 96-well plates 24 h before the experiment. 2. Replace media with 80 μL of fresh media before the experiment. 3. Prepare polyplexes as described in Subheading 3.4 and add 20 μL polyplex solution per well.

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So¨ren Reinhard and Ernst Wagner

Table 2 Cell lines and culture media Cell line

Cell culture media

Mouse neuroblastoma Neuro2A cells Neuro2A/eGFPLuc cells

Dulbecco’s modified Eagle’s medium (DMEM), 10% fetal bovine serum, 100 U/mL penicillin, and 100 μg/mL streptomycin

Human prostate carcinoma DU-145 cells DU-145/eGFPLuc cells

RPMI-1640 medium, 10% fetal bovine serum, 100 U/ mL penicillin, and 100 μg/mL streptomycin

4. Always prepare untreated cells as a control on each plate. 5. Gently shake the 96-well plates horizontally for few seconds for complete diffusion of polyplexes. 6. The cells are incubated at 37
C for 48 h. 3.6.1 siRNA Silencing Assay

Gene silencing experiments can be performed in reporter cell lines stably expressing an eGFP-Luciferase fusion protein. Thus, silencing by siGFP can be quantified by a standard luciferase assay (resulting in luciferase activity decrease). Treatment with analogous control siCrtl should maintain luciferase activity close to 100% unless unspecific effects take place. 1. Cells are harvested and analyzed for gene silencing after 48 h incubations with siGFP- and siCrtl-containing polyplexes. 2. Prepare 1  cell lysis buffer by diluting an aliquot of 5  lysis buffer with four volumes of dH2O. 3. Remove the transfection media. 4. Add 100 μL of 1  cell lysis buffer into each well. 5. Mix 500 μL of 10 mM luciferin with 10 mL luciferase assay reagent (LAR) and apply the injector of a plate reader luminometer into the mixture (see Note 30). 6. Program the injector to add 100 μL of luciferin-LAR solution per well. 7. Transfer 35 μL of cell lysate into a white 96-well microplate. 8. Measure the relative light units (RLU) for 10 s with a plate reader luminometer. 9. Calculate the RLU as percentage of the luciferase gene expression obtained with untreated control cells.

3.6.2 Cell Viability Assay (CellTiter-Glo® Assay)

Unspecific cytotoxicity of nucleic acid carriers may cause safety concerns and limit their applicability. Measuring metabolic cell activities compared to untreated cells via a CellTiter-Glo® assay based on the quantitation of ATP can serve as an indicator for cell viability.

Defined Lipo-Oligomers Containing Unsaturated Fatty Acids

17

1. Cells are harvested and analyzed for metabolic activity after 48 h incubations with siGFP- and siCrtl-containing polyplexes. 2. Add 100 μL of CellTiter-Glo® reagent to the media in each well. 3. Place the plate on an orbital shaker for 10 min to induce cell lysis. 4. Transfer the lysate into a white 96-well microplate. 5. Measure the luminescence with a plate reader luminometer. 6. Calculate the metabolic activity as percentage of the untreated control cells.

4

Notes 1. The ability of the lipo-oligomers to bind siRNA is determined by measuring the electrophoretic mobility of siRNA in a 2.5% agarose gel as described in Subheading 3.5.1. The same assay is repeated after incubation with 90% serum at 37
C as an indicative assay for extracellular stability. All formulations showed similar siRNA binding and polyplex stability in serum, rendering the T-shape oligomers well-suited for the evaluation of structure–activity relationships. 2. Modifications of the hydrophobic moieties are expected to alter the interaction of the lipo-oligomers with the membrane lipids, resulting in different membranolytic potentials. An erythrocyte leakage assay (as described in Subheading 3.3) is performed, comparing the lysis of erythrocyte membranes at different concentrations and pH values (Fig. 3a). All three lipooligomers showed a desirable increase in lytic activity at slightly acidic endosomal pH values in accordance with the progressive protonation of the Stp-units, which increases the cationic character for enhanced lipid membrane binding. The structures with unsaturated or modified hydrocarbon chains displayed significantly higher lytic activity than the saturated SteA-t oligomer. OleA-t showed particularly favorable pH dependency of membranolytic activity with low lysis at pH 7.4, while OH-SteA-t showed significantly higher erythrocyte lysis at pH 7.4 with threefold increase compared to OleA-t at 5 mM concentration. This enhanced lytic activity at neutral pH may result in increased cellular internalization on the one hand, but in undesired cytotoxicity on the other hand, as cell membrane lysis can occur already at the cell surface or at intracellular membranes after release of lipo-oligomers into the cytosol. 3. The particle sizes of the lipo-polyplexes are measured by dynamic light scattering (DLS) as described in Subheading 3.5.2. All formulations showed uniform sizes below 150 nm

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So¨ren Reinhard and Ernst Wagner

z-average with positive zeta potentials, ranging from +39 to +46 mV, due to the cationic Stp units of the oligomers. 4. Gene silencing assays (as described in Subheading 3.6.1) are performed in Neuro2A neuroblastoma cells and DU145 human prostate cancer cells stably expressing an eGFPLuciferase fusion protein (Fig. 3b). Especially in DU145 cells, lipo-polyplexes formulated with control siRNA and OH-SteAt showed reduced luciferase activity, indicating unspecific toxicity of the formulations, while OleA-t formulations displayed high cell tolerability and high gene silencing efficacy. SteAt polyplexes showed low gene silencing activity in both cell lines, which is in accordance with the low lytic activity of the lipo-oligomer. 5. High cellular compatibility of OleA-t and SteA-t polyplexes are confirmed by CellTiter-Glo® assays (as described in Subheading 3.6.2) as an indicator of cell viability. Obviously, hydroxylation of the unsaturated fatty acids by TFA-mediated cleavage after SPS, caused by TFA addition and subsequent hydrolysis of the TFA esters, have the potential to enhance the lytic activity at physiological pH and therefore can increase the cytotoxicity of lipo-oligomers. Saturated fatty acids seem to be a poor alternative to unsaturated FAs, as they showed low gene silencing activity such as in case of SteA-t. 6. DMF tends to hydrolyze to formic acid and dimethylamine when exposed to water. This can reduce coupling efficiencies during synthesis and cause false positive results in Kaiser tests. Peptide grade quality DMF should be used and stored with caution. 7. Syringe micro reactors are available in different sizes (2–100 mL) and should be chosen according to the resin amount. The amount of resin depends on the determined resin loading (L) and the scale size (scale size[mmol]/L [mmol/g] ¼ resin amount[g]). For the synthesis of OleA-t, OH-SteA-t, SteA-t, and LinA-t, 15 μmol of resin is used which correlated to 27-42 mg of yield. 8. Any stirring mechanism guaranteeing continuous mixing of the solution can be used, such as overhead shakers, vortex mixers, and vortexing units of semi-automated and automated peptide synthesizers. Coupling time can be reduced if microwaveassisted peptide synthesis is performed. The coupling temperature should not exceed 50
C if 2-chlorotrityl chloride resin is used as solid support. 9. Dry DCM by storage over calcium chloride (CaCl2) to prevent H2O from reacting with the resin’s linker. 10. Make sure that the resin is not sedimenting during the incubation by vortexing with high speed (at least 112  g).

Defined Lipo-Oligomers Containing Unsaturated Fatty Acids

19

11. Calculate resin load by the following formula:   mmol 1000  A 301 ¼ Loading g D  7800  m ½mg with D as dilution factor (0.025) and 7800 as molar extinction coefficient [L  mol1  cm1] of Fmoc. Calculate the arithmetic mean of the triplicate values as final resin loading. 12. The determined loading should be approximately 0.3 mmol/g of resin. 13. Sequence (N ! C): OleA-t: H2N-C-Y3-Stp2-K-ε[K-α,ε(OleA)2]αStp2-Y3-C-OH. OH-SteA-t: H2N-C-Y3-Stp2-K-ε[K-α,ε(OH-SteA)2]αStp2-Y3C-OH. SteA-t: H2N-C-Y3-Stp2-K-ε[K-α,ε(SteA)2]αStp2-Y3-C-OH. LinA-t: H2N-C-Y3-Stp2-K-ε[K-α,ε(LinA)2]αStp2-Y3-C-OH 14. The Dde-deprotection is performed with an automated peptide synthesizer. If the Dde-deprotection is performed manually for example using an overhead shaker, take samples of the Dde-deprotection solution and measure the absorbance against a blank of fresh hydrazine deprotection solution at λ ¼ 290 nm. Deprotection is finished when the absorbance is well below 0.1 for at least three cycles. 15. After branching with Fmoc-L-Lys(Fmoc)-OH, the molar amounts of coupling reagents (fatty acids, PyBOP, HOBt, and DIPEA) have to be doubled since the number amines is doubled. 16. Precooling of both cleavage solution and resin proved to be essential to reduce the amount of TFA-adducts on the unsaturated fatty acids. TFA is a very aggressive acid, so wear protective clothes and protective goggles. If the structures contain no cysteine, EDT is not required in the cleavage cocktail. The solution should then consist of 95/2.5/2.5 (v/v/v) trifluoroacetic acid (TFA)/triisopropylsilane (TIS)/dH2O. 17. For purification of lipo-oligomers, samples are dissolved in 60/40 (v/v) H2O/MeOH containing 0.1% TFA. Column: YMC-Pack C4, 250  10 mm, 5 μm, 12 nm. Mobile phases: A: 0.1% TFA in 60/40 (v/v) H2O/MeOH; B: 0.1% TFA in ACN; 0% B for 10 min, 0–90% B in 40 min, 90% B for 10 min; 2.00 mL/min, 35
C, 280 nm. Chromatograms are recorded using a Chromaster HPLC-DAD system by VWR Hitachi and analyzed using Chromaster System Manager (Ver. 1.1 by Hitachi). 18. 1H-NMR spectra are recorded using an AVANCE III HD 500 (500 MHz) by Bruker with a 5 mm CPPBBO probe. All spectra are recorded without TMS as internal standard and

20

So¨ren Reinhard and Ernst Wagner

therefore all signals are calibrated to the residual proton signal of the deuterium oxide (D2O) solvent. Chemical shifts are reported in ppm and refer to the solvent as internal standard (D2O at 4.79). Integration is performed manually. The spectra are analyzed using MestreNova (Ver. 9.0 by MestReLab Research). 1

H-NMR spectrum of OleA-t (500 MHz, deuterium oxide) δ (ppm) ¼ 0.65–0.85 (s, 6 H, –CH3 oleic acid), 0.85–1.95 (m, 64 H, βγδH lysine, –CH2– oleic acid), 2.0–2.65 (m, 20 H, –CO–CH2–CH2–CO– Stp, –CO–CH2– oleic acid), 2.65–3.1 (m, 20 H, εH lysine, tyrosine, cysteine), 3.1–3.6 (m, 64 H, –CH2– Tp), 3.85–4.55 (m, 10 H, αH amino acids), 5.05–5.20 (s, 4 H, –CH¼CH oleic acid), 6.55–7.10 (m, 24 H, –CH– tyrosine). 1 H-NMR spectrum of OH-SteA-t (500 MHz, deuterium oxide) δ (ppm) ¼ 0.60–0.80 (s, 6 H, –CH3 hydroxystearic acid), 0.80–1.35 (m, 68 H, βγδH lysine, –CH2– hydroxystearic acid), 2.3–2.65 (m, 22 H, –CO–CH2–CH2–CO– Stp, –CO–CH2– hydroxystearic acid, ¼CH–OH hydroxystearic acid), 2.65–3.1 (m, 20 H, εH lysine, tyrosine, cysteine), 3.1–3.6 (m, 64 H, –CH2– Tp), 3.90–4.55 (m, 10 H, αH amino acids), 6.55–7.10 (m, 24 H, –CH– tyrosine). 1 H-NMR spectrum of SteA-t (500 MHz, deuterium oxide) δ (ppm) ¼ 0.60–0.80 (s, 6 H, –CH3 stearic acid), 0.80–1.35 (m, 72 H, βγδH lysine, –CH2– stearic acid), 2.3–2.65 (m, 20 H, –CO–CH2–CH2–CO– Stp, –CO–CH2– stearic acid), 2.65-3.1 (m, 20 H, εH lysine, tyrosine, cysteine), 3.1–3.6 (m, 64 H, –CH2– Tp), 3.90–4.55 (m, 10 H, αH amino acids), 6.55–7.10 (m, 24 H, –CH– tyrosine). 1 H-NMR spectrum of LinA-t (500 MHz, deuterium oxide) δ (ppm) ¼ 0.60–0.80 (s, 6 H, –CH3 linoleic acid), 0.90–2.00 (m, 56 H, βγδH lysine, –CH2– linoleic acid), 2.30–2.60 (m, 20 H, –CO–CH2–CH2–CO– Stp, –CO–CH2– oleic acid), 2.65–2.95 (m, 20 H, εH lysine, tyrosine, cysteine), 2.95–3.55 (m, 64 H, –CH2– Tp), 3.95–4.60 (m, 10 H, αH amino acids), 5.05–5.25 (s, 8 H, –CH¼CH linoleic acid), 6.60–7.10 (m, 24 H, –CH– tyrosine). 19. One micoliter matrix droplet consisting of a saturated solution of sDHB (sum of 2,5-dihydroxybenzoic acid and 2-hydroxy-5methoxybenzoic acid) in 1:1 acetonitrile–water containing 0.1% (v/v) TFA is spotted on a MTP AnchorChip (Bruker Daltonics, Bremen, Germany). After the Super-DHB matrix crystallized, one μL of the sample solution (1 mg/mL in water) is added to the matrix spot. Samples are analyzed in negative ion mode using an Autoflex II mass spectrometer (Bruker Daltonics, Bremen, Germany). [M-H](OleA-t): calc: 3070.9 g/mol, found: 3069.9 g/mol; [M–H](OH-SteA-t):

Defined Lipo-Oligomers Containing Unsaturated Fatty Acids

21

calc: 3106.9 g/mol, found: 3106.2 g/mol; [M–H](SteA-t): calc: 3074.9 g/mol, found: 3074.0 g/mol; [M–H](LinA-t): calc: 3066.9 g/mol, found: 3066.7 g/mol. 20. For calculation of the molecular weight of the HCl salts, each Cl counterion of the protonatable amines of the oligomers has to be considered by adding 36.5 g/mol to the molecular weight. 21. The generated salts tend to be hygroscopic and both cysteines and unsaturated fatty acids are prone to oxidation. Therefore flush the vials with argon and seal well to preserve the products from air and moisture. 22. Stock solutions should be prepared in H2O (concentration: 10 mg/mL) and frozen in aliquots. Dissolving HCl salts in unbuffered H2O results in low pH values of the stock solutions which hampers premature oxidation of the cysteines. 23. The reducing agent TCEP is used to reverse a possible premature oxidation of thiols during the TFA ester hydrolysis at neutral pH. 24. Determination of free thiols as percentage of calculated thiols in lipo-oligomers via Ellman’s assay based on weighted samples: OleA-t: 87%; OH-SteA-t: 84%; SteA-t: 80%. Deviation from 100% might be due to residual water or salt and premature oxidation of thiols. 25. Prepare 4  75 μL of erythrocyte suspension at each pH for quadruplicate measurements and prepare the same amount for the negative and positive control of each plate. 26. Make sure not to damage the erythrocyte pellet while transferring the supernatant and pipet the Triton X-100-treated control carefully as it tends to form bubbles. 27. The N/P values depict the ratio of protonatable amines (N) of the oligomer to phosphates (P) of the siRNA. As the diaminoethylene motif of Stp is pH-responsive, not all protonatable amines are protonated at neutral pH, which is why the N/P ratio does not present charge ratios during polyplex formation. 28. nðphosphateÞ ¼ mðsiRNA Þ=av:mol weight of nucleotide nðnitrogenÞ ¼ nðphosphateÞ  desired N =P nðnitrogenÞ vðSampleÞ ¼ cðoligomerÞ  number of protonatable amines Average MW of nucleotides ¼ 311 g/mol. The number of protonatable amines is calculated by all available secondary (Stp) amines as well as N-terminal primary amines. The unit of c(oligomer) is mol/L.

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So¨ren Reinhard and Ernst Wagner

29. The equilibration time is set to 0 min, the temperature is 25
C and an automatic attenuator is used. The refractive index of the solvent is 1.330 and the viscosity is 0.8872 mPals. 30. Light intensity is determined by the rate of catalysis by luciferase and is dependent on the temperature. Therefore, it is important to equilibrate the luciferase assay reagents to room temperature before measurements. The solution is stable for approximately 2 weeks at 4
C.

Acknowledgments This work was supported by DFG SFB1032 B4 (to E.W.), SFB1066 B5 (E.W.), DFG FOR1406 (E.W.) and DFG Excellence Cluster Nanosystems Initiative Munich (E.W.). References 1. Friedmann T, Roblin R (1972) Gene therapy for human genetic disease? Science 175 (25):949–955 2. Mulligan RC (1993) The basic science of gene therapy. Science 260(5110):926–932 3. Fire A (1999) RNA-triggered gene silencing. Trends Genet 15(9):358–363 4. Elbashir SM, Harborth J, Lendeckel W, Yalcin A, Weber K, Tuschl T (2001) Duplexes of 21-nucleotide RNAs mediate RNA interference in cultured mammalian cells. Nature 411 (6836):494–498 5. Hannon GJ (2002) RNA interference. Nature 418(6894):244–251 6. Wagner E (2012) Functional polymer conjugates for medicinal nucleic acid delivery polymers in nanomedicine. In: Kunugi S, Yamaoka T (eds) Advances in polymer science, vol vol 247. Springer, Berlin/Heidelberg, pp 1–29 7. Reinhard S, Wagner E (2017) How to tackle the challenge of siRNA delivery with sequencedefined oligoamino amides. Macromol Biosci 17(1) 8. de Fougerolles A, Vornlocher HP, Maraganore J, Lieberman J (2007) Interfering with disease: a progress report on siRNA-based therapeutics. Nat Rev Drug Discov 6 (6):443–453 9. Wagner E (2007) Programmed drug delivery: nanosystems for tumor targeting. ExpertOpinBiol Ther 7(5):587–593 10. Davis ME, Zuckerman JE, Choi CH, Seligson D, Tolcher A, Alabi CA, Yen Y, Heidel JD, Ribas A (2010) Evidence of RNAi in humans from systemically administered siRNA

via targeted nanoparticles. Nature 464:1067–1070 11. Lee ER, Marshall J, Siegel CS, Jiang C, Yew NS, Nichols MR, Nietupski JB, Ziegler RJ, Lane MB, Wang KX, Wan NC, Scheule RK, Harris DJ, Smith AE, Cheng SH (1996) Detailed analysis of structures and formulations of cationic lipids for efficient gene transfer to the lung. Hum Gene Ther 7(14):1701–1717 12. Ma JB, Ye K, Patel DJ (2004) Structural basis for overhang-specific small interfering RNA recognition by the PAZ domain. Nature 429 (6989):318–322 13. Martin B, Sainlos M, Aissaoui A, Oudrhiri N, Hauchecorne M, Vigneron JP, Lehn JM, Lehn P (2005) The design of cationic lipids for gene delivery. CurrPharmDes 11(3):375–394 14. Akinc A, Zumbuehl A, Goldberg M, Leshchiner ES, Busini V, Hossain N, Bacallado SA, Nguyen DN, Fuller J, Alvarez R, Borodovsky A, Borland T, Constien R, de Fougerolles A, Dorkin JR, Narayanannair JK, Jayaraman M, John M, Koteliansky V, Manoharan M, Nechev L, Qin J, Racie T, Raitcheva D, Rajeev KG, Sah DW, Soutschek J, Toudjarska I, Vornlocher HP, Zimmermann TS, Langer R, Anderson DG (2008) A combinatorial library of lipid-like materials for delivery of RNAi therapeutics. NatBiotechnol 26(5):561–569 15. Semple SC, Akinc A, Chen J, Sandhu AP, Mui BL, Cho CK, Sah DWY, Stebbing D, Crosley EJ, Yaworski E, Hafez IM, Dorkin JR, Qin J, Lam K, Rajeev KG, Wong KF, Jeffs LB, Nechev L, Eisenhardt ML, Jayaraman M, Kazem M, Maier MA, Srinivasulu M, Weinstein

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oligonucleotide transfer into cells in culture and in vivo: polyethylenimine. ProcNatlAcadSciUSA 92(16):7297–7301 37. Behr JP (1997) The proton sponge: a trick to enter cells the viruses did not exploit. Chimia 51(1-2):34–36 38. L€achelt U, Wagner E (2015) Nucleic acid therapeutics using polyplexes: a journey of 50 years (and beyond). Chem Rev 115 (19):11043–11078 39. Schaffert D, Badgujar N, Wagner E (2011) Novel Fmoc-polyamino acids for solid-phase synthesis of defined polyamidoamines. Org Lett 13(7):1586–1589 40. Fro¨hlich T, Edinger D, Kl€ager R, Troiber C, Salcher E, Badgujar N, Martin I, Schaffert D, Cengizeroglu A, Hadwiger P, Vornlocher H-P, Wagner E (2012) Structure–activity relationships of siRNA carriers based on sequencedefined oligo (ethane amino) amides. J Control Release 160(3):532–541 41. Troiber C, Edinger D, Kos P, Schreiner L, Klager R, Herrmann A, Wagner E (2013) Stabilizing effect of tyrosine trimers on pDNA and siRNA polyplexes. Biomaterials 34 (5):1624–1633 42. Klein PM, Muller K, Gutmann C, Kos P, Krhac Levacic A, Edinger D, Hohn M, Leroux JC, Gauthier MA, Wagner E (2015) Twin disulfides as opportunity for improving stability and transfection efficiency of oligoaminoethane polyplexes. J Control Release 205:109–119 43. Wang XL, Nguyen T, Gillespie D, Jensen R, Lu ZR (2008) A multifunctional and reversibly polymerizable carrier for efficient siRNA delivery. Biomaterials 29(1):15–22 44. Steenbergen C, Deleeuw G, Rich T, Williamson JR (1977) Effects of acidosis and ischemia on contractility and intracellular pH of rat heart. CircRes 41(6):849–858 45. Wang XL, Ramusovic S, Nguyen T, Lu ZR (2007) Novel polymerizable surfactants with pH-sensitive amphiphilicity and cell membrane disruption for efficient siRNA delivery. Bioconjug Chem 18(6):2169–2177 46. Lee DJ, Wagner E, Lehto T (2015) Sequencedefined oligoaminoamides for the delivery of siRNAs. Methods Mol Biol 1206:15–27 47. Jiang Q, Yue D, Nie Y, Xu X, He Y, Zhang S, Wagner E, Gu Z (2016) Specially-made lipidbased assemblies for improving transmembrane gene delivery: comparison of basic amino acid residue rich periphery. Mol Pharm 13 (6):1809–1821 48. Lee DJ, He D, Kessel E, Padari K, Kempter S, L€achelt U, Radler JO, Pooga M, Wagner E

(2016) Tumoral gene silencing by receptortargeted combinatorial siRNA polyplexes. J Control Release 244(Pt B):280–291 49. Zhang W, Muller K, Kessel E, Reinhard S, He D, Klein PM, Hohn M, Rodl W, Kempter S, Wagner E (2016) Targeted siRNA delivery using a lipo-oligoaminoamide nanocore with an influenza peptide and transferrin shell. Adv Healthc Mater 5(12):1493–1504 50. Muller K, Kessel E, Klein PM, Hohn M, Wagner E (2016) Post-PEGylation of siRNA Lipo-oligoamino amide polyplexes using tetraglutamylated folic acid as ligand for receptortargeted delivery. Mol Pharm 13 (7):2332–2345 51. Reinhard S, Zhang W, Wagner E (2017) Optimized solid-phase-assisted synthesis of oleic acid containing siRNA nanocarriers. ChemMedChem 12(17):1464–1470 52. Morys S, Urnauer S, Spitzweg C, Wagner E (2018) EGFR targeting and shielding of pDNA lipopolyplexes via bivalent attachment of a sequence-defined PEG agent. Macromol Biosci 18(1) 53. Lee DJ, Kessel E, Lehto T, Liu X, Yoshinaga N, Padari K, Chen YC, Kempter S, Uchida S, Radler JO, Pooga M, Sheu MT, Kataoka K, Wagner E (2017) Systemic delivery of folatePEG siRNA lipopolyplexes with enhanced intracellular stability for in vivo gene silencing in leukemia. Bioconjug Chem 28 (9):2393–2409 54. Peterson PE (1960) Solvents of low nucleophilicity. I. Reactions of hexyl tosylates and hexenes in trifluoroacetic acid and other acids1. J Am Chem Soc 82(22):5834–5837 55. Peterson PE, Allen G (1962) Solvents of low nucleophilicity. II. Addition of trifluoroacetic acid to alkenes and cycloalkenes1. J Org Chem 27(5):1505–1509 56. Peterson PE, Allen G (1963) Solvents of low nucleophilicity. III. The effect of remote substituents in the addition of trifluoroacetic acid to substituted alkenes. J Am Chem Soc 85 (22):3608–3613 57. Peterson PE, Tao EVP (1964) Solvents of low nucleophilicity. IV. Addition of acetic, formic, and trifluoroacetic acid to branched alkenes1. J Org Chem 29(8):2322–2325 58. Peterson PE, Casey C, Tao EVP, Agtarap A, Thompson G (1965) Solvents of low nucleophilicity. VI. The effects of remote substitutents in the addition of trifluoroacetic acid to aliphatic, cyclic, and bicyclic alkenes1a. J Am Chem Soc 87(22):5163–5169 59. Latre`mouille GA, Eastham AM (1967) Kinetics of the addition of acids to olefins with and

Defined Lipo-Oligomers Containing Unsaturated Fatty Acids without boron fluoride catalysis. Can J Chem 45(1):11–16 60. Weisleder D, Friedman M (1968) Addition of halogenated acetic acids to vinyl ketones. Nuclear magnetic resonance study of the kinetics. J Org Chem 33(9):3542–3543 61. Roberts RMG (1976) Kinetics and mechanism of addition of acids to olefins. Part 2. Addition of trifluoroacetic acid to (+)-(R)-limonene in weakly polar media. J Chem Soc Perkin Trans 2 (12):1374–1379 62. Nordlander JE, Haky JE, Landino JP (1980) Mechanism of addition of neat trifluoroacetic acid to protoadamantene. J Am Chem Soc 102 (25):7487–7493 63. Morys S, Wagner E, L€achelt U (2016) From artificial amino acids to sequence-defined

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targeted oligoaminoamides. Methods Mol Biol 1445:235–258 64. Kaiser E, Colescott RL, Bossinger CD, Cook PI (1970) Color test for detection of free terminal amino groups in the solid-phase synthesis of peptides. Anal Biochem 34(2):595–598 65. Ellman GL (1958) A colorimetric method for determining low concentrations of mercaptans. Arch Biochem Biophys 74(2):443–450 66. Fro¨hlich T, Edinger D, Kl€ager R, Troiber C, Salcher E, Badgujar N, Martin I, Schaffert D, Cengizeroglu A, Hadwiger P, Vornlocher HP, Wagner E (2012) Structure-activity relationships of siRNA carriers based on sequencedefined oligo (ethane amino) amides. J Control Release 160(3):532–541

Chapter 2 Synthesis of Bioreducible Polycations with Controlled Topologies Ye-Zi You, Jun-Jie Yan, Fei Yu, Zhi-Qiang Yu, and David Oupicky Abstract Bioreducible polycations, which possess disulfide linkages in the backbone, have emerged as promising nucleic acid delivery carriers due to their high stability in extracellular physiological condition and bioreduction-triggered release of the genetic material. Further benefits of bioreducible polycations include decreased cytotoxicity due to intracellular reducing environment in the cytoplasm that contains high levels of reducing molecules such as glutathione. Here, we describe the synthesis of bioreducible polycations with emphasis on methods to control their topology. Key words Bioreducible polycations, Poly(2-(dimethylamino)ethyl methacrylate, Polyethylenimine, Poly(amido amine)s, Controlled topologies, Michael-addition polymerization

1

Introduction Synthetic polycations have received increasing attention in the delivery of therapeutic nucleic acids [1]. Despite the high stability and low immunogenicity, the successful use of polycationic vectors is limited by several problems, such as low gene transfection efficiency and high cytotoxicity [2–5]. In order to achieve efficient delivery, different types of disulfide-containing polycations have been designed as a way of improving the efficacy of the nucleic acid delivery and reducing cytotoxicity. The polyplexes, prepared by introducing disulfide bonds into the structure of the polycations, show the capability to release the therapeutic nucleic acids selectively in the intracellular reducing space [6, 7]. Such redox-sensitive polyplexes have already proven as effective materials for delivering a variety of nucleic acids, including plasmid DNA, mRNA, antisense oligonucleotides, siRNA, and miRNA [3–5, 8–19]. On the other hand, the basicity and degree of protonation of polycationic vectors depend on their amount of primary, secondary, tertiary amines and their topology, which greatly influence the cytotoxicity, the escape

Manfred Ogris and Haider Sami (eds.), Nanotechnology for Nucleic Acid Delivery: Methods and Protocols, Methods in Molecular Biology, vol. 1943, https://doi.org/10.1007/978-1-4939-9092-4_2, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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of polyplexes from lysosome, and the transfection efficiency. It has been reported that the introduction of irregular branched structure into polycation might improve the transfection, and the topology of polycation has a great impact on the gene transfection efficiency [20, 21]. Therefore, the presence of disulfide bonds and topology of polycations play very important role in gene transfection. Here, we summarize selected proven methods to prepare reducible polycations and control their topologies. The most studied reducible polycations are based on polyethylenimine (PEI), poly(2-(dimethylamino)ethyl methacrylate) (PDMAEMA), poly(amido amine)s, and poly(amino ester)s. Bioreducible PEIs are prepared via functionalizing with reducible crosslinking agents [22, 23], consecutive thiolation and oxidation [14, 24], or Michael-addition reaction [15], their topologies (linear or network) can be controlled by the oligomer topology (linear or branched). Reducible PDMAEMA can be prepared via RAFT polymerization, its topology depends on RAFT agent and crosslinking agent used in the polymerization. RAFT polymerization of DMAEMA using difunctional RAFT agent to obtain α,ω-dithioester-functionalized PDMAEMA, then aminolysis and oxidation of oligomer produce linear disulfide containing PDMAEMA [5], while 1,2-bis(2-(3-methylbuta-1,3-dien-2-yloxy)ethyl) disulfane mediated RAFT polymerization of DMAEMA in the presence of biodegradable disulfide-base dimethacrylate (DSDMA) produces hyperbranched PDMAEMA with each branch point in the polymer linked by a disulfide bond [25]. Bioreducible poly(amido amine)s and poly(amino ester)s are mainly prepared by Michaeladdition polymerization of disulfide-based diacrylamide or diacrylate and amine monomers [9, 10, 26–31]. Their topologies can be finely tuned by the polymerization conditions due to the fact that primary and secondary amines have different reactivity under different polymerization conditions [32].

2

Materials Aqueous solutions are prepared using ultrapure water (prepared by purifying deionized water to attain a resistivity of 18.4 MΩlcm at 25
C) and all reagents are of analytical grade.

3

Methods

3.1 Bioreducible Polyethylenimine (PEI)

Polyethylenimine (PEI) is the most widely used polycation in gene delivery due to its excellent transfection efficiency. However, high cytotoxicity stimulated the demand of synthesis of biodegradable PEI. Bioreducible PEIs are prepared via modifying with reducible cross-linking agents [22, 23], consecutive thiolation and oxidation

Synthesis of Bioreducible Polycations with Controlled Topologies

HO

Ts N

N H

H N

N n Ts 2:n=1 4:n=3

1:n=0 3:n=2

29

OH

(b) Ts N

(a) O

Ts

N Ts

5:n=0 7:n=2

(c)

S O

9:n=0 11 : n = 2 (d)

HS

N H

H N

13 : n = 0 15 : n = 2 (e) S

N H

N Ts

H N

17 : n = 0 19 : n = 2

n

O

Ts

6:n=1 8:n=3

Ts N

N Ts

Ts N

Ts N

N Ts

n

O S

10 : n = 1 12 : n = 3

N H

H N n

SH

14 : n = 1 16 : n = 3

N H

H N n

S m

18 : n = 1 20 : n = 3

Scheme 1 Synthesis of linear reducible PEI (Reproduced from [33] with permission of ACS)

[14, 24], or Michael-addition reaction [15]. Also, reducible PEI polycations can be obtained by covalently linking low molecular weight PEI to biodegradable polymers via disulfide bonds. The topologies (linear, branched, hyperbranched) of bioreducible PEI are mainly controlled by the structure of its oligomer. The detailed procedure for preparing linear reducible PEI is given in Scheme 1 [33]: 1. Synthesis of 5, 6, 7, and 8 (step a). 4-(Dimethylamino)pyridine (10 mmol), triethylamine (92 mmol) and p-toluenesulfonyl chloride (44 mmol) are added into the mixture containing 40 mL of dichloromethane and 10 mmol of 1 (2 or 3 or

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NH2

SH

S N

N H2N

H2N N

N

N N H

NH2 N SH

PEI

PEIX-SHY

H N DMSO

N

NH2

S

N

N

H2N N

S

H2N

N

N

NH

N

N S

S

N NH2

N H

PEIX-SSY

Scheme 2 Preparation of cross-linked linear reducible PEI (Reproduced from [34] with permission of ACS)

4). After the mixture is stirred at 0
C for 24 h, dichloromethane is removed by evaporation and the mixture is redissolved in 100 mL of chloroform. The solution is washed three times with 0.1 M HCl. The organic layer is dried with anhydrous MgSO4. The final products are obtained as white or pale yellow powder. 2. Synthesis of 2, 3, and 4 (step b). Ethanolamine (10 equiv., 10 mmol) is slowly added into mixture containing 20–30 mL of DMF and 1 mmol of 5 (6 or 7). After the reaction mixture is stirred at 50
C for 1–2 days, solvent is removed by evaporation; the mixture is redissolved in DMSO and evaporated again for the removal of the remaining ethanolamine. 3. Synthesis of 9, 10, 11, and 12 (step c). Potassium thioacetate (2.5 equiv) is added into the mixture containing 50 mL of DMF and 10 mmol of 5 (6 or 7 or 8) and stirred at room temperature overnight. DMF is evaporated and the residue is poured into 100 mL of water. The mixture is extracted three or four times with chloroform, and the organic layer is collected, dried with MgSO4. The final products are obtained via evaporating the organic solvent. 4. Tosyl and Acetyl Deprotection (step d). The mixture containing 9 (1 mmol) and phenol (0.41 mmol) in 33% HBr–AcOH (15.4 mL) is refluxed for 24 h. Then, the reaction mixture is filtered, and the resulting pale brown solid is washed with ethanol. After refluxing for 1 h with ethanol, the mixture is

Synthesis of Bioreducible Polycations with Controlled Topologies

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filtered and the resulting solid is washed twice with diethyl ether to obtain the pale brown powder (13). The synthesis of 14, 15, and 16 are carried in the same way. 5. Polymerization (step e). A solution of 13 (or 14 or 15 or 16) (100 mM) in water is stirred at room temperature for 2 days under oxygen pressure (1 atm), then water is removed, the residue is washed with diethyl ether, and dried to give the final polymer products. The outline for preparing cross-linked reducible PEI is shown in Scheme 2 [34], and the detailed procedure is as follows: 1. Synthesis of Thiolated PEI. PEI (1.0 g, 800 Da) is dissolved in 5 mL of deionized water, then hydrochloric acid (0.1 M) is added dropwise to the PEI solution until pH is 7.2. The yellow solid is obtained via removal of water, then dissolved in 3 mL of CH3OH and transferred to an ampule. After the container is purged with argon for 5 min, a calculated amount of thiirane is added. The ampule is sealed and kept in a 50
C oil bath for 24 h. Then the mixture is transferred to a flask, evaporated to dryness, and stored under argon. 2. Synthesis of Disulfide Cross-Linked PEI. Thiolated PEI (1.5 g) is dissolved in 3 mL of methanol and 1.5 mL of dimethyl sulfoxide (DMSO) and is stirred for 48 h. The reaction solution is precipitated in diethyl ether three times and the product is dried under high vacuum to give a light yellow viscous glue or solid. 3.2 Synthesis of Linear and Branched PDMAEMA

Methacrylate- and acrylate-based polycations (such as PDMAEMA) have been widely investigated due to convenient synthesis by free-radical polymerization and the possibility to optimize their properties by copolymerization of a number of ionic and nonionic comonomers. However, high cytotoxicity impeded PDMAEMA prospects in gene delivery and prompted the development of reducible PDMAEMA. Reducible PDMAEMA can be prepared as follows [5]: 1,4-bis(2-(thiobenzoylthio)prop-2-yl)benzene mediated RAFT polymerization of DMAEMA produces α,ω-dithioester-functionalized PDMAEMA, which is then converted into α,ω-dithiol-ended PDMAEMA oligomer by aminolysis, and reducible PDMAEMA is synthesized by oxidation of the terminal thiol (as shown in Scheme 3). The typical procedure is as follows: 1. Synthesis of α,ω-dithioester-functionalized PDMAEMA. 1,4-Bis(2-(thiobenzoylthio)prop-2-yl)benzene (BTBP) mediated RAFT polymerization of DMAEMA is performed in THF at 60
C. The polymerization solution containing DMAEMA (1.0 g), BTBP (80.0 mg) and AIBN (AIBN: BTBP ¼ 1:20) is added into a glass ampoule, thoroughly

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Scheme 3 Synthesis of linear reducible PDMAEMA (Reproduced from [5] with permission of Elsevier)

Scheme 4 Synthesis of hyperbranched reducible PDMAEMA (Reproduced from [25] with permission of ACS)

deoxygenated, sealed under vacuum, and placed in a thermostated water bath at 60
C for 48 h. The polymer, α,ω-dithioester-terminated PDMAEMA, is obtained by precipitation into hexane and isolated by filtration. 2. Synthesis of α,ω-dithiol-ended PDMAEMA oligomer. The α,ω-dithioester functionalized PDMAEMA (0.5 g) is dissolved in THF (4 mL) containing a few drops of aqueous sodium bisulfite (Na2S2O4), hexylamine (0.2 mL) is added after the reaction mixture is purged of oxygen by bubbling with N2 for 30 min. The reaction mixture is stirred for 3 h under N2, then

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added dropwise to a tenfold hexane and the polymer is collected by filtration. 3. Synthesis of linear reducible PDMAEMA. Hexylamine (0.2 mL) and DMSO (0.2 mL) are added to solution of the α,ω-dithiol-ended PDMAEMA oligomer in methanol, and the reaction mixture is stirred at room temperature under oxygen atmosphere. The solvent is then removed and reducible PDMAEMA is dissolved in THF and then added dropwise to hexane, the polymer is collected by filtration. 1,2-Bis(2-(3-methylbuta-1,3-dien-2-yloxy)ethyl) disulfane mediated RAFT polymerization of DMAEMA in the presence of biodegradable disulfide-base dimethacrylate (DSDMA) produces PDMAEMA with a hyperbranched structure; each branch point in the polymer is linked by a disulfide bond, and the degree of branching (DB) can be tuned by the amount of DSDMA (Scheme 4) [25]. The detailed procedure is as follows: DMAEMA (316 mg, 2.0 mmol), 1,2-bis(2-(3-methylbuta-1,3-dien-2-yloxy)ethyl) disulfane (24 mg, 0.05 mmol), DSDMA (29 mg, 0.1 mmol) and AIBN (1.64 mg, 0.01 mmol) are dissolved in 3.0 mL of dimethylacetamide. Aliquots are transferred to three different vials, which are then sealed with rubber septa. Each vial is deoxygenated by purging with nitrogen for 30 min prior to placement in water bath at 70
C. The vials are taken out at 20, 42, and 65 h. Immediate cooling is applied via an ice-water bath and radical quenching is applied via air exposure. The polymers are collected after precipitation twice from dichloromethane to hexane and then dried under vacuum. 3.3 Synthesis of Bioreducible Poly (Amido Amine) s and Poly(Amino Ester)s via Michaeladdition Polymerization

Via Michael-addition polymerization between primary amines and cystamine bisacrylamide (CBA), a series of linear and hyperbranched bioreducible poly(amido amine)s have been developed. Polymerization conditions can control the topology of the produced polymers.

3.3.1 Reaction Temperature Controls the Topology of Reducible Poly(Amino Ester)s [32]

Generally, when diamine, triamine, or polyamine monomers react with cystamine bisacrylamide, linear polymers are obtained. But, in some cases, Michael-addition polymerization of disulfide-based diacrylate and equimolar N-methylethylenediamine forms ABB’ type intermediates first; 2
amines (formed) are inactive below 40
C, leading to the formation of linear poly(amino ester) via AB-type intermediates (B0 does not participate in the reaction). However, elevated temperature activates 2
amines (formed), and they participate in the addition reaction, which results in the formation of hyperbranched reducible poly(amino ester)s via

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Scheme 5 Synthesis of linear reducible poly(amino ester) (Reproduced from [32] with permission of ACS)

ABB0 -type intermediates (both B and B0 participate in the reaction). The DB of the hyperbranched polymers obtained increases with the increase of temperature. Therefore, reducible polymer topology from linear to hyperbranched can be tuned simply by varying the polymerization temperature. The detailed polymerization procedure is as follows: disulfide-based diacrylate (0.524 g, 2.0 mmol) and N-methylethylenediamine (0.148 g, 2.0 mmol) are added into 2 mL of chloroform. After the polymerization is performed in the dark at 25
C (or 50
C) for a certain time, the reaction mixture is poured into 50 mL of diethyl ether under vigorous stirring. The polymer is collected and purified by reprecipitation from a chloroform solution into diethyl ether followed by drying under vacuum for 1 day at room temperature (Schemes 5 and 6). 3.3.2 Monomer Feed Ratio Controls the Topology of Reducible Poly(Amido Amine)s [35]

The amino units in a trifunctional amine have different reactivity in Michael-addition polymerization of trivalent amine with bisacrylamide/bisacrylate, and hence the topology of the produced polymers can be tuned simply via varying reaction conditions such as the molar ratio of trivalent amine to bisacrylamide/bisacrylate. Generally, in Michael-addition polymerization of 1-(2-aminoethyl)piperazine (AEPZ) with equal molar N,N0 -cystamine bisacrylamide (CBA), AB-type intermediate formed at the starting stage and 2
amine (produced during the polymerization) do not take part in reaction due to high steric hindrance, which results in the formation of linear poly(amido amine) as shown in Scheme 7, the detailed procedure is as follows: CBA (260 mg, 1.0 mmole) and AEPZ (129.2 mg, 1.0 mmole) are added into a vial and dissolved in methanol–water mixture (3.0 mL, 8/2 v/v), then the polymerization is performed in the dark at 50
C. The reaction is allowed to

Synthesis of Bioreducible Polycations with Controlled Topologies

35

Scheme 6 Synthesis of hyperbranched reducible poly(amino ester) (Reproduced from [32] with permission of ACS)

proceed for 5 days yielding a viscous solution. The resulting reducible polymer is obtained via precipitating in cool acetone and drying under vacuum for 4 h at room temperature. However, in Michael-addition polymerization of AEPZ with double molar CBA, A2B-type intermediate produced at starting stage, 2
amine (produced during the polymerization) participates in the polymerization in late stage, which leads to the formation of hyperbranched reducible poly(amido amine) (Scheme 8). In a typical experiment, CBA (520 mg, 2.0 mmole) and AEPZ (129 mg, 1.0 mmole) are added into a vial and dissolved in methanol–water mixture (5.0 mL, 8/2 v/v). After the polymerization has been performed in the dark at 50
C for 120 h, the polymerization is stopped via decreasing the temperature to room temperature.

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HN

H N

H N

NH2

N

S-S

H N

O

H N

S-S

O

O

N

N

NH2

O AB

H N

S-S

H N

N O

O

N

H N n

Scheme 7 Synthesis of linear reducible poly(amido amine) (Reproduced from [35] with permission of ACS)

N

1 HN

NH2

R

S S

R' NH

B (AEPZ)

Michael addtion

R S S

2

S S R

R

A (CBA)

R A2B

reducible hyperbranched polymer

Scheme 8 Synthesis of hyperbranched reducible poly(amido amine). (Reproduced from [35] with permission of ACS)

The resulting reducible polymer is obtained via precipitating in cool acetone and drying under vacuum for 3 h at room temperature.

4

Notes 1. In preparing linear reducible PDMAEMA, the molar ratio of BTBP/AIBN should be over 20, otherwise the end units of some PDMAEMA will not be functionalized with thioester. 2. In preparing hyperbranched reducible PDMAEMA, the molar percentage of biodegradable disulfide-based dimethacrylate should be below 50%, otherwise, cross-linked PDMAEMA is formed. 3. In Michael-addition polymerization, the monomer concentration should not be too high ( 0.7, the sample is highly polydisperse and the values of the hydrodynamic radius given by the instrument are probably meaningless. Short periods of sonication of the sample may help to homogenize the solution [30]. 3.5

Cell Culture

1. The day prior to transfection, use trypsin to detach actively dividing cells from the culture flask (Corning). 2. After addition of 15 mL of DMEM supplemented with 10% of serum, the cells are counted. 3. Finalize the protocol of the experiment in order to determine how many wells are needed. Notably, transfection experiments are usually done using duplicates or triplicates. 4. The required volume of cell suspension to obtain a confluence of 50–80% the following day is then calculated (for 911 and 911-Luc cells, we use 200,000 cells/well) (see Note 10). Dilute the cell suspension with a volume of culture medium which allows the addition of 1 mL of culture medium into each well of the 24-well plate. 5. The cells are then incubated at the appropriate cell culture conditions (a humidified tissue culture incubator at 37
C and 5% CO2).

3.6

Cell Transfection

Before starting the DNA/siRNA transfection experiment, the cells should be examined using a microscope. This observation allows for verification of the confluence of the cells and also of the (good) shape of the cells. The peptide–DNA or siRNA complexes are

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generated just before transfection of the cells. For a transfection performed in duplicate, the following protocols are used. 3.6.1 DNA Transfection

1. The tube containing the plasmid DNA is thawed and gently vortexed. Take 4 μg of plasmid DNA (for one duplicate) and dilute them in 100 μL of sterile (0.20 μM filtered) 150 mM NaCl. 2. The tube containing a 1 mg/mL solution of LAH4 is thawed. The tube is then vigorously vortexed—a short sonication using a bath sonicator may be additionally performed—before withdrawing the desired amount of peptide (see Notes 11 and 12). Complete to 100 μL with sterile 150 mM NaCl. 3. Mix the DNA with the peptide, centrifuge very shortly to pull down all the drops, and let the tube at room temperature for about 15 min. 4. Add to the mixture serum-free medium (if applicable) to obtain a final volume of 1 mL; 0.5 mL of the transfection mixture are then put in each well of the duplicate. 5. Remove the culture medium from the cells by aspiration. If the cells that are used adhere well, they may be rinsed with PBS in order to remove all serum containing proteins from the wells (see Note 13). 6. Add 0.5 mL of LAH4/DNA transfection medium into each well of the duplicate and incubate at 37
C for 2–4 h. 7. Remove carefully the transfection medium and replace it with 1 mL of complete culture medium (DMEM supplemented with 100 units/mL penicillin, 100 μg/mL streptomycin, and 10% of fetal calf serum). 8. Incubate at 37
C for 28–48 h, depending on the cell type. 9. Analyze the cells for luciferase expression (see Notes 14 and 15) and protein content or proceed for the MTT cell viability assay (see below).

3.6.2 siRNA Transfection

1. Thaw tubes containing the siRNA-Luc. Take 0.46 μg (3.5 μL of the 10 μM siRNA solution) of siRNA-Luc (to obtain a final concentration in the wells of 50 nM). Complete to 30 μL with NaCl 150 mM. 2. The tube containing a 1 mg/mL solution of LAH4 is thawed. The tube is then vigorously vortexed—a short sonication step using a bath sonicator may be additionally applied—before withdrawing the desired amount of peptide (see Notes 11 and 12). Complete to 30 μL with NaCl 150 mM (see Note 14). 3. Of note, transfection complexes of related composition should be prepared with a control siRNA such as a siRNA-GFP. This

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control is important in order to ascertain that the silencing is gene specific. 4. Mix the siRNA with the peptide, centrifuge very shortly to pull down all the drops, and let the tube at room temperature for about 15 min. 5. Add to the mixture serum-free medium (if possible) to obtain a final volume of 0.7 mL; 0.350 mL of the transfection mixture are then added to each well of the duplicate. 6. Remove the culture medium from the cells by aspiration. If the cells that are used adhere well, the cultures may be rinsed with PBS in order to remove serum containing proteins from the wells. 7. Add 0.350 mL of LAH4/siRNA transfection medium into each well of the duplicate and incubate at 37
C for 2–4 h. 8. Remove carefully the transfection medium and replace it with 1 mL of complete culture medium (DMEM supplemented with 100 units/mL penicillin, 100 μg/mL streptomycin, and 10% of fetal calf serum). 9. Incubate at 37
C for 28–48 h, depending on the cell type. 10. Analyze the cells for luciferase expression and protein content or proceed for the MTT cell viability assay (see below) (see Notes 16 and 17). 3.7

Luciferase Assay

To determine luciferase activity, the following protocol is used: 1. Remove carefully the culture medium from the 24-well plates. 2. Add 250 μL of lysis buffer to each well. 3. After 10 min, the cell lysate is recovered and transferred into 1.5 mL Eppendorf tubes 4. Centrifuge the tubes for 5 min at 10,000  g to pellet debris. 4. Take 50 μL of the supernatant of each tube and transfer them into the wells of a white 96-well plate. 5. Measure the bioluminescence using a luminometer which automatically injects 100 μL of buffer assay and 100 μL of luciferin solution. 6. Read luminescence over 10 s. 7. Remove the luciferase background (obtained with the lysate from nontransfected cells) from each value. 8. Calculate the light units for 10 s/250 μL of sample. 9. After having measured the protein content (see below), express the efficiency as light units/10 s/mg (or μg) of protein.

3.8

Protein Content

The protein determination procedure described here is valid for the Bradford protein assay from Bio-Rad (see Note 18).

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1. Transfer 2 μL of cell lysate to a transparent 96-well plate. Of note, add a control with 2 μL of lysis buffer in order to obtain the background value. 2. Add 200 μL of Bradford reagent which was diluted five times in water. 3. Mix gently. 4. Read absorbance at λ ¼ 595 nm and determine the protein content in each sample by using a bovine serum albumin standard curve. 3.9 MTT Cell Viability Assay

This assay is used to measure the cell viability (see Note 18). It therefore allows for an evaluation of the cytotoxicity of the transfection complexes. 1. The transfection should be performed using the same experimental conditions as those used for the luciferase assay (DNA or siRNA). 2. Instead of removing the medium and lysing the cells after 28–48 h of transfection, add to the medium MTT reagent at a final concentration of 0.5 mg/mL per well. 3. Cells are incubated at 37
C and 5% CO2 for 3 h. 4. Remove the medium from the cells. 5. Add 500 μL DMSO to each well to dissolve the formazan crystals. 6. Transfer 100 μL from each sample into a transparent 96-well plate and measure absorption at λ ¼ 570 nm. 7. Untreated cells serve as control (¼100% cell viability).

3.10 Flow Cytometry for the Evaluation of the Delivery Efficiency of the siRNA

By using a fluorescently labeled siRNA and flow cytometry it is possible to quantify the amount of nucleic acid that is associated with or endocytosed by the cells. 1. Complexes are made fluorescent by using a fluorescently labeled siRNA-Luc (an FITC group is conjugated to the 30 end of the antisense strand). 2. After preparation of the complexes as described for cell transfection, they are added to 911-Luc cells (final siRNA concentration ¼ 50 nM). 3. After 3 h of incubation at 37
C, cells are washed with cold PBS and harvested in 1 mM EDTA in PBS and 10,000 cells are then analyzed by flow cytometry.

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Notes 1. Solid-phase peptide synthesis is based on a cycle of repetitive (and therefore potentially automated) steps where one amino acid after the other is connected to the growing peptide chain. The synthesis starts with the most C-terminal residue of the sequence which is attached to beads of a solid resin and progresses toward the N-terminus (i.e., reverse to the conventional reading of polypeptide sequences). By attaching the intermediate and final products to the solid support it is possible to freely exchange the solvents simply by washing steps where the resin is retained by a ceramic filter funnel (or column) without the need of recrystallization or other types of intermediate purification. A synthetic cycle starts with one amino acid being attached to the resin by its carboxyl group and the amine being protected (e.g., Fmoc). In a first step the protection group is cleaved from the chain which liberates the amino group for the following reaction. In parallel the amino acid N-terminal (n1) to the one already on the resin (n) is activated chemically; thus its carboxyl group reacts with the amine of the residue already coupled to the solid support. The amine group of this amino acid is protected (e.g., by a Fmoc group); thus it cannot polymerize with its like. By mixing an excess of this activated amino acid with the chain on the resin and incubation of this reaction mixture by several minutes the polypeptide chain grows by one residue, and a new cycle can start by removal of the protection group. Ideally the reaction yield of each step is >99%, but critical steps may occur where the cleavage of the protection group or the addition of the next amino acid residues is hindered (e.g., sterically). In such cases, increased temperatures, double couplings, extended couplings, a change in chemistry and/or solvents, or the use of pseudoproline residues may be helpful (e.g., [31]) and references cited therein). 2. The optimal flow to obtain best resolution on this type of column would be about three times increased, but this is not always possible as the resulting back pressure of the HPLC system and column exceeds the capacity of the instrument. 3. More recent studies suggest that a more reliable method consists in at least three cycles of dissolving 1 mg/mL peptide in 2 mM HCl (respecting a few-fold excess over the TFA ions) and lyophilization [32]. If the peptide is purchased from a commercial source make sure you have information about the counterions. Note that the molecular weight to be considered during the preparation of a solution is much different if you

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have TFA (2777 + 9  113), acetate (2777 + 9  59), or Cl (2777 + 9  36) counterions to nine cationic sites of the polypeptide sequence. 4. We recommend the storage of the peptide powder over longer time periods at 20
C. Therefore, before usage the container and the powder have to be first equilibrated thoroughly to reach room temperature, which can take many minutes, even hours. In order to avoid the condensation of water into the peptide powder and its recipient, it is important that during this process the vial remains tightly sealed. Thereafter, weight the appropriate amount of peptide and resuspend LAH4 at a final concentration of, e.g., stock solution of 1 mg/mL. Before transferring the vial with the peptide back into the cold assure that it is tightly sealed with, for example, Parafilm. Avoid repeated freezing and thawing of the stock solution. Prepare rather small aliquots (500 μL in 1.5 mL Eppendorf tubes). 5. If the condensation of the nucleic acids is very efficient, then the nucleic acids are no more accessible to SYBRSafe. This, in turn, results in the absence of a nucleic acid stain in the agarose gel. This is for example the case in Fig. 3 with the highest LAH4/DNA ratio. 6. The peptide LAH4 has five positive charges at neutral pH: indeed, the C-terminus is amidated and therefore it does not present a negative charge. Also of note, at acidic pH, the imidazole groups of the histidine residues become protonated and this adds four positive charges to the peptide (net charge ¼ +9 at pH 5). 7. In our hands, 2.5 to 5 times more peptide of the LAH4 family is required to retard the migration of siRNAs when compared with plasmid DNA. This indicates that complex formation and interactions of the cationic peptides with plasmid DNA are different when compared to siRNA [13]. 8. The quality of plasmid DNA may influence the transfection process and only siRNA and DNA of high quality should be used. It is recommended that the optical density ratio OD260/ OD280 is 1.8. 9. For biophysical experiments we prefer to use an LAH4 stock solution at low pH, where the peptide is monomeric [30], and therefore, mixes with DNA in the most homogenous manner. In this manner, changing back and forth between acidic and neutral pH gives reproducible DLS measurements for the transfection complexes. 10. It is usually recommended to transfect cells at a confluence of 50–80%. However, the optimal cell density for efficient DNA or siRNA transfection may vary between cell types. In any case, it is important to keep the same seeding protocol for a given

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cell type between the experiments. It is also important, especially when using primary cells, to use cells having approximately the same number of passages. 11. The optimal peptide–nucleic acid ratio for transfection has to be determined for each new cell line. 12. The optimal peptide–nucleic acid ratio may be different for a given cell line for plasmid DNA and siRNA. For example, in our hands using 911 cells, the optimal weight/weight peptide–nucleic acid ratio for siRNA delivery is higher than the one for plasmid DNA transfection. 13. The DNA and siRNA transfection process with LAH4 and derivatives is reduced in the presence of serum [14]. Therefore, when possible, avoid addition of serum—or at low %—during the 2–3 h of incubation. 14. Optimization of each step of the transfection process should be undertaken if the DNA/siRNA transfection efficiency is low. Besides testing various peptide–nucleic acid ratios, other parameters may be changed: concentration of nucleic acids per well; duration of incubation of the complexes with the cells; buffer conditions employed during complexation (e.g., water seems to give better results with the LAH4 peptides for the delivery of siRNA [13]); confluence of the cells; percentage of serum in the medium during the 2–4 h of incubation. Another way to improve the transfection consists in not replacing the transfection medium after the 2 to 4 h incubation step. In this case, add culture medium containing serum into the wells. 15. It has been shown for PEI–DNA complexes that just after addition of the complexes to the cells, a short centrifugation step of the culture plates at 1000 rpm (¼138  g) for 5 min increases the transfection efficiency [33]. This step may also be performed in order to increase the transfection of DNA/siRNA by LAH4 and derivatives. 16. When using cells expressing a reporter gene such as luciferase for measuring the efficiency of siRNA knockdown, one has to take into account the fact that the efficiency may vary from one cellular clone to another. Indeed, the number of integrated copies probably influences the level of expression and thus also the efficiency of silencing. Cells expressing a reporter gene are very useful for a first evaluation of the siRNA transfection efficiency of compounds. However, for further experiments, the best manner to evaluate the siRNA delivery efficiency of a system consists in targeting endogenous genes such as GAPDH. 17. For siRNA experiments, it is important to choose the amount of peptide which gives the highest transfection efficiency

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associated with the lowest cytotoxicity. Indeed, cell toxicity may lead to nonspecific knockdown of genes. 18. For determination of the amount of protein, use a detergentcompatible detection kit. Notably, the total cellular protein content is also an indicator of cell viability.

Acknowledgments We thank Christopher Aisenbrey for his assistance on the section concerning the chemical synthesis of the peptides. References 1. Elbashir SM, Harborth J, Lendeckel W, Yalcin A, Weber K, Tuschl T (2001) Duplexes of 21-nucleotide RNAs mediate RNA interference in cultured mammalian cells. Nature 411 (6836):494–498. https://doi.org/10.1038/ 35078107 2. Kopatz I, Remy JS, Behr JP (2004) A model for non-viral gene delivery: through syndecan adhesion molecules and powered by actin. J Gene Med 6(7):769–776. https://doi.org/ 10.1002/jgm.558 3. Behr JP (1997) The proton sponge: A trick to enter cells the viruses did not exploit. Chimia 51(1–2):34–36 4. Kichler A, Leborgne C, Coeytaux E, Danos O (2001) Polyethylenimine-mediated gene delivery: a mechanistic study. J Gene Med 3 (2):135–144. https://doi.org/10.1002/jgm. 173 5. Neuberg P, Kichler A (2014) Recent developments in nucleic acid delivery with polyethylenimines. Adv Genet 88:263–288. https://doi. org/10.1016/B978-0-12-800148-6.000092 6. Sonawane ND, Szoka FC Jr, Verkman AS (2003) Chloride accumulation and swelling in endosomes enhances DNA transfer by polyamine-DNA polyplexes. J Biol Chem 278 (45):44826–44831. https://doi.org/10. 1074/jbc.M308643200 7. Gao WW, Chan JM, Farokhzad OC (2010) pH-responsive nanoparticles for drug delivery. Mol Pharm 7(6):1913–1920. https://doi. org/10.1021/mp100253e 8. Douat C, Aisenbrey C, Antunes S, Decossas M, Lambert O, Bechinger B, Kichler A, Guichard G (2015) A cell-penetrating foldamer with a

bioreducible linkage for intracellular delivery of DNA. Angew Chem Int Ed Engl 54 (38):11133–11137. https://doi.org/10. 1002/anie.201504884 9. Kos P, Lachelt U, Herrmann A, Mickler FM, Doblinger M, He D, Krhac Levacic A, Morys S, Brauchle C, Wagner E (2015) Histidine-rich stabilized polyplexes for cMetdirected tumor-targeted gene transfer. Nanoscale 7(12):5350–5362. https://doi.org/10. 1039/c4nr06556e 10. Midoux P, Pichon C, Yaouanc JJ, Jaffres PA (2009) Chemical vectors for gene delivery: a current review on polymers, peptides and lipids containing histidine or imidazole as nucleic acids carriers. Br J Pharmacol 157 (2):166–178. https://doi.org/10.1111/j. 1476-5381.2009.00288.x 11. Kichler A, Leborgne C, Marz J, Danos O, Bechinger B (2003) Histidine-rich amphipathic peptide antibiotics promote efficient delivery of DNA into mammalian cells. Proc Natl Acad Sci U S A 100(4):1564–1568. https://doi.org/10.1073/pnas.0337677100 12. Lan Y, Langlet-Bertin B, Abbate V, Vermeer LS, Kong X, Sullivan KE, Leborgne C, Scherman D, Hider RC, Drake AF, Bansal SS, Kichler A, Mason AJ (2010) Incorporation of 2,3-diaminopropionic acid into linear cationic amphipathic peptides produces pH-sensitive vectors. Chembiochem 11(9):1266–1272. https://doi.org/10.1002/cbic.201000073 13. Langlet-Bertin B, Leborgne C, Scherman D, Bechinger B, Mason AJ, Kichler A (2010) Design and evaluation of histidine-rich amphipathic peptides for siRNA delivery. Pharm Res

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27(7):1426–1436. https://doi.org/10.1007/ s11095-010-0138-2 14. Mason AJ, Leborgne C, Moulay G, Martinez A, Danos O, Bechinger B, Kichler A (2007) Optimising histidine rich peptides for efficient DNA delivery in the presence of serum. J Control Release 118(1):95–104. https://doi.org/10.1016/j.jconrel.2006.12. 004 15. Mason AJ, Martinez A, Glaubitz C, Danos O, Kichler A, Bechinger B (2006) The antibiotic and DNA-transfecting peptide LAH4 selectively associates with, and disorders, anionic lipids in mixed membranes. FASEB J 20 (2):320–322. https://doi.org/10.1096/fj. 05-4293fje 16. Prongidi-Fix L, Sugawara M, Bertani P, Raya J, Leborgne C, Kichler A, Bechinger B (2007) Self-promoted cellular uptake of peptide/ DNA transfection complexes. Biochemistry 46(40):11253–11262. https://doi.org/10. 1021/bi700766j 17. Bechinger B, Lohner K (2006) Detergent-like actions of linear amphipathic cationic antimicrobial peptides. Biochim Biophys Acta 1758 (9):1529–1539. https://doi.org/10.1016/j. bbamem.2006.07.001 18. Bechinger B (2015) The SMART model: soft membranes adapt and respond, also transiently, in the presence of antimicrobial peptides. J Pept Sci 21(5):346–355. https://doi.org/10. 1002/psc.2729 19. Georgescu J, Munhoz VH, Bechinger B (2010) NMR structures of the histidine-rich peptide LAH4 in micellar environments: membrane insertion, pH-dependent mode of antimicrobial action, and DNA transfection. Biophys J 99(8):2507–2515. https://doi. org/10.1016/j.bpj.2010.05.038 20. Zhang TT, Kang TH, Ma B, Xu Y, Hung CF, Wu TC (2012) LAH4 enhances CD8+ T cell immunity of protein/peptide-based vaccines. Vaccine 30(4):784–793. https://doi.org/10. 1016/j.vaccine.2011.11.056 21. Moulay G, Leborgne C, Mason AJ, Aisenbrey C, Kichler A, Bechinger B (2017) Histidine-rich designer peptides of the LAH4 family promote cell delivery of a multitude of cargo. J Pept Sci 23(4):320–328. https://doi. org/10.1002/psc.2955 22. Liu Y, Kim YJ, Ji M, Fang J, Siriwon N, Zhang LI, Wang P (2014) Enhancing gene delivery of adeno-associated viruses by cell-permeable peptides. Mol Ther Methods Clin Dev 1:12. https://doi.org/10.1038/mtm.2013.12

23. Fenard D, Genries S, Scherman D, Galy A, Martin S, Kichler A (2013) Infectivity enhancement of different HIV-1-based lentiviral pseudotypes in presence of the cationic amphipathic peptide LAH4-L1. J Virol Methods 189 (2):375–378. https://doi.org/10.1016/j. jviromet.2013.02.005 24. Fenard D, Ingrao D, Seye A, Buisset J, Genries S, Martin S, Kichler A, Galy A (2013) Vectofusin-1, a new viral entry enhancer, strongly promotes lentiviral transduction of human hematopoietic stem cells. Mol Ther Nucleic Acids 2:e90. https://doi.org/10. 1038/mtna.2013.17 25. Majdoul S, Seye AK, Kichler A, Holic N, Galy A, Bechinger B, Fenard D (2016) Molecular determinants of vectofusin-1 and its derivatives for the enhancement of lentivirally mediated gene transfer into hematopoietic stem/progenitor cells. J Biol Chem 291 (5):2161–2169. https://doi.org/10.1074/ jbc.M115.675033 26. Gemmill KB, Muttenthaler M, Delehanty JB, Stewart MH, Susumu K, Dawson PE, Medintz IL (2013) Evaluation of diverse peptidyl motifs for cellular delivery of semiconductor quantum dots. Anal Bioanal Chem 405(19):6145–6154. https://doi.org/10.1007/s00216-013-69822 27. Abbate V, Liang W, Patel J, Lan Y, Capriotti L, Iacobucci V, Bui TT, Chaudhuri P, Kudsiova L, Vermeer LS, Chan PF, Kong X, Drake AF, Lam JK, Bansal SS, Mason AJ (2013) Manipulating the pH response of 2,3-diaminopropionic acid rich peptides to mediate highly effective gene silencing with low-toxicity. J Control Release 172(3):929–938. https://doi.org/10.1016/j. jconrel.2013.09.033 28. Caplen NJ, Parrish S, Imani F, Fire A, Morgan RA (2001) Specific inhibition of gene expression by small double-stranded RNAs in invertebrate and vertebrate systems. Proc Natl Acad Sci U S A 98(17):9742–9747. https://doi. org/10.1073/pnas.171251798 29. Aisenbrey C, Bechinger B, Grobner G (2008) Macromolecular crowding at membrane interfaces: adsorption and alignment of membrane peptides. J Mol Biol 375(2):376–385. https:// doi.org/10.1016/j.jmb.2007.10.053 30. Marquette A, Mason AJ, Bechinger B (2008) Aggregation and membrane permeabilizing properties of designed histidine-containing cationic linear peptide antibiotics. J Pept Sci 14(4):488–495. https://doi.org/10.1002/ psc.966

Cationic Peptides for the Delivery of Nucleic Acids 31. Harzer U, Bechinger B (2000) Alignment of lysine-anchored membrane peptides under conditions of hydrophobic mismatch: a CD, 15N and 31P solid-state NMR spectroscopy investigation. Biochemistry 39 (43):13106–13114 32. Andrushchenko VV, Vogel HJ, Prenner EJ (2007) Optimization of the hydrochloric acid

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concentration used for trifluoroacetate removal from synthetic peptides. J Pept Sci 13 (1):37–43. https://doi.org/10.1002/psc.793 33. Boussif O, Zanta MA, Behr JP (1996) Optimized galenics improve in vitro gene transfer with cationic molecules up to 1000-fold. Gene Ther 3(12):1074–1080

Chapter 4 Covalent Fluorophore Labeling of Oligonucleotides and Generation of Other Oligonucleotide Bioconjugates Cornelia Lorenzer and Johannes Winkler Abstract Oligonucleotide conjugates have already reached considerable importance in life science research and oligonucleotide drug development. Since the preparation of oligonucleotide conjugates depends critically on the chemical nature of the used ligand and linker, there is no general and universal procedure. Here, we present a detailed, quick, and facile protocol for attaching fluorescent dyes or cross-linkers of variable chemical stability to oligonucleotides at 30 - or 50 -aminoalkyl handles. Purification and removal of educts and side-products and structural verification by gel electrophoresis and mass spectrometry are presented. Aspects for adapting this protocol for other reaction sites at the oligonucleotide are discussed. We highlight important issues for generating oligonucleotide conjugates with other molecules, including peptide, proteins, and small molecules for receptor-targeting applications. The methodology is suitable for oligonucleotides with various modifications, including stabilized antisense, siRNAs, and miRNAs. Key words Bioconjugation, Oligonucleotides, Fluorescent dyes, Linkers, Drug targeting, Tracking

1

Introduction During the last decades, oligonucleotides have become important tools for many applications of biomedical research, and more recently also as therapeutic agents [1]. The modular nature of nucleic acids allows for facile and straightforward, automated synthesis, and their target recognition properties based on hydrogen bonding to the complementary nucleobase confers a number of distinct applications in research and drug development [2–4]. Among those, the use as probes for in situ hybridization, in microarrays, for some PCR applications, or the tracking of oligonucleotides in vitro or in vivo requires attachment of fluorescent or other labels to chemically synthesized oligonucleotides [5]. In addition, bioconjugation of various ligands has an important role in the development of therapeutically active oligonucleotide agents [6, 7]. The tethering of proteins, peptides or small

Manfred Ogris and Haider Sami (eds.), Nanotechnology for Nucleic Acid Delivery: Methods and Protocols, Methods in Molecular Biology, vol. 1943, https://doi.org/10.1007/978-1-4939-9092-4_4, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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molecule receptor ligands for targeting applications may be instrumental for overcoming the inherent, yet not sufficiently solved issues of pharmaceutical development. In particular, quick renal elimination must be avoided, oligonucleotide agents need to reach the targeted tissue and to be taken up efficiently into the cytosol of target cells [8]. Receptor-binding ligands may aid all of these critical steps, and thus oligonucleotide bioconjugates have been pushed into the focus of therapeutic development of antisense, siRNA, and aptamer therapeutics [9]. As an example, the covalent attachment of N-acetyl galactosamine has substantially facilitated the progress of both siRNA and antisense oligonucleotides towards their clinical use [10, 11]. However, this approach is limited to hepatocyte targeting and thus directed to liver diseases, and other targeting moieties seem to be needed for expanding the target space of oligonucleotide therapeutics. Besides conferring receptor-targeting properties, the attachment of certain ligands to oligonucleotides may also influence pharmacokinetics and biodistribution in a nonspecific way, by increasing the molecular weight [12–14] or by modulating lipophilicity [7] or plasma protein binding. In addition, cellular uptake and endosomal escape are aimed to be increased by attaching cationic small molecules or peptides [15–19]. The preparation of oligonucleotide conjugates can be achieved by attaching linkers at either the 30 - or 50 -terminus, at the 20 -position of the ribose, at the phosphate linkage, or at the nucleobases [6]. While many applications demand a stable covalent linkage, some approaches benefit from a linker that is cleaved under certain conditions. A receptor-targeted carrier often needs to be removed from the oligonucleotide cargo after successful cellular uptake in order to allow gene silencing or other effects [20]. Depending on the position, the linker and the particular ligand, optimal preparation protocols vary. Small molecule ligands can in some cases be attached directly during automated oligonucleotide synthesis by using appropriate phosphoramidite building blocks or modified solid supports [21]. Most commonly, ligand attachment takes place after solid-phase oligonucleotide assembly by direct reaction of a handle introduced during solid phase synthesis, or by first tethering a linker molecule at the 30 -or 50 -position, which is then used as an anchor for the ligand [6]. In this chapter, we give step-by-step instructions for covalently attaching a fluorescent dye to a synthetic oligonucleotide through an active ester reaction to an aminoalkyl handle. In addition we show a procedure how to attach linkers at the 30 - or 50 -end for further bioconjugation of respective molecules with stable (triazole or maleimide) or labile (disulfide) linkers, and indicate possible modifications of the protocol for a variety of ligands, attachment sites, and linker chemistries.

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2

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Materials In most instances, oligonucleotides will be provided by commercial suppliers. For this protocol, obtain the respective sequence with an aminoalkyl modification at either the 30 - or 50 end. If selfsynthesizing the oligonucleotide, use standard methods on an automated oligonucleotide synthesizer, and incorporate an aminoalkyl linker at the 30 - or 50 -position by using a modified solid support, or an appropriate phosphoramidite building block, respectively (see Note 1). Ensure complete removal of primary aminecontaining impurities such as buffers and ammonia salts through an adequate desalting procedure (see Note 2).

2.1 Oligonucleotide Desalting and Precipitation

1. RNase-free water. 2. Sodium acetate solution, 0.3 M. 3. Butanol or isopropanol. 4. Ethanol, 75%. 5. Centrifuge, min. 11,600  g. 6. Nanospectrophotometer.

2.2 Fluorescent Dye Attachment

1. Fluorescent dye with desired fluorophore properties as an NHS ester. A variety of fluorescent labels with a large number of different excitation and emission wavelengths are commercially available for amine attachments, including the Alexa Fluor®, Dylight®, ATTO® series. 2. Dimethylsulfoxide (DMSO), water-free, or dimethylformamide (DMF), water-free, for preparing stock solutions. 3. Sodium tetraborate buffer, 0.1 M, pH 8.5, freshly prepared. 4. pH-meter or pH test strips able to differentiate between 0.2 units in a range of pH 7 to 9. 5. Thermomixer or shaker for Eppendorf tubes.

2.3

Purification

1. Sephadex G 25, G-50 or equivalent (see Note 3). ¨ KTA pure or equivalent FPLC system. 2. A 3. Superdex 75 10/300 GL column or equivalent. 4. HPLC system. 5. Reversed phase column for oligonucleotide separations. 6. Microdialysis devices. 7. Nanospectrophotometer.

2.4 Gel Electrophoresis

1. Urea–acrylamide gel: For two gels: mix 6.8 g urea, 7.5 mL of a 40% acrylamide–bisacrylamide stock solution (29:1 ratio acrylamide–bis-acrylamide), and 1.5 mL 10 TBE buffer. Add 7.5 μL tetramethyl ethylenediamine and 75 μL ammonium

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persulfate (10%) and immediately prepare the gels. Leave at r.t. for polymerization (min 1 h). 2. Mini Protean III gel electrophoresis system or equivalent. 3. Loading dye: 20% sucrose, 0.01% bromophenol blue. 4. Running buffer: 1 TBE. 5. Fluorescence gel imager able to detect respective fluorescent dye, or alternatively gel imager for detection of methylene blue- or ethidium bromide-stained gels. 2.5 Linker Attachment Reactions

1. Linker for generation of disulfide (succinimidyl 3-(2-pyridyldithio)propionate, SPDP), maleimide (sulfosuccinimidyl 4-(N-maleimidomethyl)cyclohexane-1-carboxylate, SMCC), or triazole (dibenzocyclooctyne-N-hydroxysuccinimidyl ester, DBCO) linkages. 2. Dimethylsulfoxide, water-free (DMSO). 3. Dulbecco’s phosphate buffered saline (10, DPBS). 4. pH-meter or pH test strips able to differentiate between 0.2 units in a range of pH 7 to 9. 5. Thermomixer or shaker for Eppendorf tubes.

3

Methods

3.1 Desalt Oligonucleotide

This step is necessary if not done by the commercial provider, in order to remove remnants of amine-containing salts and reagents such as Tris, which interfere with linker attachment (see Note 2). When dissolving an oligonucleotide provided in lyophilized form, be sure to use an amine-free buffer such as bicarbonate, PBS, or RNase-free water. Avoid Tris buffers. 1. For precipitation from sodium acetate solution, dissolve up to 100 nMol of the oligonucleotide in 50 μL RNase-free water (see Note 4). 2. Add 200 μL sodium acetate (0.3 M) and 1 mL butanol or isopropanol and mix. 3. After incubation for 20 min or o/n at 70  C, centrifuge for 10 min at 11,600–15,000  g, decant the alcohol and wash twice in 750 μL ice-cold 75% ethanol. Remove the supernatant again and dissolve the dried pellet in 50 μL RNase-free water. 4. Determine oligonucleotide concentration by absorption measurement preferably in a nanophotometer using the sequencederived extinction coefficient and the Lambert–Beer law (see Note 5). 5. Dilute to a stock solution of 1 mM or 2 mM.

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65

Fig. 1 Reaction scheme of the attachment of an N-hydroxysuccinimide (NHS) activated fluorescent dye to an aminohexyl modified oligonucleotide 3.2 Direct Attachment of a Fluorescent Dye to an Aminoalkyl Oligonucleotide

1. Dilute the oligonucleotide in RNase-free water to a 500 μM solution, typically 25 nmol in 50 μL. 2. Prepare a stock solution of the appropriate fluorescent dye N-hydroxy succinimidyl (NHS) ester, 10–20 mM in anhydrous DMSO (see Note 6). 3. Add a threefold volume of 0.1 M tetraborate buffer (150 μL) to 50 μL of 500 μM oligonucleotide. 4. Add a tenfold molar surplus of dye to the oligonucleotide solution; 1.25–2.5 μL of 10–20 mM stock solution of the NHS ester activated fluorescent dye (Fig. 1). 5. Mix overnight at r.t. in a shaker or thermomixer (see Note 7).

3.3

Purification

Remove excess dye by precipitation, gel filtration, size exclusion chromatography, dialysis, or HPLC. 1. Precipitation: Add 20 μL sodium acetate solution (0.3 M) and 1 mL isopropanol to the reaction mix, vortex, and keep at 70  C for a minimum of 20 min (see Note 8). Centrifuge for 10 min at 15,000  g, decant the alcohol and wash twice in 750 μL ice-cold 75% ethanol. Remove the supernatant again and dissolve the dried pellet in 20–50 μL RNase-free water. 2. Gel filtration (benchtop): Prepare a slurry of 2 mL Sephadex G-25 or an equivalent gel filtration matrix (see Note 3) in water. Fill the slurry into a benchtop column and let the liquid reach the top of the gel matrix. Add the reaction mixture and wait again till the liquid reaches the top of the gel. Add nuclease-free water and collect fractions of 500 μL each, until the entire product has eluted. Check fractions by measurements in a nanospectrophotometer. Pool relevant fractions and concentrate them in a speed-vac, or remove the solvent by lyophilization. ¨ kta pure chroma3. Size exclusion chromatography: Equip an A tography system (or equivalent) with a Superdex 75 10/300 GL column (or equivalent). As elution solvent use 5 DPBS, pH 7.4 and a flow rate of 0.5 mL/min. Collect fractions of

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0.5 mL with detection at 260 nm (total volume approx. 2.5–3.0 mL) and optionally also at the absorption wavelength of the dye. Pool fractions containing the conjugate and concentrate to a 100 to 200 μM solution in a 3 kDa MWCO centrifugal filter. 4. Dialysis. Fill reaction solution into a microdialysis device (MWCO 1–3 kDa for most dyes). Dialyze for at least 2 h or o/n against PBS or a solvent of choice. For optimal removal of unreacted dye, exchange dialysis medium repeatedly. 5. HPLC. Equip the HPLC system with a reversed phase column for oligonucleotide separations (Clarity 5 μ RP18 250  4.60 mM, Phenomenex, or similar). Use triethylammonium acetate buffer in nuclease-free water, pH 7 and acetonitrile as solvents and equilibrate the column with 8% acetonitrile (ACN). Load the reaction mixture onto the column and elute with a linear gradient of 8–44% ACN in TEAA in 60 min with a flow rate of 1 mL/min. Collect product containing fractions. Depending on the loading capacity of the column, the separation may have to be performed several times with aliquots of the reaction mixture. 3.4 Gel Electrophoresis

Optional: For verification of successful attachment and quantification of attachment yields, separate the purified conjugate on a denaturing urea–acrylamide gel. 1. Prepare urea–acrylamide gel and leave at r.t for complete polymerization o/n. 2. Mix 20–50 pmol conjugate with loading buffer. As controls, mix uncoupled oligonucleotide, and uncoupled dye (20 pmol each) with loading buffer. 3. Load gel lanes with conjugate and controls. 4. Run gel electrophoresis in 1 TBE at 100 V until bromophenol blue has nearly reached the bottom of the gel (ca. 90 min). 5. Transfer the gel to a fluorescence imager and record image upon excitation at the respective wavelength of the attached fluorophore. If no appropriate imager is available, the gel can be stained with methylene blue (0.1%) or an alternative DNA staining solution (see Note 9), but the applied amount of oligonucleotide conjugate would need to be increased to approx. 500 pmol. For an efficient and complete reaction, a single band will be visible. After staining, both the labeled and the unlabeled oligonucleotides will give distinct bands; however, their separation is dependent on the characteristics (molecular weight, polarity) of the attached fluorophore. Most unseparated, unattached dyes will migrate close to the front of the gel.

Oligonucleotide Conjugations

3.5 Mass Spectrometry

3.6 Attachment of Cross-Linkers for Generating Disulfide, Maleimide, or Triazole Linkages 3.6.1 For Disulfide Linkages

67

Mix 30 μL of a 25 μM oligonucleotide sample with 15 μL 7.5 M ammonium acetate solution and incubate at r.t. for 1 h [22]. Add 115 μL ethanol and store o/n at 80  C. Centrifuge for 30 min at 11,600–15,000  g and 0  C to collect the precipitate. Decant the ethanol immediately after centrifugation and wash the pellet with 100 μL ice-cold 75% ethanol. Centrifuge again for 20 min at 13,800  g and 0  C. Air-dry the pellet and dissolve it in 50 μL nuclease-free water. Determine concentration by measurement with a nanospectrophotometer. Dilute the sample to a 10 μM solution with 10 mM ammonium acetate in RNase-free H2O and analyze it in an electrospray mass spectrometer. 1. Dilute the oligonucleotide in RNase-free water to a 500 μM solution, typically 25 nmol in 50 μL. 2. Prepare a stock solution of SPDP linker, 20 mM in anhydrous DMSO. 3. Add an appropriate volume of 10  DPBS, pH 8.5, to reach a final concentration of 5 DPBS (e.g., 56.25 μL 10 DPBS buffer to 50 μL oligonucleotide solution). 4. Add a fivefold molar surplus of linker (e.g., 6.25 μL of 20 mM SPDP to 50 μL of 500 μM oligonucleotide solution (Fig. 2)). 5. Mix 1–3 h at r.t. in a shaker or thermomixer. 6. Remove excess of linker reagent by precipitation (see Note 10 and Subheading 3.4). 7. Attach ligand of choice by reaction in PBS or an appropriate buffer (see Note 11).

3.6.2 For Maleimide Linkages

1. Dilute the oligonucleotide in RNase-free water to a 500 μM solution, typically 25 nmol in 50 μL. 2. Prepare a stock solution of Sulfo-SMCC linker, 10 mM in RNase-free water. 3. Add an appropriate volume of 10 DPBS, pH 7.7, to reach a final concentration of 5 DPBS (e.g., 175 μL 10 DPBS buffer to the oligonucleotide solution). 4. Add a 50-fold molar surplus of linker (e.g., 125 μL of 10 mM Sulfo-SMCC to 50 μL of 500 μM oligonucleotide solution (Fig. 2)). 5. Mix 3 h at r.t. in a shaker or thermomixer. 6. Remove excess of linker reagent by gel filtration, size exclusion chromatography, dialysis, or HPLC (see Subheading 3.4). 7. Attach ligand of choice by reaction in PBS or an appropriate buffer. (see Note 10).

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Fig. 2 Structures of disulfide-, maleimide-, and triazole-generating linkeroligonucleotide conjugates after attachment of bifunctional tethers to an aminohexyl modified oligonucleotide 3.6.3 For Triazole Linkages

1. Dilute the oligonucleotide in RNase-free water to a 500 μM solution, typically 25 nmol in 50 μL. 2. Prepare a stock solution of DBCO linker, 10 mM in anhydrous DMSO. 3. Add an appropriate volume of 10 DPBS, pH 7.4, to reach a final concentration of 5 DPBS (e.g., 56.25 μL 10 DPBS buffer to the oligonucleotide solution). 4. Add a tenfold molar surplus of linker (e.g., 6.25 μL of 10 mM DBCO to 50 μL of 500 μM oligonucleotide solution (Fig. 2)). 5. Mix 2 h at r.t. in a shaker or thermomixer. 6. Remove excess of linker reagent by precipitation, gel filtration, size exclusion chromatography, dialysis, or HPLC (see Subheading 3.4). 7. Attach ligand of choice by reaction in PBS or an appropriate buffer. (see Note 10).

Oligonucleotide Conjugations

4

69

Notes 1. This protocol is primarily designed for attaching ligands via aminoalkyl linkers at either the 30 -or the 50 -end of oligonucleotides. Corresponding oligonucleotides can be ordered from most suppliers, as they are easily prepared during automated oligonucleotide synthesis by either using a modified solid phase (for 30 -attachments) or an additional coupling step with an aminoalkyl phosphoroamidite (for 50 -attachments). Aminohexyl linkers are most common, but other alkyl chain lengths are available. For some particular applications, which demand internal labeling, aminoalkyl-modified nucleobases such as 5-[N-(trifluoroacetylaminohexyl)-3-acrylamido]-20 -deoxyuridine may be used. The protocol works with unmodified DNA and RNA oligonucleotides, but also with a variety of chemical modifications, which include phosphorothioates, 20 -methyl, 20 -ethyl, 20 -methoxyethyl, 20 -fluoro, locked nucleic acids, many base modifications, and backbone modifications. However, sitespecific equimolar conjugation relies on the presence of a single primary amine functionality, and thus alkylamine-containing modifications such as bridged nucleic acids with an amine should be avoided. 2. The success of the NHS-ester amine reaction depends critically on the absence of any other primary amines in the reaction mixture, and amine-containing buffer such as Tris will interfere with the chemical reaction. From our experience, commercial oligonucleotides vary in their quality and in the presence of residual chemicals from synthesis and purification depending on the supplier and selected purification method. Cleavage from the solid support and deprotection after automated synthesis is usually achieved with ammonium or methylamine solutions, and standard desalting procedures may fail to completely remove these reagents. After HPLC purification, traces of amines from the mobile phase can also be present in the final product. If unsure, it is advisable to first evaluate the conjugation reaction using a small amount of the substance. If the reaction is unsuccessful or gives poor yields, an additional desalting procedure is recommended. 3. Choose gel filtration matrix according to the length of the oligonucleotide: G-25 is suited for oligonucleotides longer than 10 bases, G-50 is a better choice for 20mer and longer oligonucleotides. 4. Other desalting methods are gel filtration in a benchtop column (see Subheading 3.3, step 2) or a spin column (fraction

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collection by centrifugation) or dialysis (see Subheading 3.3, step 4). 5. Because the contributions of individual nucleotides to the absorption of short nucleic acid sequences vary greatly, the calculation for nucleic acid concentration that is provided the software of many spectrophotometers is not valid for oligonucleotides. This calculation assumes an equal distribution of each of the four nucleotides, which holds true only for long sequences. Instead, calculate the specific extinction coefficient either by a nearest-neighbor calculation (available for example at www.basic.northwestern.edu/biotools/oligocalc.html) [23, 24], or by summing up individual values for each base (A: 15400, G: 11700, C: 7300, T: 8800; U: 9950). Concentration is calculated using the Lambert–Beer law: E ¼ ε  c  d. 6. Stock solutions for most fluorescent dye NHS esters can be prepared in DMF or DMSO. Consult the product information for the optimal solvent and recommended stock solution concentrations (usually 10–20 mM). To avoid hydrolysis, the use of water-free solvents is essential, and a stock solution in DMSO or DMF can be stored at 70  C and used for up to 2 months. 7. Overnight incubation is recommended for increasing attachment yields. For some reactions, if a high tethering ratio is not of major importance, the reaction can be shortened to 2 h at r.t. 8. The solution can be left at 70  C o/n to increase precipitation yield. Oligonucleotides with lipophilic modifications such as a full phosphorothioate backbone and/or a highly lipophilic dye, might fail to precipitate completely. In these cases, increase the volume of isopropanol or remove the solvent and resort to a different purification method. 9. Fluorescent DNA staining dyes include ethidium bromide, SYBR Green, and similar compounds. Although those dyes are generally more sensitive than the nontoxic methylene blue, they react less efficiently with short single stranded sequences than with duplexed nucleic acids, and also require relatively high oligonucleotide amounts compared to direct excitation of the attached fluorescent label. When using these staining procedures, molar amounts of oligonucleotides for optimal detection should be carefully examined. Cave: ethidium bromide and similar stains are carcinogenic. 10. Purification with gel filtration or RP or IEX-HPLC may be an alternative if precipitation is not sufficient or not possible in cases of lipophilic oligonucleotide modifications. Because the SPDP linker is easily cleaved in mildly reductive environments, prevention of degradation is dependent on the precise purification conditions.

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11. Practically all ligands with either a reactive thiol (to SPCC and SMCC) or an azido group (to DBCO) can be consecutively attached to the linker at this step. These include particularly proteins and peptides, for which inherent or engineered cysteines or azides can be used as potentially unique attachment sites, and small molecules with respective functional groups or derivatization. However, reaction conditions and buffers vary greatly depending on the selected ligand, and need to optimized separately for each reaction. Therefore, more detailed descriptions are beyond the scope of this article. Reactions can be performed under native (e.g., Tris, PBS, borate-based buffers) or denaturing conditions (urea or guanidinium chloride buffers), at r.t., or at elevated temperatures, and with reaction times between a few minutes and several days. For optimization, monitoring of the reaction by an appropriate analytical procedure such as HPLC, SEC, or gel electrophoresis is recommended. The choice of a purification method is also dependent of the characteristic and molecular size of the ligand, and respective possibilities include RP- or IEX-HPLC, SEC, and dialysis. References 1. Khvorova A, Watts JK (2017) The chemical evolution of oligonucleotide therapies of clinical utility. Nat Biotechnol 35(3):238–248. https://doi.org/10.1038/nbt.3765 2. Baumann V, Lorenzer C, Thell M, Winkler A-M, Winkler J (2017) RNAi-mediated knockdown of protein expression. Methods Mol Biol 1654:351–360. https://doi.org/10.1007/ 978-1-4939-7231-9_26 3. Dirin M, Winkler J (2013) Influence of diverse chemical modifications on the ADME characteristics and toxicology of antisense oligonucleotides. Expert Opin Biol Ther 13 (6):875–888. https://doi.org/10.1517/ 14712598.2013.774366 4. Baumann V, Winkler J (2014) miRNA-based therapies: strategies and delivery platforms for oligonucleotide and non-oligonucleotide agents. Future Med Chem 6(17):1967–1984. https://doi.org/10.4155/fmc.14.116 5. Asseline U (2006) Development and applications of fluorescent oligonucleotides. Curr Org Chem 10(4):491–518. https://doi.org/10. 2174/138527206776055349 6. Winkler J (2013) Oligonucleotide conjugates for therapeutic applications. Ther Deliv 4 (7):791–809. https://doi.org/10.4155/tde. 13.47

7. Patel PL, Rana NK, Patel MR, Kozuch SD, Sabatino D (2016) Nucleic acid bioconjugates in cancer detection and therapy. ChemMedChem 11(3):252–269. https://doi.org/10. 1002/cmdc.201500502 8. Lorenzer C, Dirin M, Winkler A-M, Baumann V, Winkler J (2015) Going beyond the liver: progress and challenges of targeted delivery of siRNA therapeutics. J Control Release 203:1–15. https://doi.org/10.1016/ j.jconrel.2015.02.003 9. Ming X, Laing B (2015) Bioconjugates for targeted delivery of therapeutic oligonucleotides. Adv Drug Deliv Rev 87(0):81–89. https://doi.org/10.1016/j.addr.2015.02. 002 10. Nair JK, Willoughby JLS, Chan A, Charisse K, Alam MR, Wang Q, Hoekstra M, Kandasamy P, Kel’in AV, Milstein S, Taneja N, O’Shea J, Shaikh S, Zhang L, van der Sluis RJ, Jung ME, Akinc A, Hutabarat R, Kuchimanchi S, Fitzgerald K, Zimmermann T, van Berkel TJC, Maier MA, Rajeev KG, Manoharan M (2014) Multivalent N-acetylgalactosamine-conjugated siRNA localizes in hepatocytes and elicits robust RNAi-mediated gene silencing. J Am Chem Soc 136(49):16958–16961. https://doi.org/ 10.1021/ja505986a

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11. Prakash TP, Graham MJ, Yu J, Carty R, Low A, Chappell A, Schmidt K, Zhao C, Aghajan M, Murray HF, Riney S, Booten SL, Murray SF, Gaus H, Crosby J, Lima WF, Guo S, Monia BP, Swayze EE, Seth PP (2014) Targeted delivery of antisense oligonucleotides to hepatocytes using triantennary N-acetyl galactosamine improves potency 10-fold in mice. Nucleic Acids Res 42(13):8796–8807. https://doi. org/10.1093/nar/gku531 12. Winkler J (2015) Therapeutic oligonucleotides with polyethylene glycol modifications. Future Med Chem 7(13):1721–1731. https://doi. org/10.4155/fmc.15.94 13. Gaziova Z, Baumann V, Winkler A-M, Winkler J (2014) Chemically defined polyethylene glycol siRNA conjugates with enhanced gene silencing effect. Bioorg Med Chem 22 (7):2320–2326. https://doi.org/10.1016/j. bmc.2014.02.004 14. Shokrzadeh N, Winkler A-M, Dirin M, Winkler J (2014) Oligonucleotides conjugated with short chemically defined polyethylene glycol chains are efficient antisense agents. Bioorg Med Chem Lett 24(24):5758–5761. https:// doi.org/10.1016/j.bmcl.2014.10.045 15. Winkler J, Urban E, Noe C (2005) Oligonucleotides conjugated to short lysine chains. Bioconjug Chem 16(4):1038–1044. https:// doi.org/10.1021/bc049729d 16. Winkler J, Gilbert M, Kocourkova A, Stessl M, Noe C (2008) 2 ’-O-lysylaminohexyl oligonucleotides: modifications for antisense and siRNA. ChemMedChem 3(1):102–110. https://doi.org/10.1002/cmdc.200700169 17. Winkler J, Saadat K, Diaz-Gavilan M, Urban E, Noe C (2009) Oligonucleotide-polyamine conjugates: influence of length and position of 2 ’-attached polyamines on duplex stability and

antisense effect. Eur J Med Chem 44 (2):670–677. https://doi.org/10.1016/j. ejmech.2008.05.012 18. Dirin M, Urban E, Lachmann B, Noe CR, Winkler J (2015) Concise postsynthetic preparation of oligonucleotide-oligopeptide conjugates through facile disulfide bond formation. Future Med Chem 7(13):1657–1673. https:// doi.org/10.4155/fmc.15.109 19. Dirin M, Urban E, Noe CR, Winkler J (2016) Fragment-based solid-phase assembly of oligonucleotide conjugates with peptide and polyethylene glycol ligands. Eur J Med Chem 121:132–142. https://doi.org/10.1016/j. ejmech.2016.05.001 20. Winkler J (2011) Nanomedicines based on recombinant fusion proteins for targeting therapeutic siRNA oligonucleotides. Ther Deliv 2 (7):891–905. https://doi.org/10.4155/tde. 11.56 21. Lonnberg H (2009) Solid-phase synthesis of oligonucleotide conjugates useful for delivery and targeting of potential nucleic acid therapeutics. Bioconjug Chem 20(6):1065–1094. https://doi.org/10.1021/bc800406a 22. Shah S, Friedman SH (2008) An ESI-MS method for characterization of native and modified oligonucleotides used for RNA interference and other biological applications. Nature Prot 3(3):351–356 23. SantaLucia J Jr (1998) A unified view of polymer, dumbbell, and oligonucleotide DNA nearest-neighbor thermodynamics. Proc Natl Acad Sci U S A 95(4):1460–1465 24. Kibbe WA (2007) OligoCalc: an online oligonucleotide properties calculator. Nucleic Acids Res 35(suppl_2):W43–W46. https://doi.org/ 10.1093/nar/gkm234

Chapter 5 Synthetic Oligonucleotides in SPECT/CT In Vivo Imaging: Chemical Modifications, In111 Complex Formation, Incorporation into Drug Delivery Systems Maxim Antopolsky Abstract Here in we describe a solid phase synthesis of oligonucleotides bearing unnatural moiety appropriate for complex formation with In111 as well as their deprotection, isolation, and purification. We also present methods for oligonucleotides/In111 complex formulation with single and double stranded oligonucleotides of RNA nature and give an example of preparation method for one supramolecular drug delivery system (DDS) consisting of radiolabeled siRNA and positively charged peptide. Key words Oligonucleotides, In111 complex, Solid phase RNA synthesis, Radiolabeling, Drug delivery system

1

Introduction The role of therapeutic oligonucleotides (antisense, siRNA, ribozymes, microRNA, etc.) in present drug related research is difficult to overestimate: they play more and more significant role [1]. One of the key points in this research is the problem of bio distribution of oligonucleotides [2, 3], their ability to target the desired site [4] of action staying intact [5] or, at least, preserving their biological and therapeutic activity. Among the methods to investigate this SPECT/CT in vivo imaging plays an important role as far as it allows following the target radiolabeled substance in the body of laboratory animals for relatively long time without killing them [6]. On the other hand, the number of radioactive labels suitable for SPECT/CT in vivo imaging is not that big (Table 1). Only few isotopes are acceptable for this method, and selection of appropriate one as well as method of it chemical incorporation development are usually critical in each particular case. That especially concerns oligonucleotides that by their chemical nature do not display any functional group, which can be employed for either iodination or

Manfred Ogris and Haider Sami (eds.), Nanotechnology for Nucleic Acid Delivery: Methods and Protocols, Methods in Molecular Biology, vol. 1943, https://doi.org/10.1007/978-1-4939-9092-4_5, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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Table 1 Radionuclides acceptable for use in SPECT/CT in vivo imaging Radionuclide

Half-life

Main gamma energy

99m

Tc

6.0 h

141 keV

111

In

2.8 days

171,245 keV

123

I

13.2 h

159 keV

direct complex formation with Indium 111 (In111) or Technetium 99 (Tc99). Introduction of the functional group appropriate for further radiolabeling into an oligonucleotide sequence can be achieved in several different ways. Most straightforward seems to be introduction of aliphatic amino group on the 30 or 50 terminus of oligonucleotide during solid phase synthesis followed by coupling with any active ester displaying either phenolic function (for iodination) or complex forming moiety (DTPA; DOTA). Such a way we tried in our study with luciferase siRNA, but were not really successful. In case of phenolic moiety introduction, radiolabeling yields were very poor (about 30% only), and all our attempts to couple amino-modified strand of luciferase siRNA with either DOTA or DTPA succinimidyl active esters failed. It was, most probably, due to electrostatic interaction between negatively charged DOTA and DTPA and negatively charged molecule of oligonucleotide as well. That is why we decided to introduce complex forming functional group directly during solid phase synthesis of sense strand of luciferase siRNA. The only commercially available monomer for solid phase oligonucleotide synthesis (SPOS) was 50 -dimethoxytrityl-5-[N-ethylenediaminetetraacetate, monoacetylaminoethyl-3-acrylamido]-20 -deoxyuridine,30 -[(2-cyano ethyl)-(N,N-diisopropyl)]-phosphoramidite from Glen Research. The monomer was introduced on the 30 -terminus of sense strand of luciferase siRNA employing standard RNA synthesis protocol, US-II universal solid support, and TOM protected RNA monomers (Subheadings 2 and 3). Deprotected, isolated, and purified oligonucleotide was then radiolabeled by complex formation with In111, and by addition of equivalent amount of antisense strand followed by annealing the double strand luciferase siRNA was formed (Subheadings 4 and 5) and used for direct SPECT/CT in vivo imaging as well as preparation of several supramolecular formulations (Subheadings 6 and 7) that were then also used in SPECT/CT experiments.

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Fig. 1 Synthesis of sense strand of luciferase siRNA 30 -modified by introduction of EDTA moiety

2

Materials

2.1 Materials (Synthesis, Deprotection, and Purification of Sense Strand of Luciferase siRNA (Fig. 1)

1. Biosset ASM-800 DNA synthesizer [7]. 2. US-II universal solid support for oligonucleotide synthesis [8]. 3. 400 nmol columns for ASM-800 DNA synthesizer. 4. 50 -Dimethoxytrityl-5-[N-ethylenediaminetetraacetate,monoacetylaminoethyl-3-acrylimido]-20 -deoxyuridine, 30 -[(2-cyano ethyl)-(N,N-diisopropyl)]-phosphoramidite.

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5. T-CE Phosphoramidite [8]. 6. A-TOM-CE Phosphoramidite [8]. 7. C-TOM-CE Phosphoramidite [8]. 8. U-TOM-CE Phosphoramidite [8]. 9. Acetonitrile—distilled over CaH2 (see Note 1). 10. 5-Ethylthio-1H-Tetrazole. 11. Deblock solution—3% solution of dichloroacetic acid in dichloromethane. 12. Oxidation solution: 0.05 M iodine solution in a mixture of tetrahydrofuran (THF), pyridine (Py) and water (8:1:1). 13. Capping solution A: 10% solution of acetic anhydride in THF. 14. Capping solution B: 10% solution of methylimidazole in THF–Py (8:1) mixture. 15. Dichloromethane. 16. 3-((Dimethylamino-methylidene)amino)-3H-1,2,4-dithiazole-3-thione. 17. 7 M ammonia solution in methanol. 18. Methanol. 19. 32% ammonia solution in water. 20. Polyethylene tubes (45 mm  4 mm inner diameter). 21. Tosoh Bioscience TSKgel DEAE-2SN 4,6 mm  25 cm, particle size 5 μm column. 22. XTERRA C18 5 μm, 4.6 mm  15 cm column. 23. Triethylamine trihydrofluoride (TEA.3HF). 24. Glen-Pak RNA quenching buffer [8]. 25. Dimethyl sulfoxide. 26. RNase-free water. 27. RNase-free pipettes and Eppendorf tubes. 28. Triethylamine (TEA). 29. 5 mm  20 cm Sephadex G-10 desalting column. 30. HPLC Buffer A: 0.1 M triethylammonium acetate (TEAE) in RNase-free water, pH ¼ 8.0. 31. HPLC buffer B: 0.1 M TEAE in 70% acetonitrile, pH ¼ 8.0. 32. IEX HPLC Buffer A: 0.1 M NH4Ac in 20% acetonitrile/ RNase-free water, pH ¼ 6.7. 33. IEX HPLC Buffer B: 0.1 M NH4Ac and 0.4 M NaClO4 in 20% acetonitrile/RNase-free water, pH ¼ 6.7. 34. Speedvac. 35. Rotavapor.

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36. 1 ml sterile pipette tips. 37. Glen-Pak RNA quenching buffer [8]. 38. 80% acetic acid in RNase-free water. 39. Freeze dryer LyoDry compact. 2.2 Materials (In111 with EDTA-Modified Luciferase siRNA Complex Formation (Figs. 2 and 3)

1. Eppendorf thermomixer. 2. Sterile 15 ml test tubes with screw caps. 3. Sterile pipette 1 ml tips. 4. Pipette 100 μl–1000 μl. 5. RP C18 cartridges. 6. 400 μl of luciferase siRNA sense strand-EDTA modified oligonucleotide solution (2.8 mg/ml) in RNase-free water. 7. 400 μl of luciferase siRNA antisense strand oligonucleotide solution (2.8 mg/ml) in RNase-free water. 8. 0.1 M NH4Ac buffer in RNase-free water, pH ¼ 6.0. 9. 0.05 M NH4Ac buffer in RNase-free water, pH ¼ 5.5. 10. In111Cl3 solution in 1 ml of water. 11. Acetonitrile. 12. RNase-free water. 13. 60% acetonitrile in RNase-free water. 14. 25 ml round bottom flask. 15. Rotavapor.

2.3 Materials (Complex Formation of Radiolabeled Luc siRNA with “Pepfect” Peptide (Fig. 4))

3

1. All the materials mentioned in Subheading 2.2. 2. Vortex. 3. 2 mmol “Pepfect” phosphoric acid.

peptide

solution

in

10

μmol

4. 1 ml conical vessel with screw cap and two glass balls inside.

Methods

3.1 Methods (Synthesis, Deprotection, and Purification of Sense Strand of Luciferase siRNA (Fig. 1)

1. Pack US-II universal solid support into four ASM-800400 nmol synthetic columns and run coupling–oxidation (sulfurizing)–capping–deblocking cycle on the ASM-800 DNA synthesizer employing standard RNA coupling/phosphorothioate protocols and reagents with 50 -dimethoxytrityl-5-[N-ethylenediaminetetraacetate,monoacetylaminoethyl-3-acrylimido]-2’-deoxyUridine,30 -[(2-cyanoethyl)-(N,N-diisopropyl)]-phosphoramidite at all four columns (see Note 2).

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Fig. 2 In111-luciferase siRNA complex formation

2. Run RNA synthesis of sense strand luciferase siRNA employing standard RNA protocols and oxidation by sulfurizing on first and second cycles of the synthesis at all four columns. 3. Perform cleavage/deprotection procedure in the following manner: pack columns in the bottom of polyethylene tubes, put the bottom of the tubes into 1 ml pipette tips and then to the sterile Eppendorf tubes. Fill the free volume in the tubes over the column with 3.5 M ammonia methanol solution (about 1.5 ml) and let the solution pass through the columns

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Fig. 3 HPLC traces of radiolabeled lucifirase siRNA (XTERRA C18 5 μm 4,6 mm  15 cm column and linear gradient from 0 to 50% of HPLC buffer B in 30 min, flow rate 1 ml/min. (Buffer A—HPLC Buffer A—0.1 M TEAE in RNasefree water, pH ¼ 8.0; Buffer B—HPLC Buffer B—0.1 M TEAE in 70% acetonitrile, pH ¼ 8.0))

within 30–40 min. Wash the columns with 32% aqueous ammonia (2 ml) and combine all solutions in one screw capped vial. 4. Heat the vial for 8 h at 65  C, then cool it down to room temperature and evaporate to dryness. 5. Fully dissolve the residue in 230 μl of DMSO, add 120 μl of TEA and mix gently, then add 150 μl of TEA.3HF and heat the mixture at 65  C for 2.5 h. 6. Cool the mixture down and add 3.5 ml of Glen-Pak RNA quenching buffer [8] and immediately after this desalt it on Sephadex G-10 column using RNase-free water as a buffer. 7. Perform HPLC trityl-on purification using XTERRA C18 5 μm, 4.6 mm  15 cm column and linear gradient from 10% to 70% of HPLC buffer B in 30 min, flow rate 1 ml/min. Target compound comes from the column as the last peak at about 23 min. 8. Concentrate fractions containing target oligonucleotide to the volume 0.1 ml, add 1.0 ml of 80% acetic acid in RNase-free water to remove dimethoxytrityl group from 50 -terminus, keep the solution for 5 min at room temperature and then remove acetic acid by evaporation. 9. Make final IEX HPLC purification using Tosoh Bioscience TSKgel DEAE-2SN 4,6 mm  25 cm, particle size 5 μm column and linear gradient from 0 to 50% IEX HPLC B buffer in 30 min, flow 1 ml/min. Target compound comes from the column at about 18 min (see Note 3).

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Fig. 4 Preparation of luc siRNA In111 radiolabeled and “pepfect” peptide complex

10. Perform desalting of pure oligonucleotide on 5 mm  20 cm Sephadex G-10 desalting column using RNase-free water as a running buffer (see Note 4). 11. Freeze-dry the isolated pure oligonucleotide and keep it at t 20  C. 3.2 Methods (In111 with EDTA-modified Luciferase siRNA Complex Formation (Figs. 2 and 3) (See Note 5)

1. Into sterile 15 ml test tube with screw cap add 100 μl of luciferase siRNA sense strand-EDTA modified oligonucleotide solution (2.8 mg/ml) in RNase-free water, 100 μl of 0.1 M NH4Ac buffer in RNase-free water, pH ¼ 6.0 and finally 1 ml of In111Cl3 solution in 1 ml of water (radioactivity 560 MBq). 2. Put the test tube into Eppendorf thermomixer and leave stirring at 300RM and 85  C for 30 min. 3. Put into the reaction mixture 100 μl of luciferase siRNA antisense strand oligonucleotide solution (2.8 mg/ml) in RNase-

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free water, leave stirring for additional 10 min and then let it cool down within 20 min to the room temperature. 4. Prepare RP C18 cartridge passing through it 3 ml of acetonitrile followed by equilibration of the cartridge with 3 ml of 0.05 M NH4Ac buffer in RNase-free water, pH ¼ 5.5. The procedure is made by attaching 5 ml sterile syringe with appropriate buffer to the cartridge and passing the buffer through the cartridge at the flow approximately 1 drop a minute. 5. Apply cooled reaction mixture obtained in 3.2.3 on the cartridge and wash it with 3 ml of 0.05 M NH4Ac buffer in RNase-free water, pH ¼ 5.5. The procedure is performed in the same manner as described in 3.2.4. 6. Cleave radiolabeled double stranded luciferase siRNA from the cartridge with 60% acetonitrile in RNase-free water collecting fractions of approximately 0.5 ml (see Note 6). 7. Measure the radioactivity of the fractions in dose calibrator. Combine three fractions that contain main radioactivity in the 25 ml round bottom flask. 8. Remove acetonitrile by evaporation at the rotavapor and concentrate the solution to the volume of 0.5 ml (see Note 7). 9. Perform HPLC analysis of In111 radiolabeled luciferase siRNA. Use XTERRA C18 5 μm, 4.6 mm  15 cm column and linear gradient from 0 to 50% of HPLC buffer B in 30 min, flow rate 1 ml/min. (Buffer A—HPLC Buffer A—0.1 M TEAE in RNase-free water, pH ¼ 8.0; Buffer B—HPLC Buffer B— 0.1 M TEAE in 70% acetonitrile, pH ¼ 8.0). 3.3 Methods (Complex Formation of Radiolabeled Luc siRNA with “Pepfect” Peptide (Fig. 4)

4

1. Perform all the steps from 1 to 8 in Subheading 3.2. 2. Transfer the obtained solution of In111 radiolabeled Luc siRNA into 1 ml conical vessel with screw cap and two glass balls inside. Add either 100 μl (for 1:10 ratio) or 200 μl (for 1:20 ratio) of 2 mmol “Pepfect” peptide solution in 10 μmol phosphoric acid and vortex it during 5–10 s. The ready-made complex solution should be clear and colorless (see Note 8).

Notes 1. Acetonitrile that is used for solid phase oligonucleotide synthesis should be dry. Preferably, it must be distilled over calcium hydride; any traces of water may dramatically influence the efficiency of couplings. 2. Capping step after first coupling on the solid support must be performed properly; better to perform double capping: that can influence the purity of final product very much.

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3. IEX HPLC should be used at the final step of purification: as far as oligonucleotide contains phosphorothioate moieties, at RP HPLC pure compound is presented by several isomers and might come out by several peaks. In case of Ion exchange HPLC the oligonucleotide comes out from the column as a single peak. 4. After 20 -hydroxy groups deprotection only RNase-free water must be used at each and every step of the experiment in order to prevent enzymatic degradation of oligonucleotide. 5. All the experiment must be performed into a Hot cell in order to prevent radiation spreading in the lab. 6. Cleavage of radiolabelled siRNA from RPC18 cartridge should be performed at the flow rate not faster than 1 drop/s to prevent increase of eluate volume. 7. Do not evaporate the eluate to dryness, always keep radiolabelled siRNA in solution. 8. It is better to add peptide solution to the reaction mixture directly while vortexing. References 1. Ma DDF, Rede T, Naqvi NA, Cook PD (2000) Synthetic oligonucleotides as therapeutics: the coming of age. Biotechnology 5:155–196 2. Geary RS, Norris D, Yu R, Bennett CF (2015) Pharmacokinetics, biodistribution and cell uptake of antisense oligonucleotides. Adv Drug Deliv Rev 87:46–51 3. Park J, Park J, Pei Y, Xu J, Yeo Y (2016) Pharmacokinetics and biodistribution of recentlydeveloped siRNA nanomedicines. Adv Drug Deliv Rev 104:93–109 4. Ming X, Laing B (2015) Bioconjugates for targeted delivery of therapeutic oligonucleotides. Adv Drug Deliv Rev 87:81–89

5. Otto P (1989) Investigation of the stability of oligonucleotides and polynucleotides. J Mol Struct THEOCHEM 188(3–4):277–288 6. de Vries EFJ, Vroegh J, Dijkstra G, Moshage H, Vaalburg W (2004) Synthesis and evaluation of a fluorine-18 labeled antisense oligonucleotide as a potential PET tracer for iNOS mRNA expression. Nucl Med Biol 31:605–612 7. Biosset high-performance DNA and RNA synthesisers (2011) http://biosset.com/. Accessed 1 Dec 2011 8. GlenResearch catalog (2018) http://www. glenresearch.com/Catalog/index1.php

Chapter 6 Synthesis of Polyethylenimine-Based Nanocarriers for Systemic Tumor Targeting of Nucleic Acids Wolfgang Ro¨dl, Alexander Taschauer, David Schaffert, Ernst Wagner, and Manfred Ogris Abstract Nucleic acid-based therapies offer the option to treat tumors in a highly selective way, while toxicity towards healthy tissue can be avoided when proper delivery vehicles are used. We have recently developed carrier systems based on linear polyethylenimine, which after chemical coupling of protein- or peptide-based ligands can form nanosized polyplexes with plasmid DNA (pDNA) or RNA and deliver their payload into target cells by receptor-mediated endocytosis. This chapter describes the synthesis of LPEI from a precursor polymer and the current coupling techniques and purification procedure for peptide conjugates with linear polyethylenimine. A protocol is also given for the formation and characterization of polyplexes formed with LPEI conjugate and pDNA. Key words Polyethylenimine, Polyethylene glycol, Molecular conjugates, EGF receptor, Targeting, Gene delivery

1

Introduction The standard treatment for solid cancers is usually surgery, followed by radiotherapy and treatment with chemotherapeutic drugs. Dose-limiting toxicity and resistance mechanisms often preclude a successful treatment of relapsing disease. Nucleic acid-based therapeutics offer the possibility to develop highly specific, tailor made therapies for the treatment of malignant diseases taking into account the genetic aberrations occurring in tumor cells compared to healthy body tissue. For gene therapy approaches, the gene of interest is cloned into an appropriate expression cassette and can either be incorporated into a viral vector, for example the widely used adenovirus; as an alternative, plasmids are cloned and produced in E. coli for nonviral delivery approaches [1, 2]. Several physical delivery methods for plasmid are also applicable in vivo, like electroporation, particle bombardment, or ultrasound

Manfred Ogris and Haider Sami (eds.), Nanotechnology for Nucleic Acid Delivery: Methods and Protocols, Methods in Molecular Biology, vol. 1943, https://doi.org/10.1007/978-1-4939-9092-4_6, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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enhanced delivery with microbubbles (see also Chapter 15 in this book, Vlaskou et al.). For systemic delivery, particle-mediated systems are commonly used, either based on lipids, polycations, or combinations thereof [3]. For polycation-based transfection systems, polyethylenimine (PEI) represents a kind of “golden standard” [4, 5]. PEIs are polymers with one of the highest charge densities: 1 mg of PEI contains approx. 23 μmol potentially protonatable amines, of which approx. 50% are protonated at pH 7 [6]. Due to its high positive charge density, nanosized particles, so called polyplexes, can be formed by electrostatic interaction after mixing PEI with nucleic acids containing a negatively charged phosphate backbone. PEI polyplexes, usually carrying a positive surface charge, bind to negatively charged cell surfaces mainly by interaction with proteoglycans and are thereafter internalized by adsorptive endocytosis [3, 7]. After internalization into endosomes and acidification by ATP driven proton pumps, additional amines become protonated leading to a so called proton sponge effect [8–10]: protons absorbed by PEI trigger the influx of chloride ions, which in turn leads to attraction of water molecules and the osmotic imbalance causes vesicle disruption and subsequent release of its payload into the cytoplasm. Besides branched PEI, linear PEI (LPEI) has been used for transfection studies in vitro and in vivo [11, 12]. Compared to branched PEI, LPEI exhibits a clearly improved transfection performance both in vitro and in vivo [13]. The synthesis of LPEI can be carried out by hydrolysis of the precursor polymer poly(2-ethyl-2-oxazoline) under highly acidic conditions [14, 15]. In order to obtain a product with fully biofunctional LPEI, care has to be taken that complete hydrolysis of the precursor is achieved, as residual N-acyl groups negatively affect the transfection efficiency [16]. Albeit being nonbiodegradable, LPEI-based vectors can be designed in a way that renders them well biocompatible. When polyplexes between LPEI and plasmid DNA are formed, they usually exhibit a positive surface charge and excess of free, not polyplex bound LPEI [17]. After intravenous injection, LPEI polyplexes rapidly interact with blood components and aggregate within the first vascular bed encountered, namely the lung [11, 12, 18]. This makes LPEI an excellent transfection reagent to achieve high transgene expression levels in the lung, where after crossing the endothelial barrier mostly pulmonary cells at the basolateral site are transfected [19]. At least two important parameters for this efficient transfection of lung tissue have been identified, namely the aggregation with blood platelets [20] and the presence of free, non-polyplex bound LPEI [17]. Although similar aggregation occurs with polyplexes based on branched PEI (BPEI), the latter polyplexes are far less efficient in lung transfection compared to LPEI [13]. Major reasons for this effect are the differences in the aggregation behavior and the dissociation behavior between PEI and plasmid DNA [21]. For optimal

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transfection via the systemic route, LPEI polyplexes have to be rather small. After intravenous injection, LPEI polyplexes rapidly aggregate in the blood stream, which causes their entrapment in the lung. On the cellular level, a reduced binding strength towards plasmid DNA of LPEI compared to BPEI has been observed: LPEI polyplexes are dissociated after endocytosis within intracellular vesicles [22] and release intact plasmid [23], which then is accessible for the translation machinery. In vivo, LPEI polyplexes initially aggregating in the lung redistribute to a considerable extent to the liver within the first minutes after injection [24], whereas BPEI polyplexes do not. To reduce the interaction with blood components and aggregation in blood, LPEI can be, similarly as described for stealth liposomes, chemically modified with the hydrophilic polymer polyethylene glycol (PEG) [25, 26]. Coupling of PEG to LPEI is on the one hand beneficial when it comes to systemic application of polyplexes in vivo, where the PEG component significantly reduces protein binding, allows blood circulation and passive accumulation in well vascularized tumors [24, 27–29]. After cellular internalization, excessive PEGylation can be disadvantageous, as it negatively affects the endosomal release of polyplexes [25]. Such limitations can be overcome by designing pH-responsive PEI conjugates, for example by coupling PEG via chemical bonds which are cleaved after acidification of the endosome [30, 31]. Alternatively, rather short PEG molecules can be used, which prevent aggregation in blood and still allow transfection of tumor cells in vivo [32]. As for PEG, protein ligands like transferrin or EGF can hamper endosomal release of targeted polyplexes when coupled to PEI polyplexes, but also interfere with proper particle condensation [33–35]. Hence, we developed a platform for the development of targeted, LPEI polyplexes, where short, peptidic ligands are utilized [36]. Here, the peptidic ligand is coupled to LPEI via a rather short 2 kDa PEG spacer. To ensure the formation of LPEI-PEG-peptide conjugates without crosslinking of LPEI molecules, a heterobifunctional PEG linker 3-(2-pyridyldithio)propionamide-PEG-N-hydroxysuccinimide ester (short: OPSS-PEG-NHS) is used. In the first coupling step, the NHS group reacts with one of the secondary amines in the LPEI chain forming a stable amide bond. In order to improve the reactivity of the NHS ester with secondary amines and to reduce ester hydrolysis, this reaction step is best carried out under waterfree conditions in absolute ethanol or other suitable solvents. After purification by cation exchange chromatography, which removes unreacted PEG, the distal OPSS group on the PEG linker is available for coupling to free thiols by forming a reducible disulfide bond [10, 36]. The thiopyridone group released during this reaction strongly absorbs at 343 nm, allowing UV control of the reaction. After a second cation exchange chromatography step, unreacted peptide and 2-thiopyridone are removed. The resulting LPEI-PEG-peptide conjugate forms nanosized polyplexes with

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plasmid DNA, but also with RNA. With such polyplexes, targeted delivery and high transfection efficiency can be obtained both in vitro and in vivo in tumors after intratumoral [36] or intravenous polyplex administration [32]. For a further preclinical development it is of note that LPEI-based polyplexes have already been applied in clinical trials [37], and that LPEI is available in GMP grade from the company Polyplus in France (www.polyplus-transfection.com).

2

Materials With the exception of the materials mentioned below, all reagents can be obtained from standard lab suppliers. Please choose the quality grade “per synthesis” if available. For all reactions in aqueous solution use desalted and highly purified water. The precursor polymer for linear polyethylenimine (LPEI), poly(2-ethyl-2-oxazoline) (Mw 50 kDa) is obtained from Aldrich (Aldrich Cat. No. 37,284-6). Alternatively, LPEI with a molecular weight of 22 kDa can be purchased as ExGen 500 from Fermentas (Burlington, Canada). The heterobifunctional 2 kDa poly(ethylene glycol) (PEG) linker OPSS-PEG-NHS has been synthesized by Rapp Polymere GmbH (Tu¨bingen, Germany); a 3 kDa version is available from IRIS Biotech (Marktredwitz, Germany). Peptides used in this study were synthesized by standard fmoc solid phase synthesis and obtained with more than 95% purity from Biosyntan (Berlin, Germany). Ion exchange resin MacroPrep HighS was purchased from Bio-Rad (Munich, Germany).

2.1 For LPEI Synthesis, the Following Equipment Is Needed

Round bottom flask (NS 29/32, 100 mL). Reflux condenser (NS 29/32, 200–300 mm length). Silicone oil bath. Magnetic stirrer with heating function and stir bar. 50 mL centrifuge tubes Centrifuge system. Lyophilization system.

2.2 For Conjugate Synthesis the Following Lab Equipment Is Needed

Lab-shaker with controllable temperature. Vortex mixer. Glass vial (5–10 mL) equipped with stir bar. Magnetic stirrer. Sterile polypropylene tubes (2 mL and 15 mL). Conjugates are purified on a HPLC/FPLC system running under aqueous conditions. A gradient mixer with at least two channels, a multiline UV/VIS detector, and a fraction collector are needed. pH

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is checked by using a micro pH electrode; for photometric measurements a standard UV/VIS photometer is sufficient.

3

Methods The following assays are used for quantification of LPEI conjugate components.

3.1 Quantification of LPEI (Copper Assay)

This assay has been first described by Ungaro et al. [38]. After adding a solution of Cu2+ to LPEI solutions a dark blue cuprammonium complex is formed, which absorbs strongly at 285 nm enabling quantification of the LPEI content in an aqueous solution. 1. Prepare 100 mL of Cu2+ solution by dissolving 23 mg of CuSO4 in 0.1 M Na-acetate buffer (pH 5.4). 2. For creating a standard curve prepare LPEI standard solutions in water with a final volume of 100 μL at concentrations ranging from 10–100 μg/mL. Use 100 μL water as blank. Standard solutions should always be analyzed in duplicates. 3. Dilute your LPEI sample to be analyzed also to a final volume of 100 μL with water. Use duplicates and different dilutions of sample in water. 4. Mix 100 μL Cu2+ solution with the blank, the standard solutions and the samples and incubate at room temperature for 5 min. 5. Set the absorption wavelength to 285 nm on a standard UV/Vis spectrophotometer and set the absorption of the blank to zero. 6. Measure the absorption of LPEI solutions and LPEI samples to be analyzed and calculate the LPEI concentration with the help of the standard curve (see Note 1).

3.2 Determination of OPSS Content

After addition of excess DTT (Dithiothreitol; 1 M stock in water) to a sample containing an OPSS group, 2-thiopyridone gets released and induces UV absorption at 343 nm (ε343 nm ¼ 8080 M1 cm1). 1. Dilute the OPSS containing sample in 150 μL water, add 15 μL 1 M DTT solution, mix well, and incubate for 10 min. Use 15 μL 1 M DTT diluted with 150 μL water as blank. 2. Transfer to a micro cuvette suitable for UV measurement and measure the absorption at 343 nm against the blank. Calculate the OPSS concentration with ε343 1 cm1 taking into account the dilution factor. nm ¼ 8080 M

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3.3 Quantification of LPEI/PEG Ratio by 1 H-NMR

In addition to the copper and DTT assay, 1H-NMR analysis allows nondestructive determination of the molar ratio between LPEI and PEG in a conjugate. By comparison of the distinct proton signals of ethylene glycol units of PEG and ethyleneimine units of LPEI the yield of the PEGylation reaction can be calculated. 1. Dissolve 5 mg of LPEI-PEG-OPSS (synthesis described in Subheading 3.7) in 1 mL D2O. 2. Adjust the pH of the solution to pH 7 based on pH paper using 1 M stock solutions of DCl or NaOD (see Note 2). 3. Analysis is then carried out on a 200 MHz NMR with 3-(trimethyl silyl) propionic-2,2,3,3-d4 acid (TSP) as internal reference. Alternatively, the residual solvent peak can be used to correct the spectrum. The spectrum is characterized by two major signals, a rather broad LPEI signal at 2.9–3.1 ppm (derived from –CH2–CH2–N) and a sharp PEG signal (derived from –CH2–CH2–O) at 3.7 ppm (see Note 3). The signals of the OPSS moiety can be observed above 7 ppm, but with lower PEGylation rates these signals are normally too weak to allow quantification. To quantify the degree of PEGylation use the integral of the LPEI derived –CH2–CH2–N signal (2.9–3.1 ppm) to normalize the other signals. In the representative example (Fig. 1) the integral of the LPEI derived signal was set to 2047 (N ¼ [22,000(Mw  4 (Protons per monomer))] (LPEI))/43(Mw(ethyleneimine –CH2–CH2–N unit). Calculate the number of protons in the PEG-chain using N ¼ [PEG(Mw(PEG))/44 (Mw(ethylene glycol monomer))]  4 (Protons per –CH2–CH2–O unit). For PEG with a Mw of 2 kDa this yields 182. Divide the integral value of the PEG-signal (3.7 ppm) by 182 to obtain the average number of PEG chains per PEI molecule. In the representative example (Fig. 1) the integral value of the PEG derived signal is 192 which results in a PEG/LPEI ratio of 1.05:1.

3.4 Quantification of Free Thiols (Ellman’s Assay)

This assay is based on the thiol-specific reactivity of Ellman’s reagent (5,50 -dithiobis-(2-nitrobenzoic acid), short DTNB) forming mixed disulfides. During the reaction, 2-nitro-5-thiobenzoate (TNB) is released, which can be quantified by measuring the absorption at 412 nm [39]. 1. Dissolve 2 mg DTNB in 0.1 M HEPES pH 7.4 (see Note 4). 2. Dilute 5 μL of the solution from step 1, fill up to 500 μL with 0.1 M HEPES pH 7.4 and use as a blank on a photometer set to an absorption wavelength of 412 nm. 3. Dilute your sample (using different amounts) in the solution from step 1, incubate for 20 min at ambient temperature and measure the absorption at 412 nm.

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Fig. 1 1H NMR profile of LPEI-PEG-OPSS. OPSS-PEG-NHS (Mw 2 kDa) has been coupled to LPEI (Mw 22 kDa) as described in Subheading 3.7 and the integral of the –CH2–CH2–N signal (LPEI derived) (green line at 2.97 ppm) set to 2047. The number of protons in PEG (Mw 2 kDa) are (2000/44)  4 ¼ 182. The molar ratio PEG/LPEI is calculated by dividing the integral value of the –CH2–CH2–O signal (PEG derived; green line at 3.63 ppm) by the number or protons in PEG: 192/182 ¼ 1.05 PEG2kDa/LPEI22kDa M/M

4. Calculate the thiol content using the molar extinction coefficient ε412nm ¼ 14,100 M1 cm1). 3.5 Peptide Quantification by A280 Measurement

Prior to coupling, the absorption coefficient of the peptide has to be estimated, as the peptide content in conjugates is calculated according to the absorption of aromatic amino acids within the peptide. For this purpose, a useful online tool from the Expasy Bioinformatics tool portal (expasy.org) can be utilized (see http:// web.expasy.org/protparam/). The molar extinction coefficient (ε) is calculated with the following formula: εðPeptideÞ ¼ NumberðTyrÞ  εðTyrÞ þ NumberðTrpÞ  εðTrpÞ þ NumberðCysÞ  εðCysÞ For proteins in water measured at 280 nm use the following extinction coefficients:

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εðTyrÞ ¼ 1, 490 M1 cm1 ; εðTrpÞ ¼ 5, 500 M1 cm1 ; εðCysÞ ¼ 125 M1 cm1 For the GE11 peptide (CYHWYGYTPQNVI; GE11 sequence in bold letters; amino acids taken for calculating extinction coefficient are underlined) the extinction coefficient ε280nm ¼ 9970 M1 cm1 is calculated. Nevertheless, it is of note that the theoretical calculation of ε280nm has to be amended by absorption measurements at 280 nm with the actual peptide used. In the following subheadings, the synthesis of LPEI from the precursor molecule and the conjugate synthesis are described. 3.6 Synthesis of LPEI (Mw 22 kDa) from Poly (2-Ethyl-2-Oxazoline) (Mw 50 kDa) 3.6.1 Synthesis of LPEI–HCl

1. Dissolve 5 g poly(2-ethyl-2-oxazoline) 50 kDa (100 μmol) in 50 mL HCl (30% v/v) in a 100 mL round bottom flask (see Note 5). 2. Attach the round bottom flask to the reflux condenser and connect the reflux condenser to the cooling system. Boil the reaction under reflux and constant stirring for 16 h (see Note 6). 3. Cool the reaction to room temperature. 4. Transfer the content of the round bottom flask including the fine, white precipitate into a 50 mL centrifuge tube. 5. Centrifuge at 4000  g for 5 min and remove the supernatant. 6. Carry out three repeated washes of the precipitate with 40 mL 30% (v/v) HCl per washing cycle as in step 5 (see Note 7). The isolated precipitate is LPEI as HCl salt. 7. Dissolve the precipitate in 200 mL water and lyophilize the product (see Note 8).

3.6.2 Synthesis of LPEI (Free Base)

1. Transfer 2 g LPEI–HCl to a round bottom flask and resuspend it in 30 mL 1 M NaOH. 2. Attach the round bottom flask to the reflux condenser and connect the reflux condenser to the cooling system. Boil the reaction under reflux using an oil bath under constant stirring. Carefully add 1 M NaOH (in 10 mL portions) to the still hot solution until the solution is clear (see Note 5); then switch off the heating. 3. Transfer the content of the round bottom flask including white precipitate into 50 mL centrifuge tubes. 4. Centrifuge at 4000  g for 5 min and remove the supernatant. 5. Carry out three repeated washes of the precipitate with 40 mL 1 M NaOH (per tube) per washing cycle as in step 5. Conduct 5 further washing cycles with 40 mL water (per tube).

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6. Resuspend the precipitate in 50 mL water and transfer into a 100 mL round bottom flask. 7. Lyophilize product. 8. The resulting LPEI can be stored at room temperature in a desiccator protected from light. 3.7 Synthesis and Purification of LPEI-PEG-OPSS

1. Dissolve 75 mg (3.4 μmol) LPEI (free base; Mw 22,000) in 1.5 mL absolute ethanol in a 2 mL polypropylene reaction vial with lid and incubate on a standard lab mixer for 15 min at 800 rpm and 35  C (see Note 9). 2. Dissolve OPSS-PEG-NHS linker (6.8 μmol; 2 eq based on LPEI) in 100 μL DMSO (water free), add to the reaction mixture from step 1 and incubate again for 3 h at 35  C with 800 rpm. 3. Quench the reaction by adding 100 μL 1 M Tris (pH 8.0) and incubating for 15 min at 35  C with 800 rpm. 4. Transfer the reaction mixture to a 15 mL polypropylene tube and add approximately 2 mL of a 20 mM HEPES solution pH 7.4 (pH 7.4) and 20 mM HEPES–3 M NaCl (pH 7.4), so that in the final volume of 5 mL the NaCl concentration reaches 0.5 M (the starting NaCl concentration during ion exchange purification, see below) (see Notes 10 and 11). 5. Adjust the pH with 1 M HCl and check the pH with a micro pH probe until pH 7 is reached. Thereafter fill up the reaction mix to 5 mL with 20 mM HEPES (pH 7.4) (see Notes 12 and 13). 6. Equip the HPLC system with a column (HR10/10, i.e., 10 cm in length, 10 mm diameter) filled with cation exchange resin (MacroPrep High S). Set the UV detector to 240, 280 and 343 nm. Equilibrate the system with 83.3% solution A (20 mM HEPES; pH 7.4) and 16.7% solution B (20 mM HEPES–3 M NaCl; pH 7.4) for at least 1 h at a flow rate of 0.5 mL/min; this corresponds to a concentration of 500 mM NaCl and 20 mM HEPES at pH 7.4. 7. Program a gradient (flow rate: 0.5 mL/min) with 16.7% A and 83.3% B from 0 to 25 min, and linear change to 100% B over 40 min (25–65 min), followed by 100% B for 20 min. 8. Load the product from step 4 onto the column and run the gradient. Fractions eluting during the first 25 min (at 500 mM NaCl, 20 mM HEPES pH 7.4) contain unreacted OPSS-PEGNHS and by-products from the reaction between the NHS ester in PEG and amines in LPEI. A representative chromatogram is shown in Fig. 2. Fractions eluting between 2.0 and 2.8 M NaCl contain LPEI modified with PEG (LPEI-PEG-

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Fig. 2 Cation exchange chromatography profile of LPEI-PEG-OPSS. 5 mL reaction mixture from step 5, Subheading 3.7, were loaded onto a HR10/10 cation exchange column and purified as described in the text. Fraction eluting from 40 to 60 min were pooled and further processed as described

OPSS), these fractions are pooled and subsequently dialyzed against 5 L water at 4  C under constant stirring overnight. 9. Lyophilize product (see Note 14). 3.8 Coupling of Peptide to LPEIPEG-OPSS

The peptide CYHWYGYTPQNVI (sequence of GE11 in bold letters, see ref. 36) used in this study was synthesized by standard Fmoc solid phase peptide synthesis and purified on a C18 reversed phase HPLC column. The product was eluted with an acetonitrile gradient (A: 0.05% (v/v) TFA in water, B: 0.05% TFA (v/v) in 80% acetonitrile in water), 0.6 mL/min flow, linear gradient 2.5% B/min, detection at 220 nm) the peptide eluted at 13.1 min) and thereafter lyophilized (see Note 15). 1. For this reaction always use 2 equivalents of peptide based on OPSS in the sample. 2. The whole reaction can be conducted in a glass vial equipped with a magnetic stir bar. 3. Purge all solvents used for this synthesis (30% acetonitrile–water–0.1% TFA; 20 mM HEPES–30% acetonitrile [pH 7.4]; 20 mM HEPES–3 M NaCl–10% (v/v) acetonitrile [pH 7.4]) with argon. 4. Dissolve the peptide in 100 μL 30% acetonitrile–water–0.1% TFA. (a) To ensure that the thiol residue has not been oxidized or dimerized, quantify the free thiol content by Ellman’s assay. For this purpose, dilute 1 μL peptide sample 1:100

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Fig. 3 Cation exchange chromatography profile of LPEI-PEG-GE11. 10 mL reaction mixture from step 11, Subheading 3.8, were loaded onto a HR10/10 cation exchange column and purified as described in the text. Fraction eluting from 57 to 77 min were pooled and further processed as described

with water and use 15 μL dilution for the assay, proceed as described in Subheading 3.4. 5. Dissolve LPEI-PEG-OPSS in 20 mM HEPES–30% acetonitrile (pH 7.4) (see Note 16). (a) The final concentration of LPEI in the reaction mix should be between 4 and 5 mg/mL. 6. For online reaction monitoring, check the absorption of the LPEI-PEG-OPSS solution at 343 nm prior to mixing. 7. Add the peptide solution (2 eq in 100 μL, see above) to the LPEI-PEG-OPSS solution and incubate at room temperature under constant stirring. 8. Thereafter (approx. 1 min) transfer a 150 μL sample from the reaction mix into a cuvette and measure A343 with a UV/Vis spectrophotometer. From then on A343 is measured every 30 min, until there is no more increase in absorption observed. 9. Calculate the 2-thiopyridone released as described in Subheading 3.2. 10. Add 20 mM HEPES–3 M NaCl–10% (v/v) acetonitrile (pH 7.4) to obtain a final concentration of 500 mM NaCl in a final volume of 5 mL. 11. Fill up to 5 mL with 20 mM HEPES–10% (v/v) acetonitrile (pH 7.4) (see Note 13).

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12. Equip a HPLC/FPLC system with a 10/10 MacroPrep column (MacroPrep High S; HR10/10; Bio-Rad, Mu¨nchen, Germany). Set the UV detector to 240, 280, and 343 nm. Equilibrate the system with 83.3% solution A (20 mM HEPES–10% (v/v) acetonitrile; pH 7.4) and 16.7% solution B (20 mM HEPES–3 M NaCl–10% (v/v) acetonitrile; pH 7.4) for at least 1 h at a flow rate of 0.5 mL/min; this corresponds to a concentration of 500 mM NaCl. 13. Program a gradient (flow rate: 0.5 mL/min) with 16.7% A and 83.3% B from 0 to 35 min, and linear change to 100% B over 40 min followed by 100% B for 20 min. 14. Load the product from step 11 onto the column and run the gradient. The product elutes between 2.0 and 2.8 M NaCl (Fig. 2). A chromatogram from this purification step is depicted in Fig. 3. 15. Pool the fractions eluting between 2 and 2.8 M NaCl and dialyze them against 5 L water at 4  C under constant stirring overnight. 16. Lyophilize product. 17. At this stage the product can be stored under dry conditions at 80  C. 18. For analyzing LPEI and peptide content reconstitute conjugate in water and set pH to 7. 19. Analyze the LPEI content in the conjugate by copper assay. 20. Calculate the peptide concentration as described in Subheading 3.5. For the measurement, take a 150 μL aliquot of your sample (undiluted, can be reused, so use sterile, clean cuvettes) and measure the absorption at 280 nm. 21. For further storage of conjugate containing solutions freeze aliquots (snap freezing) and store at 80  C. 3.9 Polyplex Formation

The LPEI-PEG-peptide conjugates have been developed for local or systemic delivery of nucleic acids in vivo. For this purpose, polyplexes are generated in a low salt buffer (HBG: HEPES buffered glucose, 20 mM HEPES–5% glucose w/v [pH 7.4]). This allows the formation of rather small, colloidal stable polyplexes. The HBG buffer is sterile filtered (0.2 μm pore size) and stored in aliquots either at 4  C or frozen at 20  C. Plasmid DNA is usually produced in suitable E. coli strains and the plasmid isolated after alkaline lysis of bacterial cells using commercially available purification kits. Please make sure to use kits which result in a low endotoxin contamination in the purified plasmid. Alternatively, plasmids can be produced by companies specialized in plasmid production (see Note 17).

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LPEI/pDNA polyplexes are defined by their molar ratio of phosphate in plasmid and nitrogen in LPEI (N/P ratio). The N/P ratio is transformed into a w/w ratio by the following formula: μg PEI ¼ μg DNA  43  ðN=P ratioÞ=330: Forty-three is the molecular weight of one repeating [CH2–CH2–NH] unit in PEI (N), and 330 is the average molecular weight of one nucleotide (P). Polyplexes with N/P ratios ranging from 5 to 10 are recommended. Here, an example is given to obtain 500 μL polyplexes containing 100 μg plasmid DNA at an N/P ratio of 6. 1. Prepare 250 μL plasmid diluted in HBG (final plasmid concentration 400 μg/mL). 2. Prepare 250 μL LPEI-PEG-peptide conjugate solution in HBG (amount of LPEI in 250 μL total volume: 100  43  6/330 ¼ 78.18 μg). 3. Transfer the LPEI-PEG-peptide dilution to the plasmid dilution and immediately pipet the solution 10 times up and down (see Note 18). 4. For quality control, measure polyplex size and zeta potential by dynamic light scattering (DLS) or nanoparticle tracking analysis (NTA). The average particle size should be below 300 nm. 5. Store polyplex containing solution at ambient temperature for no longer than 20 min prior to use.

4

Notes 1. The absorption value of your sample should be between 0.1 and 1.2 to ensure linear correlation between absorption and LPEI concentration. 2. Adjusting the pH to 7.0 with deuterated hydrochloric acid (DCl) or deuterated sodium hydroxide (NaOD) is important at this step to obtain comparable results, as acidic pH can cause a shift of the LPEI peak towards 3.5 ppm. 3. One has to be aware of the possibility of additional peaks due to incomplete deprotection of the LPEI-precursor, especially signals of propionamidyl residues (1.2–1.3 ppm; 2.1–2.3 ppm; 3.5–3.8 ppm) can lead to a overestimation of PEGylation [40]. When dissolved in D2O secondary amines of LPEIPEG-OPSS do not show a signal in 1H NMR. 4. Always prepare the Ellman’s reagent fresh, do not exceed storage for longer than 1 day.

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5. Take safety measures (protective goggles, gloves, lab coat) when handling highly concentrated acids or alkaline. 6. Carry out the reaction in a fume hood and take proper safety measures when carrying out the reaction overnight. The reaction mixture appears first clear, after approximately 5 h a white precipitate is formed (LPEI–HCl). 7. Washings steps have to be repeated until the supernatant is clear and odorless. The residual propionic acid can develop a strong odor. 8. LPEI–HCl is well soluble in water at pH >2, but remains insoluble in >25% (v/v) HCl. At this step, LPEI–HCl can be neutralized with concentrated NaOH to pH 7 and thereafter used as transfection reagent. To calculate the LPEI, content, please note that LPEI–HCl has a molecular weight of 79.6 Da per repeating unit ([CH2–CH2–NH].HCl). 9. LPEI dissolves fast, but constant mixing is necessary to achieve a homogeneous solution, which is highly viscous. 10. When adding in 10 μL aliquots HCl in, do this under constant vortexing to avoid local low pH, which could result in LPEI precipitation. Cave: the solutions heats up if HCl is added too fast. 11. All solutions used for diluting the sample have to be purged with argon or nitrogen for at least 15 min prior to use. 12. This is necessary to reduce the EtOH concentration prior to loading of the mix onto the ion exchange columns, where otherwise a too high EtOH concentration results in pressure increase and compression of the column material. 13. Before loading the sample on the HPLC/FPLC the sample should always be filtered at least through a 0.45 μm filter. 14. At this step, the resulting LPEI-PEG-OPSS conjugate can be stored at 80  C until further use. 15. Peptides containing free thiols, like the terminal Cys residue, should be stored under argon at 80  C for no longer than a month, depending on the peptide. After extended storage, dimerization or oxidation of thiol groups occurs. 16. Make sure that the final reaction mixture contains at least a concentration of acetonitrile needed for dissolving the peptide, otherwise the peptide can precipitate. In case of CYHWYGYTPQNVI (GE11 sequence in bold letters) it is 30% (v/v) acetonitrile. 17. Elevated levels of high molecular weight bacterial genomic DNA can be present in the plasmid preparation using commercialized isolation kits [41], especially when using for example low copy plasmid [42]. In such cases, preparations methods

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using additional purification steps are recommended [41]. For example, the company PlasmidFactory (www.plasmidfactory. com) offers plasmid in “ccc” grade, that is, supercoiled plasmid structure and absence of bacterial genomic DNA impurities. Besides biological effects, the presence of high molecular weight DNA impurities can lead to excessive particle aggregation during the mixing process. For the generation of welldefined larger batches of polyplexes, methods using controllable mixing devices are recommended (see refs. 18, 43 and Chapter 14 in this book). 18. Mixing should be done immediately, to avoid formation of aggregates; the appearance of aggregates correlates positively with an increase in the final concentration of polyplexes and the salt concentration in the dilution buffer; there is a negative correlation with the increase in the N/P ratio, see also ref. 44).

Acknowledgments This work was supported by the Center for Nanoscience (CeNS) and the German Research Foundation (SFB824) to M.O., and the Nanosystems Initiative Munich (NIM) to E.W. References 1. Gehrig S, Sami H, Ogris M (2014) Gene therapy and imaging in preclinical and clinical oncology: recent developments in therapy and theranostics. Ther Deliv 12:1275–1296 2. El-Aneed A (2004) An overview of current delivery systems in cancer gene therapy. J Control Release 94:1–14 3. Pack DW, Hoffman AS, Pun S, Stayton PS (2005) Design and development of polymers for gene delivery. Nat Rev Drug Discov 4:581–593 4. Boussif O, Lezoualc’h F, Zanta MA, Mergny MD, Scherman D, Demeneix B, Behr JP (1995) A versatile vector for gene and oligonucleotide transfer into cells in culture and in vivo: polyethylenimine. Proc Natl Acad Sci U S A 92:7297–7301 5. Kircheis R, Wightman L, Wagner E (2001) Design and gene delivery activity of modified polyethylenimines. Adv Drug Deliv Rev 53:341–358 6. Tang MX, Szoka FC (1997) The influence of polymer structure on the interactions of cationic polymers with DNA and morphology of the resulting complexes. Gene Ther 4:823–832

7. Kopatz I, Remy JS, Behr JP (2004) A model for non-viral gene delivery: through syndecan adhesion molecules and powered by actin. J Gene Med 6:769–776 8. Sonawane ND, Szoka FC Jr, Verkman AS (2003) Chloride accumulation and swelling in endosomes enhances DNA transfer by polyamine-DNA polyplexes. J Biol Chem 278:44826–44831 9. Nguyen J, Szoka FC (2012) Nucleic acid delivery: the missing pieces of the puzzle? Acc Chem Res 45:1153–1162 10. Schaffert D, Ogris M (2013) Nucleic acid carrier systems based on polyethylenimine conjugates for the treatment of metastatic tumors. Curr Med Chem 20:3456–3470 11. Ferrari S, Moro E, Pettenazzo A, Behr JP, Zacchello F, Scarpa M (1997) ExGen 500 is an efficient vector for gene delivery to lung epithelial cells in vitro and in vivo. Gene Ther 4:1100–1106 12. Goula D, Benoist C, Mantero S, Merlo G, Levi G, Demeneix BA (1998) Polyethylenimine-based intravenous delivery of transgenes to mouse lung. Gene Ther 5:1291–1295

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13. Wightman L, Kircheis R, Rossler V, Carotta S, Ruzicka R, Kursa M, Wagner E (2001) Different behavior of branched and linear polyethylenimine for gene delivery in vitro and in vivo. J Gene Med 3:362–372 14. Brissault B, Kichler A, Guis C, Leborgne C, Danos O, Cheradame H (2003) Synthesis of linear polyethylenimine derivatives for DNA transfection. Bio Conjug Chem 14:581–587 15. Ogris M, Wagner E (2012) Synthesis of linear polyethylenimine and use in transfection. Cold Spring Harb Protoc 2012:246–250 16. Thomas M, Lu JJ, Ge Q, Zhang C, Chen J, Klibanov AM (2005) Full deacylation of polyethylenimine dramatically boosts its gene delivery efficiency and specificity to mouse lung. Proc Natl Acad Sci U S A 102:5679–5684 17. Boeckle S, von Gersdorff K, van der Piepen S, Culmsee C, Wagner E, Ogris M (2004) Purification of polyethylenimine polyplexes highlights the role of free polycations in gene transfer. J Gene Med 6:1102–1111 18. Taschauer A, Geyer A, Gehrig S, Maier J, Sami H, Ogris M (2016) Up-scaled synthesis and characterization of nonviral gene delivery particles for transient in vitro and in vivo transgene expression. Hum Gene Ther Methods 27:87–97 19. Goula D, Becker N, Lemkine GF, Normandie P, Rodrigues J, Mantero S, Levi G, Demeneix BA (2000) Rapid crossing of the pulmonary endothelial barrier by polyethylenimine/DNA complexes 965. Gene Ther 7:499–504 20. Chollet P, Favrot MC, Hurbin A, Coll JL (2002) Side-effects of a systemic injection of linear polyethylenimine-DNA complexes. J Gene Med 4:84–91 21. Kwok A, Hart SL (2011) Comparative structural and functional studies of nanoparticle formulations for DNA and siRNA delivery. Nanomedicine 7:210–219 22. Itaka K, Harada A, Yamasaki Y, Nakamura K, Kawaguchi H, Kataoka K (2004) In situ single cell observation by fluorescence resonance energy transfer reveals fast intra-cytoplasmic delivery and easy release of plasmid DNA complexed with linear polyethylenimine. J Gene Med 6:76–84 23. de Bruin KG, Fella C, Ogris M, Wagner E, Ruthardt N, Brauchle C (2008) Dynamics of photoinduced endosomal release of polyplexes. J Control Release 130:175–182 24. Zintchenko A, Susha AS, Concia M, Feldmann J, Wagner E, Rogach AL, Ogris M (2009) Drug nanocarriers labeled with nearinfrared-emitting quantum dots

(quantoplexes): imaging fast dynamics of distribution in living animals. Mol Ther 17:1849–1856 25. Kursa M, Walker GF, Roessler V, Ogris M, Roedl W, Kircheis R, Wagner E (2003) Novel shielded transferrin-polyethylene glycol-polyethylenimine/DNA complexes for systemic tumor-targeted gene transfer. Bioconjug Chem 14:222–231 26. Fitzsimmons REB, Uludag H (2012) Specific effects of PEGylation on gene delivery efficacy of polyethylenimine: Interplay between PEG substitution and N/P ratio. Acta Biomater 8:3941–3955 27. Smrekar B, Wightman L, Wolschek MF, Lichtenberger C, Ruzicka R, Ogris M, Rodl W, Kursa M, Wagner E, Kircheis R (2003) Tissue-dependent factors affect gene delivery to tumors in vivo. Gene Ther 10:1079–1088 28. Ogris M, Brunner S, Schuller S, Kircheis R, Wagner E (1999) PEGylated DNA/transferrin-PEI complexes: reduced interaction with blood components, extended circulation in blood and potential for systemic gene delivery. Gene Ther 6:595–605 29. Schwerdt A, Zintchenko A, Concia M, Roesen N, Fisher K, Lindner LH, Issels R, Wagner E, Ogris M (2008) Hyperthermiainduced targeting of thermosensitive gene carriers to tumors. Hum Gene Ther 19:1283–1292 30. Fella C, Walker GF, Ogris M, Wagner E (2008) Amine-reactive pyridylhydrazone-based PEG reagents for pH-reversible PEI polyplex shielding. Eur J Pharm Sci 34:309–320 31. Walker GF, Fella C, Pelisek J, Fahrmeir J, Boeckle S, Ogris M, Wagner E (2005) Toward synthetic viruses: endosomal pH-triggered deshielding of targeted polyplexes greatly enhances gene transfer in vitro and in vivo. Mol Ther 11:418–425 32. Klutz K, Schaffert D, Willhauck MJ, Grunwald GK, Haase R, Wunderlich N, Zach C, Gildehaus FJ, Senekowitsch-Schmidtke R, Goke B, Wagner E, Ogris M, Spitzweg C (2011) Epidermal growth factor receptor-targeted (131) I-therapy of liver cancer following systemic delivery of the sodium iodide symporter gene. Mol Ther 19:676–685 33. Kircheis R, Wightman L, Schreiber A, Robitza B, Rossler V, Kursa M, Wagner E (2001) Polyethylenimine/DNA complexes shielded by transferrin target gene expression to tumors after systemic application. Gene Ther 8:28–40

Synthesis of Polyethylenimine-Based Nanocarriers for Systemic Tumor. . . 34. Ogris M, Steinlein P, Carotta S, Brunner S, Wagner E (2001) DNA/polyethylenimine transfection particles: Influence of ligands, polymer size, and PEGylation on internalization and gene expression. AAPS Pharm Sci 3: E21 35. Ogris M, Walker G, Blessing T, Kircheis R, Wolschek M, Wagner E (2003) Tumortargeted gene therapy: strategies for the preparation of ligand-polyethylene glycol-polyethylenimine/DNA complexes. J Control Release 91:173–181 36. Schafer A, Pahnke A, Schaffert D, van Weerden WM, de Ridder CM, Rodl W, Vetter A, Spitzweg C, Kraaij R, Wagner E, Ogris M (2011) Disconnecting the Yin and Yang relation of epidermal growth factor receptor (EGFR)-mediated delivery: a fully synthetic, EGFR-targeted gene transfer system avoiding receptor activation. Hum Gene Ther 22:1463–1473 37. Sidi AA, Ohana P, Benjamin S, Shalev M, Ransom JH, Lamm D, Hochberg A, Leibovitch I (2008) Phase I/II marker lesion study of intravesical BC-819 DNA plasmid in H19 over expressing superficial bladder cancer refractory to bacillus calmette-guerin. J Urol 180:2379–2383 38. Ungaro F, De Rosa G, Miro A, Quaglia F (2003) Spectrophotometric determination of polyethylenimine in the presence of an oligonucleotide for the characterization of

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controlled release formulations. J Pharm Biomed Anal 31:143–149 39. Eyer P, Worek F, Kiderlen D, Sinko G, Stuglin A, Simeon-Rudolf V, Reiner E (2003) Molar absorption coefficients for the reduced Ellman reagent: reassessment. Anal Biochem 312:224–227 40. Jeong JH, Song SH, Lim DW, Lee H, Park TG (2001) DNA transfection using linear poly (ethylenimine) prepared by controlled acid hydrolysis of poly(2-ethyl-2-oxazoline). J Control Release 73:391–399 41. Schleef M, Schmidt T (2004) Animal-free production of ccc-supercoiled plasmids for research and clinical applications. J Gene Med 6(Suppl 1):S45–S53 42. Magnusson T, Haase R, Schleef M, Wagner E, Ogris M (2011) Sustained, high transgene expression in liver with plasmid vectors using optimized promoter-enhancer combinations. J Gene Med 13:382–391 43. Kasper JC, Schaffert D, Ogris M, Wagner E, Friess W (2011) The establishment of an up-scaled micro-mixer method allows the standardized and reproducible preparation of welldefined plasmid/LPEI polyplexes. Eur J Pharm Biopharm 77:182–185 44. Ogris M, Steinlein P, Kursa M, Mechtler K, Kircheis R, Wagner E (1998) The size of DNA/transferrin-PEI complexes is an important factor for gene expression in cultured cells. Gene Ther 5:1425–1433

Chapter 7 Cationic Photopolymerized Polydiacetylenic (PDA) Micelles for siRNA Delivery Manon Ripoll, Patrick Neuberg, Jean-Serge Remy, and Antoine Kichler Abstract Polymerized micelles obtained by photopolymerization of diacetylenic surfactants and which are forming polydiacetylenic systems (PDAs) have recently gained interest as stabilized monodisperse systems showing potential for the delivery of hydrophobic drugs as well as of larger biomolecules such as nucleic acids. Introduction of pH-sensitive histidine groups at the surface of the micellar PDA systems allows for efficient delivery of siRNA resulting in specific gene silencing through RNA interference. Here, we describe the detailed experimental procedure for the reproducible preparation of these photopolymerized PDA micelles. We provide physicochemical characterization of these nanomaterials by dynamic light scattering, transmission electron microscopy, and diffusion ordered spectroscopy. Moreover, we describe standardized biological tests to evaluate the silencing efficiency by the use of a cell line constitutively expressing the luciferase reporter gene. Key words RNA interference, siRNA, Nonviral delivery system, Cationic micelles, Endosomal release, Histidine residue, Photopolymerization, Diacetylenic micelles, Polydiacetylenic, PDA, Photopolymerization

1

Introduction Gene therapy is one of the major biomedical breakthroughs of the last decades and of great promise for an increasing number of patients affected by genetic or acquired diseases. Delivery of a “medicine” based on nucleic acids into cells requires the use of carriers (vectors), which will allow the therapeutic agent to cross various biological barriers and reach the target cells where it will be processed. Vectors for nucleic acids are either viral or synthetic (nonviral vectors). Nonviral vectors such as cationic lipids and polymers generally present interesting in vitro activities but remain less efficient in vivo as compared to the most efficient viral vectors. Nevertheless, synthetic vectors continue to be the focus of intense research efforts in order to improve their efficiency.

Manfred Ogris and Haider Sami (eds.), Nanotechnology for Nucleic Acid Delivery: Methods and Protocols, Methods in Molecular Biology, vol. 1943, https://doi.org/10.1007/978-1-4939-9092-4_7, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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To increase the transfection efficiency of nonviral vectors, it is for example important to have a better control of the size of the complexes. While large aggregates have the propensity to sediment in cell culture experiments onto the cell layer improving particle internalization through close contact with the cell membranes, they are not suited for in vivo delivery. What stands out as an advantage for in vitro experiments is considered as a drawback for most in vivo applications. Indeed, large particles (>100 nm) are unable to diffuse through tissues and, in particular, through the walls of blood vessels. Detergents forming micellar systems are mostly nanostructured and monodisperse systems. The spherical form of the micelle is given by the packing characteristics and polar head repulsions of the charged detergent molecules. Moreover, some cationic micelle forming detergents such as CTAP (cetyl trimethylammonium bromide) are able to interact with DNA by electrostatic interaction with the negatively charged phosphate groups. Cooperative hydrophobic assembly of the lipidic chains, is leading to local micellar structures which will condense DNA molecules efficiently [1]. Nevertheless, in biological assays rapid diffusion of the detergent molecules getting trapped in the biological membranes leads to the opening of the micelle-like domains and release of the DNA cargo molecules before their internalization [2]. Monodisperse spherical particles have also been obtained by dimerizable surfactant molecules containing cysteinyl groups together with a cationic guanidine group (guanidinocysteine-Ndecylamide) [3] or an ornithyl group (ornithyl-cysteinyl-tetradecylamide) [4]. These molecules, with their elegant concept of DNA compaction, proved relatively efficient for DNA transfection in vitro [5]. However, no in vivo experiments were published that could demonstrate the functionality of these structures in a living organism. In order to improve the stability of small sized delivery vectors we describe here a new system that shares common features with cationic detergent molecules and cationic polymers bridging the gap between “destructured” polymers and “organized” labile micelles. Indeed, we have explored delivery systems where the vectors are based on auto-organized detergent type molecules that are polymerized inside their lipidic core. Diacetylenic surfactants auto-organize into micelles that can be photopolymerized by strong UV irradiation, so that their lipidic chains become covalently interconnected (Fig. 1). These cross-linked systems gain new biological properties and allow to fine tune their interaction with biological membranes, which present one of the crucial steps in the transfection process. Prior work by our group has allowed to prove that such polydiacetylenic systems (PDA micelles) can be used for DNA delivery. Their delivery potential compared favorably with classic cationic

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Fig. 1 Illustration of the stepwise assembly of the micelles/siRNA complexes

amphiphilic molecules [6]. More recent research by our group showed that fine tuning of the polar headgroups of diacetylenic surfactants allows to create photopolymerized systems which can interact with biological membranes, and release model hydrophobic fluorescent compounds into the cytoplasm [7]. In 2016 we proved that the modification of the hydrophilic headgroups by the introduction of pH-sensitive groups such as imidazole groups from histidine residues could promote siRNA delivery, leading to specific RNA interference [8]. We evidenced that the release of siRNA is coupled with the ATPase proton pump activity located in the endosomal compartments. The efficient release after endocytosis proceed through the pH buffering capacity by the delivery system bearing imidazole groups that become protonated, counteracting this way the acidification steps of endosomal maturation (Fig. 2). The H+ buffering capacity at the pH values of early endosomes is indeed crucial for avoiding degradation of the siRNA molecules prior to cytoplasmic release. These finding corroborate the often cited “proton sponge” hypothesis, which has been termed for the delivery mechanism of one of the most efficient DNA delivering polymers, namely polyethylenimine (linear and branched PEIs) [9–11]. Meanwhile PDA-based micellar systems tend to become more and more promising systems for the delivery of all kind of drugs in in vivo and preclinical applications, and have been explored by different groups [12–15]. Polydiacetylene micelles decorated with a targeting peptide were able to encapsulate the anticancer drug camptothecin (CPT). These CPT-PDA micelles could kill ovarian cancer after internalization and they penetrated efficiently into a tumor [15]. Also, an elegant strategy has been developed to covalently attach the cytotoxic peptide magainin II (MGN-II) to the surface of PDA micelles. These well-defined PDA micelles had high cytotoxicity in cancer cell lines, and were able to reduce the tumor size in mice [13]. Lastly, PDA micelles with “stealth” zwitterionic

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Fig. 2 Mechanism of endosomal escape mediated by histidine-based micelles

surface coating were tested in a murine xenograft model of breast cancer. The micelles were taken up at the margins of the tumor, and allowed to delineate its volume with the aim to aid surgery [14]. Notably, it is of crucial importance to define reproducible formulation methods of these nanosized PDA micellar systems. As even small variations in the formation of these nanoparticles can entail large variations in the biological activity, it is important to propose a detailed experimental procedure of their synthesis, formulation and characterization. Here, we provide guidelines for the preparation of our histidine-based PDA-micellar systems. In particular we will describe: (1) the protocol of synthesis and the chemical characterization of the diacetylenic surfactant presenting the histidine headgroup; (2) the method of preparation of the micelles and the characterization of these nano-objects (by measuring the CMC, by using DOSY-NMR, DLS and by performing gel shift mobility assays); (3) the in vitro assays used to determine the siRNA transfection efficiency (by measuring the silencing of a reporter gene); (4) the evaluation of the cytotoxicity of the micelles–siRNA complexes by dosing the protein content and by performing MTT assays.

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Materials

2.1 Synthesis and Characterization of the Surfactant

1. Analytical grade of dichloromethane (DCM), tetrahydrofuran, methanol, ethanol, ethyl acetate, and cyclohexane are used (Sigma-Aldrich).

2.1.1 Chemicals

2. 10,12-pentacosediynoic acid (Sigma-Aldrich) (see Note 1). 3. N-hydroxysuccinimide (NHS), N,N-diisopropylethylamine (anhydrous DIPEA), 1-ethyl-(3-dimethylaminopropyl)carbodiimide hydrochloride (EDC), trifluoroacetic acid (TFA) and 4,7,10-trioxa-1,13-tridecanediamine (Sigma-Aldrich). 4. Nα-(tert-butoxycarbonyl)-L-histidine stored at +4
C.

2.1.2 Chromatographic Methods

(Sigma-Aldrich)

is

1. Analytical thin layer chromatography (TLC) is performed using 60F-254 silica gel plates (Merck). 2. Solution of acidic vanillin reagent: 15 g vanillin in 250 mL ethanol + 2.5 mL conc. sulfuric acid. 3. Ninhydrin solution: 1.5 g ninhydrin in 100 mL ethanol + 3 mL conc. acetic acid. 4. Purifications by column chromatography are performed using silica gel Geduran Si 60 (Merck, 0.040–0.063 mm).

2.1.3 Nuclear Magnetic Resonance Spectroscopy

1. 1H and 13C NMR spectra were recorded at room temperature with the following spectrometer: Bruker Advance 400 (NMR 1 H: 400 MHz and NMR 13C: 75 MHz). 2. Chloroform-d (CDCl3) and methanol-d4 (MeOD).

2.1.4 High-Resolution Mass Spectra

2.2 Synthesis and Characterization of the Micelles 2.2.1 Formulation

High-resolution mass spectra (HRMS) are obtained using an Agilent Q-TOF (time of flight) 6520 and liquid chromatography coupled broad-resolution mass spectra (LC-BRMS) are realized using an Agilent MSD 1200 SL (EPI/APCI) with an Agilent HPLC 1200 SL. 1. Sonication steps are performed using an ultrasonic bath sonicator (Fisher Scientific FB15047). 2. Polymerization is done in 1 mL quartz cuvettes (Hellma) using a Cross-Linker Bio-Link 254 (Fisher Bioblock). 3. Dialysis cassettes (2000 MWCO, 0.5–3.0 mL capacity, Thermo Scientific).

2.2.2 Critical Micelle Concentration (CMC)

1. Fluorescence is quantified on a RF-5301 PC spectrofluorophotometer (Shimadzu). 2. DMSO and pyrene (Sigma-Aldrich).

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3. 10 mM stock solution of micelles: prepared as described in Subheading 3.2 at the exception that the film is dissolved in 700 μL rather than 1 mL of fresh deionized water (see step 4). 4. 1 mM solution of pyrene: 20.8 mg of pyrene is dissolved in 1 mL DMSO and sonicated. This solution was then diluted 100 times to obtain the 1 mM pyrene solution. 2.2.3 DOSY-NMR Experiments

2.3 Characterization of the Micelles/siRNA Complexes 2.3.1 Gel Mobility Shift Assay

1D-1H and 2D-DOSY experiments are carried out at 300 K on a 600 MHz Bruker Avance III NMR spectrometer with 5 mm DOTY probe delivering up to 500 G/cm gradients. 1. Ultrapure agarose. 2. 10 Tris–acetate for 1 L solution: 400 mM Tris base (48.4 g), concentrated solution of acetic acid (about 13 mL) until pH ¼ 7.5; diluted prior to electrophoresis to 40 mM Trisacetate. 3. Loading buffer: 30% glycerol in 40 mM Tris–acetate, pH ¼ 7.5. 4. SYBRSafe DNA gel stain (Invitrogen). 5. Electrophoresis unit.

2.3.2 Dynamic Light Scattering

1. Dynamic light scattering (DLS) measurements were performed on a Zetasizer Nano-ZS instrument (Malvern Instruments). 2. Disposable cuvettes: UV-Cuvette (12.5  12.5  45 mm, 70 μL, Brand).

micro

3. Suspension of 60 nm latex nanospheres (Duke Scientific) for calibration of the DLS instrument. 2.4 In Vitro Evaluation 2.4.1 Cell Culture

1. The A549-luc cells that stably express the reporter gene luciferase GL3 (1–2  1010 RLU/mg protein) were established by transfecting human lung carcinoma A549 cells (CCL-185; ATCC) with pGL3-Luc plasmid (Clontech) using jetPEI as delivery system (Polyplus-Transfection). 2. RPMI 1640 medium and fetal bovine serum (FBS). 3. Penicillin–streptomycin. 4. Antibiotic G418. 5. 24-well culture plates. 6. Complete medium: RPMI 1640 medium supplemented with 10% FBS and 1% antibiotic solution.

2.4.2 siRNA Transfection

1. HBG: 10 mM Hepes (119.5 mg/50 mL), 10 mM Hepes-Na (130.2 mg/50 mL), 5% glucose (2.5 g/50 mL), pH ¼ 7.5, sterile filtered on 0.22 μm CA filter units (Millex, Millipore).

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2. siRNA (Eurogentec), provided in annealed form and HPLC purified. Sequences are as follows with: TT (DNA bases). Luciferase targeting siRNA (siLuc): Gl3-ssSIRNA: 50 -CUU ACG CUG AGU ACU UCG A TT-30 (sense strand). Gl3-asSIRNA: 50 -U CGA AGU ACU CAG CGU AAG TT-30 (antisense strand). Control siRNA (siCTL): Ctl-ssSIRNA: 50 -CGU ACG CGG AAU ACU UCG ATT-30 (sense strand). Ctl-asSIRNA: 50 -U CGA AGU AUU CCG CGU ACG TT-30 (antisense strand). 3. 5 μM solution of siLuc or siCTL: dilute the stock solution of 100 μM siRNA to have a final work solution of 5 μM in RNasefree water. 4. Transfection reagent INTERFERin (from POLYPLUStransfection SA, Ozyme). 2.4.3 Luciferase Assay

1. Luciferase kit (Promega). 2. Centro LB Luminometer (from Berthold). 3. White 96-well plates. 4. 1 lysis buffer: dilute five times 5 cell culture lysis buffer (Promega) in water.

2.4.4 Protein Assay

1. BCA assay kit (Interchim). 2. 3.5 mL tubes (PS, Ø 11.8, Gosselin). 3. Semi-micro cuvette for spectrophotometer in crystal PS (Ratiolab). 4. Absorption spectra were recorded using Shimadzu UV-1800 UV/visible spectrophotometer (mode: simple reads).

2.4.5 Cell Viability Assay

1. 1 mg/mL of 3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyl-tetrazolium bromide (MTT) in serum-free culture media. 2. 96-well culture plates. 3. Quantification was done by spectrophotometry on a Multiskan FC plate reader (Thermo Fisher).

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Methods

3.1 Synthesis of C25diynetrioxa-LHistidine 4 (Fig. 3; Note 2) 3.1.1 2,5-Dioxopyrrolidin1-yl Pentacosa-10,12Diynoate (1)

1. Dissolve 10,12-pentacosadiynoic acid (1 eq, 10 g, 26.7 mmol) and NHS (N-hydroxysuccinimide, 1.5 eq, 4.61 g, 40 mmol) in dichloromethane (DCM). 2. 1-ethyl-(3-dimethylaminopropyl)carbodiimide hydrochloride (EDC; 1.5 eq, 7.68 g, 40 mmol) and N, N-diisopropylethylamine (DIPEA; 1.5 eq, 5.18 g, 6.5 mL, 40 mmol) were added (see Note 3). 3. The mixture was stirred overnight at room temperature protected from direct light. After evaporation of the solvents the crude product was dissolved in ethyl acetate and extracted with water. The organic phase was dried over anhydrous sodium sulfate, filtered and evaporated to provide compound 1 (white powder, 12.6 g, 26.7 mmol, 100%). TLC (Rf ¼ 0.42, EtOAc–cyclohexane 4:6, UV and vanillin). 1 H NMR (400 MHz, CDCl3) δ 2.835 (s, 4H), 2.59 (t, J ¼ 6.9 Hz, 2H) 2.23 (t, J ¼ 7.1 Hz, 4H), 1.74 (q, 2H), 1.51 (q, 4H), 1.44–1.37 (m, 4H), 1.37–1.27 (m, 22H, alkyne chain), 0.88 (t, J ¼ 7 Hz, 3H) ppm (see Note 4). 13 C NMR (75 MHz, CDCl3) δ 177.2, 176.7, 77.4, 67.0, 35.9, 33.2–27.2, 23.9, 19.8 and 14.6 ppm.

3.1.2 C25diynetrioxaamine (2)

1. Compound 1 (1 eq, 5 g, 10.6 mmol) was dissolved in dry DCM. 2. 4,7,10-trioxa-1,13-tridecanediamine (2.5 eq, 6 g, 26.5 mmol) was added dropwise (see Note 5).

Fig. 3 Synthesis scheme of C25diynetrioxa-L-histidine (compound 4). EDC N-(3-dimethylaminopropyl)-Nethylcarbodiimide hydrochloride), DCM dichloromethane, DIPEA diisopropylethylamine, TFA trifluoroacetic acid, r.t. room temperature

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3. The reaction mixture was stirred at room temperature overnight. The solvent was evaporated under reduced pressure and the crude product was purified by column chromatography using DCM–MeOH–NH4OH 9:0.9:0.1 as eluent affording the desired compound 2 (white powder, 4.25 g, 7.4 mmol, 70%). TLC (Rf ¼ 0.25 (DCM–MeOH–NH4OH 9:0.9:0.1, UV and ninhydrin). 1 H NMR (400 MHz, MeOD) δ 3.69–3.63 (m, 8H), 3.61–3.58 (m, 2H), 3.51 (t, J ¼ 6.5 Hz, 2H), 3.23 (t, J ¼ 7.1 Hz, 2H), 3.10 (t, J ¼ 6.5 Hz, 2H), 2.22 (t, J ¼ 7.1 Hz, 4H), 2.16 (t, J ¼ 7.1 Hz, 2H), 1.93 (q, 2H), 1.76 (q, 2H), 1.63–1.59 (m, 2H), 1.54–1.47 (m, 4H), 1.44–1.37 (m, 4H), 1.37–1.27 (m, 22H, alkyne chain), 0.90 (t, J ¼ 7 Hz, 3H) ppm. 13 C NMR (75 MHz, MeOD) δ 176.4, 78.0–77.9, 71.7–70.04, 66.6, 40.3–37.3, 33.2–29.6, 27.2 (C23), 23.9 (C2), 19.8 and 14.6 ppm. LCMS (ESI+): m/z 577.5 [M+H]+. 3.1.3 C25diynetrioxaNα-(Tert-Butoxycarbonyl)L-Histidine (3)

1. C25diynetrioxaamine 2 (1 eq, 510 mg, 0.884 mmol) was dissolved in dry dichloromethane. 2. Nα-(tert-butoxycarbonyl)-L-histidine (1.2 eq, 271 mg, 1.06 mmol), 1-ethyl-(3-dimethylaminopropyl)carbodiimide hydrochloride (1.5 eq, 254 mg, 1.33 mmol) and N, N-diisopropylethylamine (1.5 eq, 0.2 mL, 1.33 mmol) were added. 3. The reaction mixture was stirred overnight under argon atmosphere at room temperature. Then the solvent was evaporated under reduced pressure and the crude product was purified by column chromatography using DCM–MeOH–NH4OH 9:0.9:0.1 as eluent affording compound 3 (yellowish oil, 620 mg, 86%). TLC (Rf ¼ 0.5 (DCM–MeOH–NH4OH 9:0.9:0.1, UV and ninhydrin). 1 H NMR (400 MHz, MeOD) δ 8.97 (s, 1H), 7.52 (s, 1H), 4.17 (t, J ¼ 7 Hz, 1H), 3.65–3.55 (m, 8H), 3.53–3.44 (m, 4H), 3.35–3.22 (m, 6H), 2.24 (t, J ¼ 6.5 Hz, 4H), 2.18 (t, J ¼ 7.5 Hz, 2H), 1.77–1.69 (m, 4H), 1.64–1.57 (m, 2H), 1.54–1.46 (m, 4H), 1.41 (s, 9H, Boc), 1.36–1.28 (m, 26H, alkyne chain), 0.9 (t, J ¼ 7 Hz, 3H) ppm.

3.1.4 C25diynetrioxa-LHistidine (4)

1. Compound 3 was dissolved in dry DCM. 2. Trifluoroacetic acid (TFA) (50 eq, 2.8 mL, 38.1 mmol) was added at 0
C.

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3. The reaction mixture was stirred under argon atmosphere at room temperature overnight. The solvent was evaporated under reduced pressure and the crude product was purified by column chromatography using DCM–MeOH–NH4OH 9:0.9:0.1 as eluent affording desired product 4 (yellowish oil, 360 mg, 66%). TLC (Rf ¼ 0.4 (DCM–MeOH–NH4OH 9:0.9:0.1, UV and ninhydrin). 1 H NMR (400 MHz, MeOD) δ 8.97 (s, 1H), 7.52 (s, 1H), 4.17 (t, J ¼ 7 Hz, 1H), 3.65–3.55 (m, 8H), 3.53–3.44 (m, 4H), 3.35–3.22 (m, 6H), 2.24 (t, J ¼ 6.5 Hz, 4H), 2.18 (t, J ¼ 7.5 Hz, 2H), 1.77–1.69 (m, 4H), 1.64–1.57 (m, 2H), 1.54–1.47 (m, 4H), 1.43–1.28 (m, 26H), 0.9 (t, J ¼ 7 Hz, 3H) ppm. 13 C NMR (75 MHz, MeOD) δ 174.9, 161.6, 135.5, 132.6, 118.4, 115.9, 115.4, 76.6, 76.5, 70.1, 69.9, 69.8, 68.5, 68.3, 65.0, 53.5 38.7, 36.6 36.4, 35.8, 31.7, 29.4–28.1, 25.6, 22.3, 18.3, 13.1 ppm. HRMS (ESI): m/z 714.554 ([M+H]+, calculated for C41H72N5O5 714.554). 3.2 Preparation of the Micelles

The following protocol describes how to formulate micelles at 5 mg/mL in water characterized by a diameter ranging from 7 to 10 nm. 1. Solubilize 5 mg of surfactant 4 in 200 μL of HCl 0.1 N and 800 μL ethanol. 2. Sonicate for 5 min (80 W, 25
C) until total solubilization of surfactant 4. 3. Evaporate the solution under reduced pressure until formation of a film (see Note 6). 4. Dissolve the film in 1 mL of deionized water. 5. Sonicate for 30 min (80 W, 25
C) to obtain micelles. 6. Polymerize the micelles for 4 h by UV-irradiation at 254 nm and 48 W in 1 mL quartz cuvettes using a Cross-Linker Bio-Link 254 (see Note 7). 7. Dialyze the micelles in 2000 MWCO dialysis cassettes starting with 70% ethanol–deionized water to 100% deionized water during a 4-day period (see Note 8). Depending on the protocol that is used for the formulation, surfactant 4 can also form nanofibers (Fig. 4). The latter structures however turned out to be inactive for siRNA transfection, as they are too big to get internalized by the cells [16] (see Note 9).

3.3 Characterization of the Micellar System

Due to their high molecular weight, the micelles cannot be characterized by classical NMR methods. We therefore performed

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Fig. 4 (a) Structure-dependent coloration of the solution: On the left the micellar form is yellow orange, on the right is shown the fluorescent red nanofiber form. (b) Electron microscopy of the micelles. (c) Fluorescence microscopy of Histidine-surfactant Nanofibers

CMC measurements, DOSY-NMR experiments, gel mobility shift assays and dynamic light scattering (DLS) to characterize these nano-objects. 3.3.1 Critical Micelle Concentration (CMC)

The CMC values were determined by using the pyrene inclusion method as described in the literature [17]. This method makes use of the environment specific fluorescence of pyrene probe as a detection of organized lipidic environment. 1. Prepare a set of dilution of the 10 mM stock solution of nonpolymerized micelles ranging from 2 mM to 1 μM in deionized water in order to have a final volume of 1 mL. 2. Add 1 μL of the 1 mM pyrene solution to 1 mL of each sample and stir vigorously. 3. Let the samples incubate at room temperature for 2 h until fluorescence measurement. 4. Settings of the fluorescence spectrophotometer are: UV excitation: 339 nm, Band pass 5 nm. 5. Measure the relative intensities at 373 nm and 384 nm. 6. Fluorescence of the micelles without pyrene is used as control. 7. The ratio of the relative fluorescence intensities I373 nm/I384 nm are plotted against log of millimolar concentrations. CMC is deduced from the inflexion point in the obtained graph (see Note 10).

3.3.2 DOSY-NMR Experiments

DOSY allows discriminating the NMR signals of different species according to their diffusion coefficients. Hence, this technique permits a visualization of the different populations present within a sample.

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The monomer surfactant population is first separated from the polymerized surfactant molecules by solvent extraction with deuterated methanol. 1. Formulate 5 mg/mL solution of nonpolymerized micelles (see Subheading 3.2 up to step 5), polymerized micelles (see Subheading 3.2 up to step 6), and dialyzed polymerized micelles (see Subheading 3.2 up to step 7). 2. Evaporate under vacuum 1 mL of each solution. 3. Solubilize the obtained solid film in deuterated methanol (see Note 11). 4. Analyze samples in DOSY-NMR experiments (see Note 12). 5. The percentage of covalently bridged surfactants versus the remaining monomer molecules can be calculated from the peak integration from the 2D-plot, as they show two distinct populations characterized by their respective diffusion coefficients (see ref. 8). 3.3.3 Gel Mobility Shift Assay

The agarose gel mobility shift assay is a method that allows determining the capacity of a given compound to complex nucleic acids. The electrophoretic analysis of a specific nucleic acid in presence of increasing concentrations of complexation agent allows determine the minimal amount of such a compound to retard the migration of nucleic acids during agarose gel electrophoresis. On one side the uncomplexed nucleic acid is detected as a discrete band. On the other side the complexation leads to the formation of particles, which are unable to migrate through the agarose mesh, and stay at the top of the gel. 1. Prepare a 1.3% agarose gel by dissolving 1 g of agarose in 75 mL of tris–acetate buffer (1, pH ¼ 7.5) and heating the suspension at 100
C using a microwave oven. After cooling down to about 60
C, add 15 μL of SYBRSafe to the agarose solution (see Note 13). 2. Pour the agarose solution in the electrophoresis tray, and position a comb into the gel. Using a Pasteur pipette remove bubbles from the gel and then wait until agarose has solidified (at least 30 min is required). Place the gel in the electrophoresis unit and remove the comb. Take care to add enough Tris–acetate buffer to cover the gel. 3. In order to analyze the complexation of siRNA by the cationic micelles 300 ng amounts of siRNA samples are distributed in Eppendorf tubes (4.3 μL, 5 μM) (see Note 14). 4. Dilute with HBG in order to obtain a final volume of 20 μL (be careful, volume of added micelles must be taken into account for determining the volume of HBG).

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Fig. 5 The siRNA was complexed with the 4 h polymerized micelles (PDA) in low salt conditions at increasing N/P ratios; Lane 1 (N/P ¼ 0): naked siRNA. Analysis by electrophoresis on a 1.3% agarose gel in tris–acetate buffer (40 mM, pH ¼ 7.5)

5. Add various volumes of micelles to obtain the desired N/P ratios (see Note 15). 6. Mix well and incubate for 1 h at room temperature. 7. Add 3 μL of loading buffer to each sample, mix, and then load 20 μL of samples into the wells of the agarose gel. Also include a sample containing siRNA without micelles as a control. 8. Electrophoresis conditions: 80 V for 30 min. 9. Visualize the agarose gel stained with SYBRSafe using an UV illuminator. The band corresponding to the free siRNA disappears at a N/P ratio above 1 meaning that total complexation has occurred (lanes 2, 3, and 4 in Fig. 5). We also note that no fluorescence is observed in the wells (start line) although siRNA is present in complexed form. The micelle may quench the fluorescence of SYBRSafe or SYBRSafe is no more able to insert into the complexed siRNAs. 3.3.4 Dynamic Light Scattering

The hydrodynamic diameter of the micelles and micelles–siRNA complexes were determined by using the Zetasizer Nano-ZS system using the following protocol.

Size Measurement of the Micelles

After formulation, the size of each sample should be measured to confirm the presence of the micelles. In our case, we developed the formulation protocol in such a manner that we obtain 10 nm diameter objects. 1. Prepare a 5 mg/mL solution of micelles as indicated in the formulation section. 2. Transfer 90 μL of the solution in the low-volume quartz cuvette. Place the cuvette in the cuvette block of the Zetasizer Nano-ZS instrument. 3. Set the parameters as described below:

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Temperature: 25
C. Index of refraction of material: 1.43. Index of refraction of pure water: 1.33. Viscosity of water used: 0.8872 cP. Sampling time: 55 s. Number of runs: 3. 4. Record a first measurement after equilibration of the sample (about 60 s) in the cuvette. The values correspond to average size  standard deviation of three runs. Perform at least three acquisitions to ensure the reproducibility of the measurements and the stability of the sample over time (see Note 16). 5. After measurements, the sample can be recovered for use in other experiments. A homogeneous population of small objects with a diameter below 10 nm should be observed. Size Measurement of the Micelles–siRNA Complexes

1. Prepare complexes by mixing 70 ng of siRNA (5 μM) prediluted in HBG (pH ¼ 7.5) with various amounts of micelles (7 mM) to form complexes at N/P ratios ranging from 5 to 50 (in 100 μL final volume). 2. Incubate at 25
C for 1 h. 3. Transfer 90 μL of the solution in the low-volume disposable cuvettes. Place the cuvette in the cuvette holder of the DLS measurement system. 4. Set the parameters as described below: Temperature: 25
C. Index of refraction of material: 1.43. Index of refraction of HBG 5%: 1.3374. Viscosity of HBG 5%: 1.1557 cP. Sampling time: 55 s. Number of runs: 3. 5. Record a first measurement after equilibration of the sample (about 60 s) in the cuvette. The values correspond to average size  standard deviation of three runs. Perform at least three acquisitions to ensure the reproducibility of the measurements and the stability of the sample over time (see Note 17). 6. After measurements, the sample can be recovered for use in other experiments. 7. The measured size of these siRNA-micelle complexes is around 60 nm at 1 h maturation in HBG buffer.

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The A549-Luc cells are grown in complete medium and maintained under selection with 0.8 μg/mL G418 at 37
C in a 5% CO2 humidified atmosphere. Cells are passaged using trypsin when they reach 90% confluency. Experiments should be anticipated in order to be sure to have enough cells. 1. The day prior transfection, use 2 mL of trypsin (for a T75 culture flask) to detach actively dividing cells (5 min at 37
C). 2. After addition of 8 mL of complete medium, the cells are centrifuged during 5 min at 200  g. 3. Remove the supernatant carefully and dilute the cells in 10 mL of fresh complete medium. 4. After having dissociated cells by making an up and down mixing with a pipette, cells are counted. 5. Determine how many wells are needed. Notably, transfection experiments are usually done using duplicates or triplicates. 6. The required volume of cell suspension to obtain a confluence of 90% at the end of the transfection experiment is then calculated (for A549-Luc cells, in 24-well tissue culture plates 25,000 cells/well are plated) (see Note 18). Dilute the cell suspension to obtain 50,000 cells/mL, in order to distribute 0.5 mL/well in 24 well plates. 7. The cells are then incubated at the appropriate cell culture conditions (a humidified tissue culture incubator at 37
C and 5% CO2).

3.4.2 siRNA Transfection

The day after the preparation of the 24-well plates, the cells are examined using a microscope to be sure that they are in good shape. All complexes are prepared 1 h prior to transfection. The following protocol is used to perform siRNA transfection in triplicates: 1. For a triplicate, to have a 10 nM final concentration of siRNA, take 4 μL of the 5 μM siRNA solution and dilute it in HBG 5% to have a final volume of 330 μL (by taking into account the volume of micelle that is needed) (see Note 19). 2. Add increasing volumes of a 5 mg/mL micelle solution so that the final concentrations of polymerized micelles ranges from 9 to 25 μg/mL (in each well in the 24-well cell culture plates) (see Note 20). 3. Use the same protocol for control experiments with a control siRNA. This control is important in order to ascertain that the silencing is gene specific. 4. In each experiment, a commercial transfection reagent (e.g., INTERFERin) should be added and used according to the

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recommendation of the supplier to control the quality of the siRNA (positive control). 5. Mix the siRNA with the micelles, centrifuge very shortly to pull down all the drops, and let the tube at room temperature for about 1 h. 6. Remove the culture medium from the cells by aspiration. If the cells that are used adhere well, the cultures may be rinsed with PBS in order to remove serum-containing proteins from the wells and replace it with serum-free medium (0.5 mL). 7. Add 100 μL of micelle/siRNA complexes solution into each well of the triplicate and incubate at 37
C for 4 h. 8. After 4 h of incubation, 60 μL of fetal bovine serum are added in each well to obtain 10% serum concentration. 9. Incubate at 37
C for 48 h. 10. Analyze the cells for luciferase expression and protein content. 3.4.3 Luciferase Assay

To determine luciferase activity, the following protocol is used: 1. After 48 h of transfection, remove carefully the culture medium from the 24-well plates. 2. Wash each well with 1 mL of PBS. 3. Add 100 μL of 1 lysis buffer to each well. 4. After 30 min of gentle shaking, the cell lysate is recovered and transferred into annotated 1.5 mL Eppendorf tubes. 5. Centrifuge the tubes for 5 min at 20,800  g to pellet debris. 6. Take 2 μL of the supernatant of each tube and transfer them into the wells of a white 96-well plate. 7. Measure the bioluminescence using a luminometer that automatically injects 50 μL of luciferase substrate (luciferin). 8. Plate is shaken for 3 s and then luminescence is measured over 1 s. 9. Calculate the light units for 10 s/100 μL of sample by multiplying the relative light units measured by 10 and 50. 10. After having measured the protein content (see below), express the efficiency as Relative Light Units (RLU)/10 s/mg of protein. Luciferase expression must be compared to nontransfected cells (untreated cells) but also to cells transfected with noncoding siRNA to determine the specific inhibition by taking into account possible off-target or cytotoxicity (see Fig. 6).

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Fig. 6 siRNA delivery experiment performed on A549-luc cells using various concentrations of micelle-Histidine and 10 nM siRNA. Luciferase expression compared to nontransfected cells and the percentage of cellular proteins level also compared to nontransfected cells after 48 h of transfection are represented. Results and standard deviations are given as a mean of triplicate 3.4.4 Protein Assay

Protein concentration in cell lysate was measured using a BCA assay kit according to the following protocol. 1. Knowing that 1 mL/tube is needed, prepare the mixed BCA solution (with solution A and solution B) with the following A–B ratio 50:1. 2. In parallel, prepare the standard solution of BCA ranging from 1 to 12 μg of bovine serum albumin (BSA) starting from a stock solution at 2 μg/μL of BSA. Then, add 15 μL of 1 lysis buffer to each standard well. 3. Prepare also two blanks containing only 15 μL of 1 lysis buffer. 4. Transfer 15 μL of each cell lysate into plastic tubes (see Note 21). 5. Add 1 mL of mix BCA solution in each tube including blanks. 6. Heat at 60
C for 30 min (change of coloration from green to purple). 7. Transfer the solution contained in tubes into 1 mL semi-micro cuvettes. 8. Read absorbance at λ ¼ 562 nm and determine the protein content in each sample by using the BSA standard curve and multiplying by 100/15.

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3.4.5 Cell Viability Assay

Cytotoxicity of vectors can be measured by MTT assay. Indeed, tetrazolium MTT is reduced by active cells resulting in purple formazan quantifiable by spectrophotometry. Hence, cell proliferation (and thus cell viability) can be measured. 1. Twenty-four hours prior to assay, cells are plated at 10  103 per well in 100 μL complete culture media in 96-well tissue culture plates. 2. Replace culture medium, with RPMI medium without FBS. 3. Add different concentrations of micelles +/ siRNA in serumfree conditions for 4 h. 4. To provide blank for absorbance readings representing total cell death, add 10% DMSO in three control wells. 5. At 4 h add 10 μL of serum per well. 6. After 48 h, add 100 μL of the 1 mg/mL MTT reagent solution and incubate 2 h at 37
C until purple precipitate is visible. 7. Remove carefully the medium and add 100 μL of DMSO per well to dissolve purple precipitate. 8. After 30 min of stirring, record absorbance at 570 nm in a plate reader (SAFAS instruments). By comparing the cytotoxicity results obtained with dialyzed and nondialyzed we concluded that monomers molecules remaining in nondialyzed micelles largely contribute to residual toxicity. As expected extensive dialysis of polymerized micellar solutions can thus be used to reduce the cytotoxicity of the nanovector solutions. This observation is in accordance with the proven presence of 25% remaining monomer as seen by DOSY-NMR experiments in photopolymerized micelles, while the monomer population is decreased in dialyzed samples.

4

Notes 1. Stored at 20
C and protected from light to prevent photopolymerization. 2. All chemical steps that involve a “diyne” part should be protected from light by using amber glassware to prevent photopolymerization. They should be stored at 20
C. If the compound starts to photopolymerize, it will turn blue and will not be soluble anymore. 3. We recommend using EDC as coupling agent rather than DCC (N,N0 -dicyclohexylcarbodiimide) because of its good solubility in water. Hence, a simple aqueous workup by extraction is efficient to remove the excess of EDC. For this step, it is recommended to perform only a workup and no column

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chromatography because of the sensitivity of the NHS ester derivative (N-hydroxysuccinimide ester) on silica gel. Thus a white powder should be obtained after filtration of the organic phase, dried over anhydrous sodium sulfate. 4. Recorded chemical shifts values were reported in parts per million (δ) and calibrated using residual signals from partially protonated solvents (CHCl3: 1H 7.26 ppm; 13C 77.16 ppm or MeOH 1H 3.31 ppm; 13C: 49.0 ppm). Data were represented as follows, chemical shift, multiplicity (s ¼ singlet, d ¼ doublet, t ¼ triplet, q ¼ quadruplet, and m ¼ multiplet), coupling constants (J in Hz), integration and attribution. NMR spectra were analyzed using the commercial software NMRnotebook (NMRtech). 5. 4,7,10-trioxa-1,13-tridecanediamine must be added in excess to limit the formation of the bicoupling compounds. The bicoupling and the monocoupling compounds can be easily separated using column chromatography (first fraction: Bicoupled product with two acyl chains). To facilitate the purification, ammonia is added into the eluent to deprotonate the amino group. 6. At the end, the micelle solution will be acidic with a pH ranging from 4 to 6 according to the efficiency of the evaporation process of excess HCl during chlorhydrate formation by vacuum pump. 7. The ene–yne system resulting from the polymerization is a conjugate system that gives rise to a bathochromic shift. Hence, in our case, as the angle of polymerization in spherical object is not optimal, polydiacetylenic micelles are slightly conjugated and show absorption maxima in UV-A to blue light (400 nm) and therefore appear yellow. So if you generate micelles, the solution should be yellow, whereas PDA nanofibers containing solutions will be red. After testing in in vitro assays different times of polymerization of micelles, 4 h has proven to be the optimal condition to have the best balance between efficiency and cytotoxicity [8]. 8. This formulation protocol turns out to be pretty straightforward and robust at the scale of 5 mg of surfactant. When using larger scales, the biological activity of the resulting micelles was shown to be lower for reasons that are still unclear. We recommend controlling each batch in biological assays before pooling them. 9. Protocol to obtain nanofibers: Surfactant 4 was dissolved in ethanol at 60
C under sonication (60 mg/6 mL). This warm solution was filtered through a cotton pad (in a glass pipette, to remove minor polymerized contaminants) and injected into ethanol-water (12 mL/42 mL) mixtures to obtain final 30%

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ethanolic solutions at a concentration of 1 mg/mL. This solution was placed at +4
C for 18 h, where it transformed into an opaque gel-like suspension. UV polymerization was performed on 5 mL batches in a UV-Crosslinker. The batches were UV irradiated in open 6-well culture plates for 10 min at 254 nm. During the UV irradiations the suspensions turned dark blue. The dark-blue suspensions were acidified by adding 50 μL diluted chlorhydric acid (1 N) to each of the 5 mL batches, which were transferred in 15 mL Falcon tubes. The suspensions were then heated under sonication at 55
C for 1 h. This led to a drastic color change. The suspensions turned crimson red, and the polymerized aggregates dissociated into nanofibers. Their homogeneity can be assessed by direct observation by fluorescence microscopy (rhodamine filter set, 600-fold magnification). 10. The CMC is deduced from the plot of all the measurement points recorded at the relative intensities at 373 nm and 383 nm. The ratios of the relative fluorescence intensities I373nm/I383nm were plotted against log of millimolar concentrations. The two linear curves crossing at one inflexion point should be obtained. CMC is deduced from the inflexion point in the obtained graph. The linear trendlines give access to the two equations of the curves that intersect at a value of concentration corresponding to the CMC. 11. The remaining solid was solubilized in deuterated methanol in order to separate monomers from polymerized lipids out of the micelles; micelles are indeed supposed to be unstable in methanol. 12. The DOSY spectra are obtained by applying an Inverse Laplace Transform (ILT) along the diffusion axis, using the commercial software NMRnotebook. The percentage of covalently cross-linked surfactants versus monomer surfactant molecules were calculated from the peak integration from the 2D-plot. 13. The pH and the nature of the running buffer are crucial for gel retardation assay with these micelles. Indeed, to have correct results, you should use a running buffer without any anionic charged molecules or complexing agents that could compete with the siRNA complexation. EDTA should be banished. Moreover, the pH of the running buffer should be ranged between 7 and 7.5. Above 7.5, the primary amine of the histidine group will not be protonated anymore and thus the micelle will no longer be positively charged. 14. We recommend using a minimal amount of 300 ng to realize the electrophoresis to have a clearly visible siRNA band.

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15. To know the global charge of the complexes formed between the micelles and the siRNA, N/P ratio is calculated. It corresponds to the positive charges contained in the surfactant versus the negative charge of the siRNA. Here, at pH ¼ 7.5 the surfactant has only one positive charge (the protonated primary amine of the histidine group) and no negative charge and the siRNA possesses 42 negative charges because there are 42 phosphate groups. Therefore, N/P ratio can be calculated with the following equation: N The amount of micelles  1 ðin molÞ ¼ P The amount of siRNA  42 ðin molÞ 16. To verify the quality of the data, it is recommended to check the correlogram. All the correlation curves should be S-shaped, with a maximal correlation coefficient higher than 0.6. The automatic attenuator setting at working concentrations should be at medium values, indicating a good count rate. If the automatic attenuator shifts wide open (values from 10 to 11), the sample is too diluted to enable a correct size measurement. 17. With our system, at N/P above 10, complexes with a size under 100 nm of diameter should be obtained. 18. The initial number of cells plated per well was calculated in such a way that we have a confluency of 90% after 72 h (knowing that this cell line has a fast division rate). 19. The amount of products is calculated by taking into account a dead volume of 10 μL per well. 20. For siRNA experiments, it is important to choose the amount of micelles that gives the highest transfection efficiency associated with the lowest cytotoxicity. Indeed, cell toxicity may lead to nonspecific knockdown of genes. Of note, the efficiency of the vector is cell line dependent. 21. We recommend centrifuging samples a second time (5 min at 20,800  g) prior to the protein quantification to be sure that no more cellular debris is present in the supernatant which would cause huge variations between individual quantifications.

Acknowledgments This work was supported by the Labex Medalis and by the FRM (fondation pour la recherche me´dicale). M.R. has a financial support from MESR (Ministe`re de l’Enseignement supe´rieur et de la

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Recherche). The authors would like to thank the chemical analysis service platform of the faculty of pharmacy of Illkirch (France) and Bruno Vincent (chemical analysis service platform at the University of Strasbourg) for DOSY-NMR experiments. References 1. Mel’nikov SM, Sergeyev VG, Yoshikawa K (1995) Transition of double-stranded DNA chains between random coil and compact globule states induced by cooperative binding of cationic surfactant. J Am Chem Soc 117:9951–9956 2. Clamme JP, Bernacchi S, Vuilleumier C et al (2000) Gene transfer by cationic surfactants is essentially limited by the trapping of the surfactant/DNA complexes onto the cell membrane: a fluorescence investigation. Biochim Biophys Acta 1467:347–361 3. Blessing T, Remy J-S, Behr J-P (1998) Monomolecular collapse of plasmid DNA into stable virus-like particles. Proc Natl Acad Sci U S A 95:1427–1431 4. Lleres D, Dauty E, Behr JP et al (2001) DNA condensation by an oxidizable cationic detergent. Interactions with lipid vesicles. Chem Phys Lipids 111:59–71 5. Dauty E, Remy J-S, Zuber G et al (2002) Intracellular delivery of nanometric DNA particles via the folate receptor. Bioconjug Chem 13:831–839 6. Morin E, Nothisen M, Wagner A et al (2011) Cationic polydiacetylene micelles for gene delivery. Bioconjug Chem 22:1916–1923 7. Neuberg P, Perino A, Morin-Picardat E et al (2015) Photopolymerized micelles of diacetylene amphiphile: physical characterization and cell delivery properties. Chem Commun 51:11595–11598 8. Ripoll M, Neuberg P, Kichler A et al (2016) pH-responsive nanometric polydiacetylenic micelles allow for efficient intracellular siRNA delivery. ACS Appl Mater Interfaces 8:30665–30670

9. Behr J-P (1997) The proton sponge: a trick to enter cells the viruses did not exploit. Chim Int J Chem 51:34–36 10. Neuberg P, Kichler A (2014) Recent developments in nucleic acid delivery with polyethylenimines. Adv Genet 88:263–288 11. Kichler A, Leborgne C, Coeytaux E et al (2001) Polyethylenimine-mediated gene delivery: a mechanistic study. J Gene Med 3:135–144 12. Gravel E, Ogier J, Arnauld T et al (2012) Drug delivery and imaging with polydiacetylene micelles. Chem Eur J 18:400–408 13. Yang D, Zou R, Zhu Y et al (2014) Magainin II modified polydiacetylene micelles for cancer therapy. Nanoscale 6:14772–14783 14. Theodorou I, Anilkumar P, Lelandais B et al (2015) Stable and compact zwitterionic polydiacetylene micelles with tumor-targeting properties. Chem Commun Camb 51:14937–14940 15. Yao D, Li S, Zhu X et al (2017) Tumor-cell targeting polydiacetylene micelles encapsulated with an antitumor drug for the treatment of ovarian cancer. Chem Commun 53:1233–1236 16. Neuberg P, Hamaidi I, Danilin S et al (2018) Polydiacetylenic nanofibers as new siRNA vehicles for in vitro and in vivo delivery. Nanoscale 10:1587–1590 17. Aguiar J, Carpena P, Molina-Bolı´var JA et al (2003) On the determination of the critical micelle concentration by the pyrene 1:3 ratio method. J Colloid Interface Sci 258: 116–122

Chapter 8 Lipids for Nucleic Acid Delivery: Cationic or Neutral Lipoplexes, Synthesis, and Particle Formation Michel Bessodes, Helene Dhotel, and Nathalie Mignet Abstract Lipidic vesicles have been extensively studied for their capacity to condensate and deliver nucleic acids to the cells. Many different amphiphilic lipidic structures have been proposed each of them bringing some advances in nonviral gene transfection. The ionic or neutral nature of the lipids induces tremendous differences in the behavior of the corresponding liposomes, from the complexation of nucleic acid to the delivery to the cell. An efficient delivery in vitro or in vivo also depends closely on the structure of the lipids and very often, efficient liposomes in vitro have been found useless for in vivo administration. We describe in this chapter the chemical synthesis of two different lipids, one cationic and the other essentially neutral, and the formulation to obtain liposomes and DNA–liposome complexes. The different ways and tricks for the formulation of the two different structures are especially highlighted. Key words Cationic lipid, Noncationic lipid, Lipid synthesis, Ethanolic injection, Liposome formulation, Lipoplex characterization, DNA–lipid complex stabilization

1

Introduction Amphiphilic lipids are able to auto-assemble to form spherical vesicles such as liposomes in aqueous solution [1]. Synthetic lipids, like phospholipids, are amphiphiles which tend to organize themselves in an aqueous medium in order to maximize the hydrophobic interactions, repulsing the water content from the lipidic part, therefore reaching the more thermodynamically stable state. Their structure should allow the lipid to expose the cationic charges of their hydrophilic head in order for them to interact with DNA. The lipoplexes formed between the lipids and DNA protect DNA from enzymatic degradation and increase its cellular uptake leading to high levels of transfection in vitro [2]. We have earlier described two families of such lipids, the first family contains cationic head able to strongly condensate plasmidic DNA into liposomes through electrostatic interactions [3, 4], the second family bears thiourea

Manfred Ogris and Haider Sami (eds.), Nanotechnology for Nucleic Acid Delivery: Methods and Protocols, Methods in Molecular Biology, vol. 1943, https://doi.org/10.1007/978-1-4939-9092-4_8, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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Scheme 1 (a) Chemical structure of our lipopolyamine (DMAPAP) and lipopolythiourea lead compounds (DDSTU). (b) Their efficacy in terms of luciferase expression by a luciferase encoding gene is shown on B16 cell lines in presence of 10% serum (SVF). (c) DNA release from the complexes was shown to be faster with DDSTU as compared to DMAPAP on Hela cells [7]. Lipids are labeled with rhodamine (red), and the gene is labeled with fluorescein (green)

groups known to promote hydrogen bonds with phosphate containing macromolecules [5–7]. A thorough structure to physicochemical and biological properties relationship has been established, thus allowing to highlight two structures (DMAPAP and DDSTU, Scheme 1) in each family with particularly interesting properties. The cationic liposomal formulation (DMAPAP/DOPE 1/1) is nowadays used as a reference in our laboratory as it transfects as efficiently as lipofectamine adherent cultured cells in vitro. However, for in vivo applications, we developed noncationic lipids that we called thiourea lipids, in order to increase blood circulation of the lipoplexes and reduce nonspecific interactions. We showed in vitro that cationic lipoplexes were internalized more efficiently than thiourea lipoplexes by the cells, and that thiourea lipoplexes were able to release DNA into the cells more efficiently than cationic complexes leading to gene transfection (Scheme 1) [7]. When administered i.v. to mice, thiourea complexes were also shown to remain longer in the circulation [6]. The practical syntheses, autoassembly of the lipids, DNA association, characterization of the lipoplexes and their stability assessment are described in this chapter.

2

Materials

2.1 Abbreviations of the Lipids Used

DOPE: L-α-Dioleoyl Phosphatidylethanolamine; The chemical names were generated with Symyx Draw 4.0® software, based on IUPAC rules. The cationic lipid whose name according to the

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nomenclature is 2-{3-[Bis-(3-amino-propyl)-amino]-propylamino}-N-ditetradecylcarbamoyl methyl-acetamide or RPR209120 that we called DMAPAP was previously described, using a different chemical route, in the supporting information of Thompson et al. [8]. Synthesis of 2-[2-(didecylamino)-2-oxo-ethoxy]-N[2-(2,3-dihydroxypropylcarbamothioylamino)-1-[(2,3-dihydroxypropylcarbamothioylamino)methyl]ethyl]acetamide called DDSTU, was described slightly differently in ref. 9. 2.2 Chemicals and DNA Provided or Synthesized

The chemicals were purchased from Sigma-Aldrich Company. The solvents were from Carlo Erba-SDS (analytical grade). TLC were performed on silica gel 60F254 aluminum sheets from Merck. L-α-Dioleoyl Phosphatidylethanolamine (DOPE) was purchased from Avanti Polar Lipids, Picogreen® from Molecular Probes. pDNA was obtained as described in ref. 7.

2.3

Size and zeta potentials measurements were performed on a Zeta Sizer NanoSeries from Malvern Instruments equipped with a MPT2 autotitrator. Fluorescence was measured on a multilabel plate reader Wallac Victor2 1420 Multilabel Counter, PerkinElmer, France, equipped with excitation and emission filters (350
10 nm, 450
10 nm).

3

Equipment

Methods

3.1 Synthesis of DMAPAP Cationic Lipid (See Note 1 and Scheme 2) 3.1.1 Tert-Butyl 2-(3-Hydroxypropylamino) Acetate (1)

3.1.2 Tert-Butyl 2-[3-Hydroxypropyl-(2,2,2Trifluoroacetyl)Amino] Acetate (2)

To a cooled solution of 3-aminopropanol (196 ml, 2.56 mol, 25 eq) in DCM (250 ml), tert-butylbromoacetate (20 g, 102.5 mmol, 1 eq) in DCM (200 ml) was added dropwise. After 2 h the reaction medium was warmed to room temperature and kept three more hours. The solution was then washed with NaHCO3 sat (3  150 ml), sat. NaCl (150 ml). After drying over magnesium sulfate, filtration and evaporation, a colorless oil was obtained (17.8 g, 92%). TLC (Rf ¼ 0.55; DCM/MeOH 8/2; ninhydrin or I2/ H2SO4). 1H NMR (300MHZ, (CD3)2SO, δ ppm): 1.44 (s, 9H), 1.54 (m, J ¼ 6.5 Hz, 2H), 2.56 (t, J ¼ 6.5 Hz, 2H), 3.19 (s, 2H), 3.46 (t, J ¼ 6.5 Hz, 2H), 3.70–4.70 (m, 1H). MS (DC/I, m/z) 190 (MH+). Compound 1 (17.65 g, 93.3 mmol, 1 eq) was dissolved in DCM (100 ml) and cooled to 0  C in an ice-water bath. Triethylamine (26 ml, 186.6 mmol, 2 eq) then trifluoroacetic anhydride (21.5 g, 102.6 mmol, 1.1 eq) were added dropwise. The reaction mixture was stirred overnight at room temperature. The solution was washed with NaHCO3 sat (3  50 ml), KHSO4 0.5 M (3  50 ml) and sat. NaCl (50 ml). The organic phase was dried

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Scheme 2 (i), (TFA)2O, Et3N,DCM; (ii), CBr4, Ph3P, CH3CN; (iii), K2CO3, CH3CN; (iv), 5 + TFA, DCM; (v), Na2CO3, EtOH, reflux; (vi), Boc-Gly, BOP, DIEA, DCM; (vii), TFA, DCM; (viii), BOP, DIEA, CH2Cl2; (ix), 1 N NaOH, THF

over magnesium sulfate, filtrated and concentrated to a pale-yellow oil (24.7 g, 93%). TLC (Rf ¼ 0.25, C6H12/EtOAc 1/1, ninhydrin or I2/ H2SO4, UV). 1H NMR (300 MHz, (CD3)2SO, δ ppm): (see Note 2) 1.44, 1.46 (2 s, 9H); 1.60–1.80 (m, 2H); 3.35–3.55 (m, 4H); 4.09, 4.25 (m, 2H); 4.57, 4.61 (t, J ¼ 5 Hz, 1H). MS (D/CI, m/z) 303 (M NH4+). 3.1.3 Tert-Butyl 2-[3-Bromopropyl-(2,2,2Trifluoroacetyl)Amino] Acetate (3)

To a stirred solution of 2 (10 g, 35 mmol, 1 eq) and triphenylphosphine (12.4 g, 47.3 mmol, 1.35 eq) in THF (150 ml), the solution was kept at 15–20  C with an ice bath then carbon tetrabromide (15.1 g, 45.6 mmol, 1.3 eq) in acetonitrile (60 ml) was added dropwise. After 4 h the reaction medium was concentrated, taken up in ethyl acetate and filtrated on paper filter. The filtrate was concentrated to dryness, taken up in cyclohexane and filtrated on sintered glass (n 3) then on sintered glass (n 4). After a final concentration and purification on silica column (C6H12/EtOAc 9/1), a lightly yellow colored oil was obtained (10.4 g, 85%).

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TLC (Rf ¼ 0.6, C6H12/EtOAc 1/1, ninhydrin or I2/H2SO4, UV); 1H NMR (300 MHz, (CD3)2SO, δ ppm) (see Note 2) 1.43, 1.45 (s, 9H); 2.05–2.20 (m, 2H); 3.50–3.65 (m, 4H), 4.11, 4.28 (m, 2H). MS (D/CI m/z) M NH4+. 3.1.4 2,2,2-Trifluoro-N[3-[3-[(2,2,2Trifluoroacetyl)Amino] Propylamino]Propyl] Acetamide (4)

3,30 -imino-bispropylamine (35 g, 266.7 mmol, 1 eq) was dissolved in anhydrous THF (150 ml) under an argon atmosphere. The solution was cooled to 0  C in an ice bath and ethyl trifluoroacetate (65 ml, 546.8 mmol, 2.05 eq) was added dropwise (see Note 3). After 3 h the reaction medium was warmed to room temperature, then left for 2 more hours under argon and the solution concentrated. After drying overnight in a vacuum oven, the colorless oil was crystallized at 4  C and a white powder was obtained (85.3 g, 99%). TLC (Rf ¼ 0.7, EtOH/NH4OH 8/2, ninhydrin). 1H NMR (300 MHz, CDCl3, δ ppm): 1.74 (m, J ¼ 6 Hz, 4H); 2.73 (t, J ¼ 6 Hz, 4H); 3.46 (m, 4H); 8.18 (m, 2H). MS (D/CI, m/z) 324 MH+.

3.1.5 Tert-Butyl 2-[3-[Bis [3-[(2,2,2-Trifluoroacetyl) Amino]Propyl]Amino] Propyl-(2,2,2Trifluoroacetyl)Amino] Acetate (5)

To a solution of compound 3 (26 g, 74.7 mmol, 1 eq) and compound 4 (24.1 g, 74.7 mmol, 1 eq) in acetonitrile (130 ml), potassium carbonate was added (30 g, 224 mmol, 3 eq) and the mixture was heated to reflux during 6 h. The reaction medium was then filtrated on sintered glass (n 4) and evaporated to dryness. Chromatography on silica (cyclohexane/EtOAc: 4/6 then after the first impurity cyclohexane/EtOAc:2/8) yielded 5 as a paleyellow oil (16.6 g, 38%). TLC (Rf ¼ 0.35, EtOAc, I2/H2SO4, UV). 1H NMR (300 MHz, CDCl3, δ ppm): (see Note 2) 1.47, 1.48 (2 s, 9H); 1.65–1.85 (m, 6H); 2.40–2.60 (m, 6H); 3.40–3.55 (m, 6H); 3.97, 4.07 (m, 2H); 7.45–7.65 (m, 2H). MS (D/CI, m/z) 591 MH+.

3.1.6 2-[3-[Bis[3-[(2,2,2Trifluoroacetyl)Amino] Propyl]Amino]Propyl(2,2,2-Trifluoroacetyl) Amino]Acetic Acid (6)

Trifluoroacetic acid (50 ml) was added to a solution of compound 5 (15.8 g, 28.76 mmol) in DCM (50 ml). The mixture was stirred at room temperature during 2 h, and then concentrated, taken up in cyclohexane then CHCl3 A faint yellow gum was obtained (18.7 g, 100%). 1H NMR (300 MHz, CDCl3, δ ppm): (see Note 2) 1.80–2.10 (m, 6H); 3.12 (m, 6H); 3.29 (m, 4H); 3.50 (m, 2H); 4.13, 4.29 (m, 2H); 9.50–9.75 (m, 2H). MS (D/CI, m/z) 535 MH+.

3.1.7 Ditetradecylamine Chlorohydrate (7)

Bromotetradecane (5 g, 18 mmol) and tetradecylamine (3.85 g, 18 mmol) were dissolved in absolute ethanol (30 ml). Sodium carbonate (4.8 g, 45 mmol, 2.5 eq) was added and the mixture was refluxed overnight. The reaction medium was then evaporated, taken up in dichloromethane (100 ml) and washed successively with water (3  20 ml) and sat. NaCl (1  40 ml). The organic phase was dried over sodium sulfate and concentrated. After

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salification (see Note 4) the product crystallized in isopropanol (3.3 g; 41%). TLC (Rf ¼ 0.55; DCM/MeOH 9/1; ninhydrin or I2/H2SO4). 1H NMR (300 MHz, CDCl3, δ ppm): 0.88 (t, J ¼ 7 Hz, 6H); 1.15–1.45 (m, 44H); 1.90 (m, 4H); 2.90 (m, 4H); 9.48 (m, 2H). MS (D/CI, m/z) 410 (MH+). 3.1.8 Tert-Butyl N-[2-[Di (Tetradecyl)Amino]-2-OxoEthyl]Carbamate (8)

To a solution of ditetradecylamine chlorohydrate (3 g, 7.33 mmol), Boc glycine (1.41 g, 8 mmol, 1.1 eq) in dichloromethane (50 ml), and BOP (3.56 g, 8 mmol, 1.1 eq) cooled to 0  C in an ice-water bath, diisopropylethylamine (6.2 ml, 36.65 mmol, 5 eq) was added dropwise. The solution was stirred at ambient temperature for 3 h then concentrated. Ethyl acetate (150 ml) was added and the solution was washed successively with 0.5 M KHSO4 (3  50 ml), sat. NaHCO3 (3  50 ml), and sat. NaCl (2x50 ml); it was dried over Na2SO4, filtered and concentrated to dryness (3.57 g, 86%). TLC (Rf ¼ 0.43; DCM/MeOH 99,5:0.5 ninhydrin or I2/H2SO4); 1H NMR (400 MHz, CDCl3, δ ppm): 0.87 (t, J ¼ 7 Hz, 6H); 1.16–1.35 (m, 44H); 1.46–1.64 (m, 4H); 3.13 (dd, J ¼ 8 Hz, 2H); 3.30 (dd, J ¼ 8 Hz, 2H); 3.94 (dd, J ¼ 4 Hz, 2H); 5.57 (m, 1H).

3.1.9 Amino-N,N-Di (Tetradecyl)Acetamide; TFA Salt (9)

The above product (3.57 g; 6.3 mmol) was dissolved in an aqueous solution of trifluoroacetic acid (90%; 15 ml) and left at ambient temperature for 1 h. The solution was evaporated, taken up in CHCl3 and then evaporated under reduced pressure and the product crystallized in diethyl ether (3.6 g; 98%). TLC (Rf ¼ 0.1; DCM/MeOH 99:1; ninhydrin or I2/H2SO4); 1H NMR (400 MHz, CDCl3, δ ppm): 0.87 (t, J ¼ 7 Hz, 6H); 1.2–1.35 (m, 44H); 1.45–1.58 (m, 4H); 3.13 (dd, J ¼ 8 Hz, 2H); 3.31 (dd, J ¼ 8 Hz, 2H); 3.88 (m, 2H).

3.1.10 N[3-[3-[[2-[[2-[Di (Tetradecyl)Amino]-2-OxoEthyl]amino]-2-Oxo-Ethyl](2,2,2-Trifluoroacetyl) Amino]Propyl-[3-[(2,2,2Trifluoroacetyl)Amino] Propyl]Amino]Propyl]2,2,2-Trifluoro-Acetamide (10)

2-Amino-N,N-di(tetradecyl)acetamide 9 (3.6 g; 6.2 mmol) and 2-[3-[bis[3-[(2,2,2-trifluoroacetyl)amino]propyl]amino]propyl(2,2,2-trifluoroacetyl)amino]acetic acid 6 (2.98 g; 5.58 mmol; 0.9 eq) were dissolved in dichloromethane (20 ml), BOP (3 g, 6.8 mmol, 1.1 eq) were added; the mixture was cooled to 0  C in an ice-water bath, and diisopropylethylamine (5.3 ml, 31 mmol, 5 eq) was added dropwise. The solution was stirred at ambient temperature for 1 h, and then concentrated. The residue was redissolved in ethyl acetate (150 ml) and washed successively with 0.5 M KHSO4 (3  20 ml), sat. NaHCO3 (3  20 ml), water, and sat. NaCl solution. It was dried over Na2SO4, filtered and concentrated to give 10 (4.67 g, 85%). TLC (Rf ¼ 0.1; DCM/MeOH 99:1; ninhydrin or I2/H2SO4); 1H NMR (400 MHz, CDCl3, δ ppm): 0.87 (t, J ¼ 7 Hz, 6H); 1.19–1.34 (m, 44H); 1.44–1.93 (m, 10H); 3.15 (m, 2H); 3.30 (m, 2H); 3.43 (m, 4H); 3.55 (m, 2H);

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4.01–4.19 (m, 4H); 6.99 (m, 1H); 7.64 (m, 1H); 7.74 (m, 1H). MS (ES, m/z) 983.5 M. 3.1.11 3-[Bis (3-Aminopropyl)Amino] Propyl-[2-[[2-[Di (Tetradecyl)Amino]-2-OxoEthyl]Amino]-2-Oxo-Ethyl] Ammonium; Hydrochloride (11, DMAPAP)

The compound 10 (4 g, 4 mmol) was dissolved in THF (60 ml), 1 N NaOH (60 ml) was added and the mixture was stirred overnight at ambient temperature. The solution was evaporated to dryness, redissolved in EtOH, evaporated, then redissolved in DCM and washed with water then dried with sodium sulfate, concentrated and chromatographed on a short column of silica eluted with DCM/MeOH/NH4OH (45:45:10). The fractions containing the nonprotonated form of 11 were collected and evaporated. Subsequent treatment with 6 N HCl in isopropanol, evaporation, and lyophilization gave 11 (2.8 g, 95%). TLC (Rf ¼ 0.2; DCM/MeOH/NH4OH 45:45:10; ninhydrin or I2/ H2SO4); 1H NMR (400 MHz, DMSO d6, δ ppm): 0.86 (t, J ¼ 7 Hz, 6H); 1.17–1.35 (m, 44H); 1.37–1.59 (m, 4H); 1.97–2.20 (m, 6H); 2.86–3.40 (m, 16H); 3.80 (m, 2H); 4.01 (m, 2H). MS (ES, m/z) 695.8 M, 696.8 M-H+, 697.8 M-2H+, 698.8 M-3H+.

3.2 Synthesis of DDSTU Thiourea Lipid (See Scheme 3)

In a 250 ml round-bottom flask, 2-amino-1,3-propanediol (5 g, 54.9 mmol) was dissolved in ethanol (150 ml) and di-tert-butyldicarbonate (11.98 g, 54.9 mmol) was added at 0  C. The mixture was stirred at ambient temperature during 10 h. It was then evaporated under reduced pressure and the white residue taken up in a minimum of dichloromethane; heptane was then added until turbidity. The product crystallized at 0  C and was filtered on a sintered funnel (n 4) (9 g; 86%). Mp 82  C. TLC (Rf ¼ 0.48; DCM/MeOH 9:1; ninhydrin); 1H NMR (400 MHz, CDCl3, δ ppm): 1.45 (s, 9H, CH3), 3.24 (m, 2H, OH), 3.67–3.83 (m, 5H, CH, CH2OH), 5.34 (d, 1H, J ¼ 6.6 Hz, NH).

3.2.1 Tert-Butyl N-[2-Hydroxy-1(Hydroxymethyl)Ethyl] Carbamate (12)

3.2.2 [2-(TertButoxycarbonylamino)-3Methylsulfonyloxy-Propyl] Methanesulfonate (13)

Compound 12 (4 g, 20.92 mmol) was dissolved in dichloromethane (47 ml), and the solution was placed under a nitrogen atmosphere and cooled to 0  C.Ttriethylamine (8.79 ml, 62.6 mmol) was added dropwise and methanesulfonyl chloride (3.9 ml, 50 mmol) was slowly added to maintain the temperature below 10  C. The mixture was stirred at ambient temperature during 3 h, whereupon no starting material could be detected (TLC: DCM/MeOH 9/1). The reaction medium was diluted with dichloromethane (100 ml) and washed successively with a 10% citric acid solution (3  10 ml), water, and sat. NaCl, then dried over sodium sulfate. After filtration and evaporation, the light brown residue solidified on standing at 4  C (7 g, 97%). Mp 85  C. TLC (Rf ¼ 0.84; DCM/MeOH 9:1; ninhydrin); 1H NMR (400 MHz, CDCl3, δ ppm): 1.43 (s, 9H, CH3), 3.06 (s, 6H, CH3), 4.23–4.37 (m, 5H, CH, CH2OH), 5.07 (d, 1H, J ¼ 7.6 Hz, NH).

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Scheme 3 (i), Boc2O, EtOH; (ii), CH3SO2Cl, Et3N, DCM; (iii), NaN3, DMF, 60  C; (iv), TFA / H2O (9:1),; (v), CS2, DCC, THF; (vi), glycolic anhydride, DCM; (vii), BOP,DIEA,DCM; (viii), H2, Pd/C, EtOH; (ix), 16 + 19, TEA,THF; (x), 1 N HCl, CH3CN 3.2.3 Tert-Butyl N-[2-Azido-1(Azidomethyl)Ethyl] Carbamate (14)

Compound 13 (8 g, 23.03 mmol) and sodium azide (14.97 g, 230 mmol) were placed in a 250 ml round bottom flask, DMF (50 ml) was added, and the mixture was stirred at 60  C for 4 h. A small aliquot was evaporated and controlled by TLC (DCM/MeOH 9:1). The reaction medium was evaporated under reduced pressure; the residue dissolved in dichloromethane and washed with water (3  25 ml), sat. NaCl (2  25 ml) and dried over Na2SO4. Evaporation of the solvent gave a yellow liquid (3.42 g; 62%). TLC (Rf ¼ 0.95; DCM/MeOH 9:1; ninhydrin); 1 H NMR (400 MHz, CDCl3, δ ppm): 1.43 (s, 9H, CH3), 3.40 (dd, J ¼ 7.8 Hz, J ¼ 16.6 Hz, CH2), 3.51 (dd, J ¼ 6.8 Hz, J ¼ 16.6 Hz, CH2), 3.85 (m, 1H, CH), 4.91 (d, 1H, J ¼ 8.4 Hz, NH).

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3.2.4 1,3-Diazidopropan2-Amine (15)

Compound 14 (3.37 g, 13.97 mmol) was dissolved at 0  C in a mixture of trifluoroacetic acid (18 ml) and water (2 ml). After 15 min it was evaporated to dryness and co-evaporated with cyclohexane three times. The residue was washed several times with ether until no acidity remained. (1.8 g; 91%). TLC (Rf ¼ 0.78; DCM/MeOH 9:1; ninhydrin); 1H NMR (400 MHz, CDCl3, δ ppm): 1.45 (s, 2H, NH), 3.02 (p, 1H, J ¼ 5.8 Hz, CH), 3.32 (dd, 2H, J ¼ 5.8 Hz, J ¼ 12.0 Hz, CH2), 3.40 (dd, 2H, J ¼ 5.8 Hz, J ¼ 12.0 Hz, CH2). MS (EI DCI) 142 (M+H)+.

3.2.5 4-(Isothiocyanatomethyl)-2,2Dimethyl-1,3-Dioxolane (16)

2,2-dimethyl-1,3-dioxolan-4-methanamine (1 g, 7.62 mmol) and carbon disulfide (0.46 ml, 7.62 mmol) were dissolved in THF (30 ml), DCC (1.57 g, 7.62 mmol) was added and the mixture was stirred at room temperature for 1 h (see Note 5). The reaction medium was diluted with pentane, filtered and the filtrate was washed successively with 0.1 N HCl, water, and sat. NaCl solution. The organic phase was dried over sodium sulfate, filtered and concentrated to the pure product (1 g, 75%) (see Note 6). TLC (Rf ¼ 0.27; DCM/MeOH 98:2; I2/H2SO4); 1H NMR (400 MHz, CDCl3, δ ppm): 1.31 (s, 3H, CH3), 1.42 (s, 3H, CH3), 3.54 (dd, 1H, J ¼ 5.1 Hz, J ¼ 15.0 Hz, CH2NCS), 3.64 (dd, 1H, J ¼ 5.1 Hz, J ¼ 15.0 Hz, CH2NCS), 3.80 (dd, J ¼ 5.1 Hz, J ¼ 8.7 Hz, CH2O), 4.07 (dd, J ¼ 6.3 Hz, J ¼ 8.7 Hz, CH2O), 4.26 (q, 1H, J) 5.3 Hz, CHO). MS (ESI, m/z) 174 (M+H)+.

3.2.6 2-[2-(Didecylamino)-2-Oxo-Ethoxy] Acetic Acid (17)

Didecylamine (3 g, 10.08 mmol) was placed in a 150 ml roundbottom flask and dissolved in DCM (100 ml), glycolic anhydride (1.64 g, 14.1 mmol) was added and the mixture was stirred at room temperature for 2 h. The end of the reaction was controlled by TLC (DCM/MeOH 9/1), the mixture was concentrated, the residue was taken up in ethyl acetate and washed successively with 0.1 N HCl (2), sat. NaCl and dried over Na2SO4. Evaporation of the solvent gave the title compound as an oil which crystallized on standing (4 g, 96%). Mp 55  C. TLC (Rf ¼ 0.47; DCM/MeOH 9:1; I2/H2SO4); 1H NMR (400 MHz, CDCl3, δ ppm): 0.85 (t, 6H, J ¼ 6.0 Hz, CH3), 1.25 (m, 28H, –CH2-), 1.53 (m, 4H, -CH2-), 3.08 (t, 2H, J ¼ 7.3 Hz, –CH2N), 3.33 (t, 2H, J ¼ 7.3 Hz, –CH2N), 4.19 (s, 2H, CH2O), 4.39 (s, 2H, CH2O). MS (ESI, m/z) 414 (M+H)+.

3.2.7 N-[2-Azido-1(Azidomethyl)Ethyl]-2[2-(Didecylamino)-2-OxoEthoxy]acetamide (18)

In a 100 ml round-bottom flask compound 17 (3.21 g, 7.76 mmol) and compound 15 (1.97 g, 7.76 mmol) were dissolved in dichloromethane (40 ml), BOP (3.43 g, 7.76 mmol) was added. The mixture was cooled to 0  C in an ice-water bath, triethylamine (2 ml, 14.23 mmol) was added dropwise. and the mixture was stirred for 1 h at room temperature (see Note 7). The solution

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was diluted with DCM (100 ml) and successively washed with 0.5 M KHSO4 (3  20 ml), sat. NaHCO3 (3  20 ml), water, and sat. NaCl solution. It was dried over Na2SO4, filtered and concentrated, yielding 18 (4 g, 96%). TLC (Rf ¼ 0.24; DCM/MeOH 99:1; I2/H2SO4); 1H NMR (400 MHz, CDCl3, δ ppm): 0.88 (t, 6H, J ¼ 6.0 Hz, CH3), 1.25 (m, 28H, -CH2-), 1.51 (m, 4H, -CH2-), 3.05 (t, 2H, J ¼ 7.3 Hz, -CH2N), 3.31 (t, 2H, J ¼ 7.3 Hz, CH2N), 3.47 (dd, 2H, J ¼ 6.0 Hz, CH2N3), 3.55 (dd, 2H, J ¼ 6.0 Hz, CH2N3), 4.10 (s, 2H, CH2O), 4.22 (m, 1H, CH), 4.27 (s, 2H, CH2O), 8.77 (d, 1H, J ¼ 8.4 Hz, NH). MS (ESI, m/z) 537 (M+H)+, 559 (M+Na)+. 3.2.8 N-[2-Amino-1(Aminomethyl)Ethyl]-2[2-(Didecylamino)-2-OxoEthoxy]Acetamide (19)

Compound 18 (2.74 g, 5.1 mmol) was dissolved in ethanol (50 ml) and cooled in an ice-water bath (see Note 8), then 10% palladium on carbon (0.2 g) was added. The mixture was stirred at atmospheric pressure of hydrogen during 3 h (see Note 9). The solution was filtered on a sintered glass funnel (n 4) coated with Celite (see Note 10) and concentrated under vacuum to give 19 (2.45 g, 99%). TLC (Rf ¼ 0.05; DCM/MeOH 8:2; ninhydrin or I2/ H2SO4); 1H NMR (400 MHz, CDCl3, δ ppm): 0.84 (t, 6H, J ¼ 6.0 Hz, CH3), 1.25 (m, 28H, –CH2–), 1.48 (m, 4H, –CH2–), 1.77 (m, 4H, NH2), 2.54 (dd, 1H, J ¼ 6.0 Hz, J ¼ 12.0 Hz, CH2NH2), 2.71 (dd, 1H, J ¼ 5.8 Hz, J ¼ 12.6 Hz, CH2NH2), 3.05 (t, 2H, J ¼ 7.3 Hz, -CH2N), 3.26 (m, 3H, -CH2N, CH), 3.39 (s, 2H, CH2O), 4.05 (s, 2H, CH2O), 8.20 (m, 1H, NH).

3.2.9 2-[2-(Didecylamino)-2-Oxo-Ethoxy]-N[2-[(2,2-Dimethyl-1,3Dioxolan-4-yl)Methylcarbamothioylamino]-1[[(2,2-Dimethyl-1,3Dioxolan-4-yl)Methylcarbamothioylamino] Methyl]Ethyl]Acetamide (20)

In a 100 ml round-bottom flask, compounds 19 (2.17 g, 4.48 mmol) and 16 (1.63 g, 9.4 mmol) were dissolved in DCM (45 ml) and triethylamine was added dropwise. The solution was stirred overnight at room temperature. It was washed with water and sat. NaCl solution, dried over sodium sulfate, filtered and concentrated. The crude product was chromatographed on silica gel with a gradient of cyclohexane/ethanol (0 ! 40%, 60 min). The fractions containing compound 20 were evaporated to a clear syrup (2 g, 54%). TLC (Rf ¼ 0.6; DCM/MeOH 95:5; I2/ H2SO4); 1H NMR (400 MHz, CDCl3, δ ppm): 0.85 (t, 6H, J) 6.6 Hz, CH3), 1.23 (m, 28H, -CH2-), 1.30 (s, 3H, CH3), 1.39 (s, 3H, CH3), 1.49 (m, 4H, -CH2-), 3.06 (t, 2H, J) 7.2 Hz, -CH2NCO), 3.27 (t, 2H, J) 7.2 Hz, -CH2-NCO), 3.68 (m, 8H, CH2NCS), 4.04 (m, 6H, CH2O), 4.25 (m, 7H, CH2O, CH, CHO). MS (ESI, m/z) 829 (MH), 831 (M–H+), 853 (MNa+).

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3.2.10 2-[2-(Didecylamino)-2-Oxo-Ethoxy]-N– [2-(2,3-Dihydroxypropylcarbamothioylamino)-1[(2,3-Dihydroxypropylcarbamothioylamino)Methyl]Ethyl]Acet amide (21; DDSTU)

Compound 20 (0.3 g, 0.36 mmol) was dissolved in acetonitrile (10 ml) and 1 N HCl was added (5 ml). The mixture was vigorously stirred for 1 h at room temperature, and then evaporated to dryness under high vacuum without heating. The residue was redissolved in DCM, washed with sat. NaCl solution, dried over sodium sulfate, filtered and concentrated, then purified on a silica column eluted first with DCM/MeOH (95:5), then DCM/MeOH (8:2). Concentration of the eluate gave 21 (0.2 g, 57%). TLC (Rf ¼ 0.17; DCM/MeOH 9:1; UV, I2/H2SO4); 1H NMR (400 MHz, CDCl3, δ ppm): 0.88 (t, 6H, J) 6.0 Hz, CH3), 1.23 (m, 28H, -CH2-), 1.54 (m, 4H, -CH2-), 2.14 (s, 4H, OH), 3.09 (m, 2H, CH2NCO), 3.30 (m, 2H, CH2NCO), 3.63 (m, 8H, CH2NCS), 3.92 (m, 1H, CH), 4.13 (m, 6H, CH2O, CH2OH), 4.31 (m, 7H, CH2O, CHOH), 7.50 (m, 5H, NH). HR-ESMS calc for C35H70N6O7NaS2: 773.4645. Found 773.4652. MS (ESI, m/z) 751 (M); 752 (MH+).

3.3

Lipoplexes are obtained by preformation of liposomes followed by a simple mixing with the nucleic acid of interest. All the methods are pretty straightforward. Ethanolic injection requires only two steps which are obviously crucial, the volume of dispersion should be well defined, and dropping the dissolved lipids in an aqueous medium should be regularly performed. Liposomes made out of DMAPAP and DOPE are either prepared as a film which protocol has been previously described [8], or as an ethanolic injection which is described underneath. The cationic formulation is the sole formulation described, but noncationic liposomes made of thiourea lipids are also prepared following this protocol. Actually the main difficulty lies in the first step following the synthesis. How to suspend a newly synthesized lipid and how to formulate it? Does it need another lipid to be suspended? Basically, amphiphilic lipids soluble in ethanol could be prepared by the protocol of ethanolic injection described underneath. However, they are not all soluble in ethanol. Here are the main points to handle a lipid:

Formulation

1. Lipids are very often hygroscopic molecules, so take care of leaving them in a dry environment; do not leave them to hydrate for hours on your bench. 2. Look at the structure: if the lipid bears double bonds, specific conditions of storage should be used such as nitrogen conditioning, to avoid oxidation. 3. Start with solubility studies: Weight several flasks containing 1 mg of the lipid and dilute it in ethanol, in acetone, in chloroform. If 1 mg is solubilized in less than 100 μl of EtOH, then the ethanolic injection is appropriate (see Note 11). Solubilization of the lipid in CHCl3 indicates that

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the film method is more appropriate for liposome obtention. For a complete description see ref. 10. 3.4 Preparation of Liposomes by the Ethanolic Injection Method

1. Dissolve separately the lipids DMAPAP (10 μmol, 10 mg) and DOPE (10 μmol, 7.3 mg) in ethanol (limit of solubility for each lipid). Take care that the lipids are well dissolved separately before mixing them (see Note 12). 2. Mix them into an eppendorf. 3. Note the total volume of ethanol required to perfectly solubilize both lipids. 4. Put milliQ filtered (0.22 μm) water in a round bottom flask (see Note 13). The volume should be ten times the amount of organic solvent required to solubilize the lipids. Put the flask on a magnetic stirrer with a magnetic bar of an appropriate size. Check that the magnetic bar turns properly, fast, and continuously (see Note 14). 5. Drop the solubilized lipids on the stirring water (see Note 15). Leave the mixture for 5 h. 6. Remove the solvent with a rotary evaporator with a pressure control device. Be careful that the suspension does not foam (see Note 16). 7. Remove the flask from the evaporator when the suspension approximately reaches the volume you expect (an approximate 10–30 mM final concentration). 8. Pipet with a hand pipette or a syringe to determine the volume left in the flask. 9. Calculate the concentration of the lipid in your suspension according to the amount initially weighed, in the predetermined volume left. 10. Control the size by dynamic light scattering (see Note 13 and Subheading 3.6.2 for more details). For measurements on a nanoZS (Malvern Instruments), dilute 5 μl of the particles obtained in a 500 μl cuvette, start the measure in the automatic mode.

3.5 Preparation of Cationic Lipoplexes

To obtain lipoplexes containing 1 μg of DNA in which DNA is fully associated, use a charge ratio of 4:8 (see Note 17). The charge ratio represents a molar ratio of cationic lipid to phosphate functions. The protocol below is given for a charge ratio lipid/DNA ¼ 6, which corresponds to a ratio total lipid to DNA ¼ 12 as the cationic lipid only represents 50% of the total lipid content in the DMAPA/ DOPE mixture. 1. Dilute the DMAPAP/DOPE suspension to 1 mM total lipid in H2O (see Note 18). 2. Dilute 1 μg plasmid DNA in 100 μl H2O.

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3. Dilute 12 μl of the 1 mM DMAPAP/DOPE suspension in 100 μl H2O. 4. Add the plasmid DNA to the cationic liposome dropwise in few seconds with constant vortexing (see Note 19). 5. Leave the sample for 1 h at room temperature to incubate before using it. The preparation of thiourea lipoplexes is similar to the preparation of cationic lipoplexes. The difference lies in the amount of lipid required to condensate DNA. The interaction between thiourea lipid and phosphates occurs at a ratio of 1 as shown by fluorescence correlation spectroscopy [11], however, these lipids do protect DNA and transfect cells at a higher TU/PO ratio, which is why the lipoplexes are prepared at a ratio lipid/phosphate ¼ 40. The complexes are usually prepared and left few hours before use. 3.6 Lipoplex Characterization

1. Prepare the picogreen® solution as described by the provider (1/200 in tris-EDTA buffer).

3.6.1 DNA Complexation Checked by Fluorescence

2. Load into a 96-well plate free DNA or complexed DNA (40 ng) in triplicate. 3. Add 200 μl of the picogreen solution (Subheading 3.6.1) to each well filled with DNA and three more to obtain the picogreen background level. 4. Read the emission at 450 nm under an excitation at 350 nm on a multiplate reader able to measure fluorescence. 5. For the calculation, calculate the mean and the standard error on each triplicate. Remove the picogreen background from the sample data. Calculate the percentage of fluorescence of each sample by dividing the sample data by the value of the free DNA taken as 100% fluorescence.

3.6.2 Lipoplex Size and Zeta Potential

The hydrodynamic diameter of the particles can be measured by quasi-elastic light scattering. The particles in suspension are submitted to the Brownian movement. When the particles are under a laser beam, they scatter the light in every direction. The variations of the light intensity as a function of time indicates the particle speed, which can be linked to their diameter by the Stokes-Einstein equation: D ¼ kT/6π Rη where D is the particle scattering coefficient, T the temperature, K the Boltzmann constant, R the particle radius, and η the viscosity of the solvent. The zeta potential is obtained through the measurement by the same technique of the electrophoretic mobility. It means that the cuvette used needs to be equipped with electrodes in order to provide an electric field which is proportional to the electrophoretic mobility. The zeta potential is obtained using the Smoluchowski law ζ ¼ ημe/εrεo where ζ is the zeta potential, η, the viscosity, μe the

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Table 1 Lipoplex characterization using dynamic light scattering Lipid/POratio

Size by intensity Polydispersity (nm) index (PDI)

Zeta potential Conductivity (mV) (mS/cm)

DMAPAP/ DOPE

N/P 4

69.1
0.4

0.18
0.02

22
1

0.22

DDSTU

TU/P 40

74.5
2.0

0.13
0.02

8
4

0.23

electrophoretic mobility, εr the dielectric constant of the dispersing medium, and εo the permittivity of free space. 1. Take a sample of your preparation and measure the size. The amount to be used depends on the system you are equipped with (see Note 20). 2. Insure the reliability of the measure by assessing the autocorrelation function and the polydispersity index (see Note 21). 3. If the sample is sufficiently concentrated for size measurement, it should be possible to measure the zeta potential on a similar sample. However, a conductivity medium should be used such as 20 mM NaCl to provide ions displacement during the electrophoresis (see Note 22). An example of how the results should be presented is given in Table 1. 3.7 Lipoplex Stability in Culture Medium

Two criteria can be evaluated: – The particle stability toward serum in terms of particle size or protein association. – DNA release or protection toward enzymatic degradation.

3.7.1 Particle Stability

1. Take a sample of lipoplexes (10 μl, 10 times more concentrated than previously described as you would use for in vivo injection 0.1 g/l DNA). 2. Dilute it in 200 μl culture medium supplemented or not with 10% serum. 3. Increase the temperature of the DLS system to 37  C. 4. Take a measure of the particle size every 2 min at 37  C. 5. Trace the evolution of the particles in terms of size, polydispersity index, and counts as a function of time.

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1. Prepare the lipoplexes with a DNA concentration of 0.1 g/l as described above in Subheading 3.5. 2. To 50 μl of the lipoplex solution, add 50 μl of culture medium implemented with 10–50% of murine fresh serum (see Note 23). 3. Incubate the samples at 37  C. 4. Every hour or so, take 10 μl sample and freeze it at 20  C. 5. After 24 h, take all the samples out of the freezer, add 2% SDS (sodium dodecyl sulfate) (5 μl), EDTA (2 μl, 0.5 M), and bromophenol blue (3 μl) to each sample (10 μl). 6. Load the mixture onto 1% agarose gel containing 0.05% SDS and put under 80 V/cm voltage. 7. After 24 h of rinsing the gel in water, plunge the gel in a solution of ethidium bromide and visualize it under UV light to reveal DNA. For an example see the Fig. 1 in ref. 6.

4

Notes 1. It should be noted that the synthetic routes have been designed to permit the syntheses of numerous structural analogs as well. The syntheses of compounds 1 to 7 were already described in a previous issue of Methods in Molecular Biology [12]. 2. A mixture of rotamers was observed by the doubling of some NMR signals. 3. Partial hydrolysis of the ethyl trifluoroacetate generates trifluoroacetic acid: It is mandatory that the pH of the ester is neutral, otherwise the amine would protonate and fail to react. The ester should be treated with dry sodium (or potassium) carbonate prior using (CAUTION: CO2 gas release). 4. The crude residue was dissolved in a warm mixture of isopropanol (600 ml) and a 5 M HCl solution in isopropanol (300 ml), which induced crystallization of the product as a white flaky powder. It was thoroughly washed with isopropanol and dichloromethane and dried. 5. After 1 h the pH of the solution was neutral, which attested the total consumption of the amine and the end of the reaction. 6. The product is evaporable and therefore temperature and duration of evaporation should be controlled. 7. pH should be >8, otherwise more Et3N has to be added. 8. The solution should be cooled in an ice-water bath before adding the catalyst, otherwise the methanol vapors may catch fire on contact with palladium.

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9. The end of reaction could be simply monitored by sampling the solution and observe the disappearance of the azide band in IR. 10. Care should be taken not to trash the dry catalyst to avoid possible fire. Instead it should be wetted and disposed of properly. 11. If not, try to heat it or sonicate it. Additional energy might help, but pay attention to an eventual reversible process. If sonication of the solution leads to a suspension of the lipid (cloudy solution), then try another solvent, first in acetone 50 μl for instance, then dilute it in EtOH. 12. Solubility of the lipids should be checked with intensive care since presence of nonsoluble entities will reduce particle homogeneity after evaporation and might cause aggregation. A lipid used at the limit of its solubility might precipitate directly when dropped on H2O or during the evaporation and volume reduction. In this case, a colipid bearing a larger hydrophilic head might be required to insure a correct interaction with the aqueous buffer and maintain the colloidal stability of the suspension. 13. All buffers and water used should be filtered on 0.22 μm filters since any dust might interfer with light scattering experiments. 14. Nonhomogeneous stirring would lead to a polydisperse population of liposomes. 15. Dropping can be performed via a peristaltic pump for a better homogeneity of the dispersion. 16. Formation of a suspension of micelles in the formulation could lead to a foam during the evaporation process, reduce the pressure cautiously. 17. The ratio between the lipid amines and DNA phosphates is fully dependent on the amine substitution and the conditions used [10]. 18. The protocol is described with H2O but can be changed for NaCl 150 mM or cellular medium. Obviously, the buffer will influence the aggregation state of the lipoplexes. Basically, all ions which will interact with the charges will reduce they availability for the interaction and enlarge the range of aggregation [13]. 19. In order to maintain an excess of cationic charges and hence avoid precipitation by going through a charge ratio (+/) equal to 1, DNA should be added on the cationic lipid and not the opposite order. 20. The concentration to be used depends on the sensitivity of the system and the angle used to detect the sample. The case of multiple diffusion is rare as usually the samples are not too

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concentrated. If you are equipped with a Zeta Sizer NanoSeries Malvern (Malvern Instruments, Venissieux, France). The concentration of the samples can be approximately 0.1 mg/ml in H2O. 21. Pay attention to the data obtained. Very often, a value will be given without reproducibility. The value is reliable if the polydispersity index is below 0.2, and if the data obtained in terms of intensity, volume, and number are identical and reproducible. Basically if the peak moves between three measurements and does not give similar results between volume, number and intensity, you more than probably have a polydisperse sample which gives different results due to the low volume of measurement. 22. Reminder: only samples exhibiting similar conductivity and pH can be compared in terms of zeta potential value. 23. Serum should be fresh to insure a strong enzymatic activity. Serum obtained from mice is more active than lyophilized serum from Sigma and more active than repetitively frozen sera. References 1. Bangham A, Standish M, Watkins J (1965) Diffusion of univalent ions across the lamellae of swollen phospholipids. J Mol Biol 13:238–252 2. Nicolazzi C, Garinot M, Mignet N, Scherman D, Bessodes M (2003) Cationic lipids for transfection. Curr Med Chem 10:1263–1277 3. Byk G, Wetzer B, Frederic M, Dubertret C, Pitard B, Jaslin G, Scherman D (2000) Reduction-sensitive lipopolyamines as a novel nonviral gene delivery system for modulated release of DNA with improved transgene expression. J Med Chem 43:4377–4387 4. Tranchant I, Thompson B, Nicolazzi C, Mignet N, Scherman D (2004) Physicochemical optimization of plasmid delivery by cationic lipids. J Gene Med 6(Suppl. 1):S24–S35 5. Tranchant I, Mignet N, Crozat E, Chain J, Girard C, Scherman D, Herscovici J (2004) DNA complexing lipopolythiourea. Bioconjug Chem 15:1342–1348 6. Leblond J, Mignet N, Largeau C, Seguin J, Scherman D, Herscovici J (2008) Lipopolythiourea transfecting agents: lysine thiourea derivatives. Bioconjug Chem 19:306–314 7. Breton M, Leblond J, Seguin J, Midoux P, Scherman D, Herscovici J, Pichon C, Mignet N (2010) Comparative gene transfer between cationic and thiourea lipoplexes. J Gene Med 12:45–54

8. Thompson B, Mignet N, Hofland H, Lamons D, Seguin J, Nicolazzi C, de la Figuera N, Kuen R, Meng Y, Scherman D, Bessodes M (2005) Neutral post-grafted colloidal particles for gene delivery. Bioconjug Chem 16:608–614 9. Leblond J, Mignet N, Largeau C, Spanedda MV, Seguin J, Scherman D, Herscovici J (2007) Lipopolythioureas: a new non-cationic system for gene transfer. Bioconjug Chem 18:484–493 10. Mignet N, Scherman D (2010) Anionic pH sensitive lipoplexes. In: Weissig V (ed) Methods in molecular biology, Liposomes, vol 605. Humana Press, Totowa, NJ, pp 435–444 11. Kral T, Leblond J, Hof M, Scherman D, Herscovici J, Mignet N (2010) Lipopolythiourea / DNA interaction: a biophysical study. Biophys Chem 148:68–73 12. Bessodes M, Scherman D (2010) Acid-labile liposome / pDNA complexes. In: Weissig V (ed) Methods in molecular biology, Liposomes, vol 605. Humana Press, Totowa, NJ, pp 405–423 13. Turek J, Dubertret C, Jaslin G, Antonakis K, Scherman D, Pitard B (2000) Formulations which increase the size of lipoplexes prevent serum-associated inhibition of transfection. J Gene Med 2:32–40

Chapter 9 Preparation, Characterization, and In Vitro Evaluation of Lipidoid–Polymer Hybrid Nanoparticles for siRNA Delivery to the Cytosol Kaushik Thanki, Xianghui Zeng, and Camilla Foged Abstract RNA interference (RNAi) therapeutics are one of the most promising biological interventions in the efficient management of difficult-to-treat diseases. RNAi is mediated by small interfering RNA (siRNA), which induces specific and highly potent gene silencing. However, intracellular delivery of exogenous, chemically synthesized siRNA to the RNAi pathway in the cytosol remains a challenge, and is fully dependent on technologies that can facilitate cytosolic delivery without undesired side effects. One example is a novel delivery system referred to as lipidoid–polymer hybrid nanoparticles (LPNs), which we recently showed mediates highly efficient and safe gene silencing. Here we describe a double emulsion solvent evaporation method for the preparation of siRNA-loaded LPNs and methodologies employed for their physicochemical characterization and biological performance. A solution of siRNA in aqueous buffer is emulsified by sonication with an organic phase containing lipid and polymer into a primary emulsion. Subsequently, the primary emulsion is emulsified with a secondary water phase containing polyvinyl alcohol by sonication, and the organic phase is evaporated, eventually resulting in LPNs. The physicochemical characterization includes determination of (1) hydrodynamic particle size distribution, (2) zeta potential, (3) siRNA encapsulation efficiency, and (4) practical siRNA loading. The transfection experiments are conducted in a cell-based model system using enhanced green fluorescence protein as reporter. The gene silencing effect is also confirmed at the mRNA level by reverse transcription polymerase chain reaction (RT-PCR). The effect of the siRNA-loaded LPNs on cell viability is measured using the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay. Key words Lipidoids, Poly(D,L-lactic-co-glycolic acid) (PLGA), Lipid–polymer hybrid nanoparticles, Small interfering RNA, Antisense oligonucleotides, Double emulsion solvent evaporation method

1

Introduction Therapeutics based on RNA interference (RNAi) have an enormous potential in the management of difficult-to-treat diseases wherein the genetic etiology is well known. Such an approach of sequence-based gene suppression is advantageous due to (1) high specificity, (2) superior potency, and (3) great versatility because

Manfred Ogris and Haider Sami (eds.), Nanotechnology for Nucleic Acid Delivery: Methods and Protocols, Methods in Molecular Biology, vol. 1943, https://doi.org/10.1007/978-1-4939-9092-4_9, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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any gene in theory can be targeted [1]. Although the advantages are attractive, the approach is also associated with a number of challenges, and the delivery of RNAi-based therapeutics to the cytosol of cells is one of the major bottlenecks for their further exploitation. The combination of unfavorable physicochemical properties of various RNA-based cargoes [e.g., small interfering RNA (siRNA), microRNA (miRNA), short hairpin RNA (shRNA), and long noncoding RNA (lncRNA)] and physiological barriers often demands the use of delivery technologies that can mediate the delivery of loaded cargo into the cytosol, where it mediates efficient RNAi. A variety of different types of nanocarriers has been designed and explored with the holistic goal of high transfection efficiency vis-a`-vis low side effects (e.g., cytotoxicity and immunogenicity) [2]. The pioneering work in this area included the use of viral vectors for intracellular delivery. However, because of safety issues, the paradigm in the field shifted to nonviral vectors. We recently demonstrated the potential of lipidoid–polymer hybrid nanoparticles (LPNs) for efficient and safe intracellular delivery of siRNA [3]. The LPNs comprise two main components: Lipidoid and poly(D,L-lactic-co-glycolic acid) (PLGA). Lipidoid belong to a novel class of cationic lipid-like materials [4], capable of efficiently interacting with polyanionic siRNA via attractive electrostatic interactions and subsequently mediating cellular internalization, endosomal escape, and cytosolic delivery. PLGA serves as a polymeric core component imparting sustained release properties to the nanoparticles, and it constitutes an integral part of the nanoparticle architecture. The cationic lipid component (i.e., lipidoid) interacts with the PLGA core and forms a shell membrane structure coating the core and binding the siRNA via attractive electrostatic interactions with the cationic headgroups. The preparation of such nanoparticulate formulations is usually complex. The aim of the present work is to specify a protocol highlighting critical contributing factors, including formulation parameters and process parameters. The LPNs are prepared by using a double emulsion solvent evaporation (DESE) method, enabling efficient encapsulation of water-soluble and polyanionic compounds (e.g., siRNA). The DESE method comprises of three essential steps: (1) formation of a primary emulsion, (2) phase inversion and stabilization of a secondary emulsion, and (3) size reduction and evaporation of the organic phase. The first step involves the formation of a primary water-in-oil (w/o) emulsion: An aqueous phase containing polyanionic siRNA is emulsified with an organic phase containing cationic lipidoid and hydrophobic PLGA. Due to the use of relatively small volumes of liquid, a probe sonication-based unit operation is employed for efficient mixing to maximize electrostatic and hydrophobic interactions between the components. In the second step,

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the metastable emulsion is stabilized by inclusion of the surfactant polyvinyl alcohol (PVA) in the continuous aqueous phase. The stabilization process mediated by phase inversion results in increased droplet size of the emulsion. Therefore, the resulting mixture is subjected to probe sonication to reduce the droplet size in the third step. Subsequently, the organic phase is evaporated, eventually resulting in an LPN dispersion.

2

Materials

2.1 Glassware and Plasticware

All glassware and plasticware used for the preparation of LPNs have to be free from RNAses. Glassware should be decontaminated by baking at 180
C for at least 10 h. RNAse-free plasticware can readily be procured commercially.

2.2

PLGA

PLGA with a lactide-to-glycolide molar ratio of 75:25, Mw 20 kDa and an ester type end-group modification was obtained from Wako Pure Chemical Industries (Osaka, JP).

2.3

Lipidoid 5

Mw 1343.25, synthesized, purified, and characterized as previously described [3]. This specific type of lipidoid used here has a tetraamine backbone displaying five acyl chains, and hence is referred to as lipidoid 5.

2.4 DEPC-Treated Water

0.1% (v/v) diethylpyrocarbonate (DEPC, Sigma-Aldrich, Saint Louis, MO, USA), in purified water (USP type 1 water, see Note 1 for additional details).

2.5

5 mM HEPES, pH 7.4, sodium hydroxide, hydrochloric acid (to adjust pH), in DEPC-treated water.

HEPES Buffer

2.6 siRNA Stock Solutions

20 -O-methyl-modified dicer substrate asymmetric siRNA duplexes directed against enhanced green fluorescent protein (EGFPsiRNA) and scrambled negative control were provided by Integrated DNA Technologies (IDT, Coralville, IA, USA) as dried, purified, and desalted duplexes). Weigh an amount of siRNA (the final siRNA concentration should be 1 mM, Table 1) in a glass vial and dissolve it in HEPES buffer (5 mM, pH 7.4) prepared in DEPC-treated water. Reanneal the siRNA duplex by heating the solution at 94
C for 2 min followed by gradual cooling to room temperature. Aliquot the reannealed solution into smaller volumes (maximum 100 μL) and store them at 20
C in microcentrifuge tubes (Nonstick, RNase-free Thermo Fisher Scientific, Hvidovre, DK).

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Table 1 siRNA sequences and modification patterns Name EGFP-siRNA

Target GenBank JQ064510.1

Negative control

Sense sequence 0

Antisense sequence

5 -pACCCUGAAGUUC AUCUGCACCACcg-30

50 -CGGUGGUGCAGAUGAA CUUCAGGGUCA-30

50 -AUCGUACGUACCGU CGUAUtt-30

50 -AUACGACGGUA CGUACGAUtt-30

Lowercase letters represent deoxyribonucleotides, underlined capital letters represent 20 -O-methylribonucleotides, and p phosphate residues

2.7 Tris-EDTA (TE) Buffer

10 mM Tris–HCl, 1 mM EDTA, pH 8.0 in DEPC-treated water. Adjust pH with sodium hydroxide or hydrochloric acid, if necessary.

2.8 Polyvinyl Alcohol (PVA) Solution

2% w/v PVA (87–90% hydrolyzed, average Mw 30–70 kDa, SigmaAldrich, Saint Louis, MO, USA) in DEPC-treated water (sterile filtered, see Note 2 for additional information).

2.9 HeparinDetergent (HD) Solution

Heparin (porcine intestinal mucosa, Grade I-A, 180 units/mg) 1 mg/mL, octyl ß-D-glucopyranoside (OG) 100 μM, dissolved in TE buffer.

2.10 Trehalose Solution

5% (w/v) trehalose dihydrate in DEPC-treated water, sterile filtered 0.22 μm syringe filter (see Note 3 for additional information).

3

Methods

3.1 Preparation of LPNs (Fig. 1)

1. Thaw lipidoid and PLGA (stored at 20 temperature.




C) at room

2. Weigh an amount of lipidoid (2.25 mg) and PLGA (12.75 mg) in an RNAse-free glass vial, and store the mixture on ice until use. A lipidoid content of 15% (w/w) to that of PLGA has been chosen for this protocol. 3. In a fume hood, add 250 μL of CH2Cl2 to the glass vial and gently dissolve the content by rotating the vial. 4. Prepare the primary emulsion by adding 125 μL of siRNA solution (8.3 μL of 1 mM siRNA stock solution (Mw 17951), and 116.7 μL TE buffer, referred to as the water phase, w1) to the organic phase (o), followed by probe sonication (Misonix, Qsonica, LLC., CT, USA) for 90 s at an amplitude of 50 in an ice bath to obtain the primary w1/o emulsion. A ratio of 1:15 (w/w) of siRNA:lipidoid has been chosen for this protocol.

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Fig. 1 Schematic diagram illustrating the preparation of LPNs

5. Add 1 mL of PVA solution (secondary water phase, w2) and vortex the resultant mixture vigorously for 1 min to phaseinverse the primary emulsion and form the secondary w1/o/ w2 emulsion. 6. Probe-sonicate the secondary emulsion for 60 s at an amplitude of 50 (Misonix, Qsonica) in an ice bath. 7. Pour the content of the vial into an RNAse-free glass beaker containing a magnet, and place it on a magnetic stirrer. Rinse the vial twice with 2 mL of PVA solution, and add the 2  2 mL to the final nanoparticle dispersion. Add additional 1 mL of PVA solution to the nanoparticle dispersion. 8. Stir the dispersion for 45 min to ensure complete evaporation of CH2Cl2. 3.2 Purification of LPNs

1. After preparation, the LPN dispersion is purified by centrifugation to remove unencapsulated siRNA and PVA (see Notes 4 and 5 for additional information). 2. The total volume of nanoparticle dispersion is approximately 6 mL (the theoretical volume is 6 mL, but a small fraction of the dispersion is lost during handling of the liquids). 3. Transfer the prepared LPN dispersion into three thick-walled polycarbonate centrifugation tubes (dimensions 13  56 mm, capacity 3.2 mL, Catalogue no. 362305, Beckman Coulter Inc.). Each tube should contain approximately 2 mL. Ensure that each tube contains no more than 1.5–2 mL/tube. 4. Centrifuge the samples to sediment the nanoparticles using a gradient centrifugation method (i.e., 6000  g for 5 min, 12,000  g for 5 min, 21,000  g for 5 min, 34,000  g for 5 min, and 48,000  g for 10 min at 4
C) (Optima™ Max Ultracentrifuge, Beckman Coulter, CA, USA). The total centrifugation time is 30 min.

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5. When the centrifugation process is almost complete, be ready close to the centrifuge with (1) the empty glass beaker in which the formulation was prepared, (2) DEPC water, (3) 1 mL pipette, and (4) pipette tips. As soon as the centrifugation process is complete, increase the pressure in the centrifuge chamber, gently take out the rotor, and remove the tubes without disturbing the samples. Gently pour the supernatants of the samples from all three tubes into the beaker, and place the remaining last drops present on the wall of the centrifugation tubes onto tissue paper while the tubes are still placed upside down to ensure that all liquid is withdrawn. 6. Add 0.5 mL of DEPC water to all three tubes and vortex for 5 min. Bath-sonicate the centrifugation tubes in pulses with concomitant vortexing to fully redisperse the nanoparticle pellet. 7. When the pellets are completely redispersed (no visible aggregates), combine the contents of all three tubes into one tube, and rinse the two empty tubes one by one with 0.5 mL of DEPC water (transfer the 0.5 mL from the first tube to the second) and transfer the washing water to the main formulation. Pulsed vortexing can also be used during rinsing. The total volume of the formulation should now be 2 mL. 3.3 Physicochemical Characterization of LPNs

1. The intensity-weighted mean hydrodynamic diameter (z-average) and polydispersity index (PDI) of the prepared LPNs can be measured by dynamic light scattering using the photon correlation spectroscopy technique. 2. An aliquot of 25 μL LPN dispersion is diluted to 1 mL with DEPC-treated water (approximately 0.3 mg/mL) and is subjected to measurement using a Zetasizer Nano ZS (Malvern Instruments, Worcestershire, UK) equipped with a 633 nm laser and 173
detection optics at 25
C. 3. The PDI reflects the particle size distribution of the LPNs dispersion ranging from 0 for a monodisperse to 1.0 for an entirely heterogeneous dispersion. The PDI is calculated using the following formula:  PDI ¼

peak width, d: nm peak height, d: nm

1=2 ð1Þ

4. Laser-Doppler microelectrophoresis is employed for measuring the zeta potential of the LPNs. 5. The Zetasizer Software version 7.11 (Malvern Instruments) is used for data acquisition and analysis. 6. The siRNA entrapment efficiency of the LPNs is determined as follows: To a volume of 25 μL LPN dispersion, a volume of

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200 μL of CHCl3 is added, followed by addition of a volume of 475 μL of HD solution. The resultant mixture is rotated endover-end for 5 min for complete extraction of siRNA into the aqueous phase. 7. The two phases are separated by centrifugation at 22,000  g for 12 min at 4
C. 8. The aqueous phase (50 μL) is carefully transferred to a new microcentrifuge tube, diluted with 950 μL DEPC treated water and incubated at 37
C to evaporate residual CHCl3. 9. The concentration of siRNA in the aqueous phase is determined using the Quant-iT™ RiboGreen™ RNA Assay Kit following the manufacturer’s protocol using a fluorescence plate reader (FLUOstar OPTIMA, BMG Labtech, DE) with excitation and emission wavelengths of 485 and 520 nm, respectively. 10. The encapsulation efficiency and practical loading of the siRNA in the LPNs are calculated according to Eqs. 2 and 3: Encapsulation efficiency ¼

Practical loading ¼

Amount of encapsulated siRNA  100 Total amount of added siRNA ð2Þ

Amount of encapsulated siRNA  100 ð3Þ Total weight of nanoparticles

11. The encapsulation efficiency of lipidoid in the LPNs is determined as described below: The organic phase from step 5 is carefully transferred to HPLC vials, evaporated under nitrogen flush and reconstituted in 100 μL CH3CN/CHCl3 (1:1, v/v). 12. The concentration of lipidoid in the organic phase is measured using HPLC in conjunction with an evaporative light scattering detector (ELSD, (Agilent 1260 Infinity, Santa Clara, CA, USA) employing the following conditions: Mobile phase: CH3CN–H2O (95:5, v/v) and CH3CN–H2O (5:95, v/v). Supplementation: 0.1% (v/v) trifluoroacetic acid to both mobile phases. Flow rate: 1 mL/min. Injection volume: 30 μL. Column: Luna C18 (150 mm  4.6 mm i.d., 3 μm particle size, ˚ pore size, Phenomenex, Torrance, CA, USA). 100 A Column temperature: 50
C. Gradient method: see Table 2.

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Table 2 Composition of mobile phases and gradient method Time (min)

CH3CN–H2O (5:95)a (%)

CH3CN–H2O (95:5)a (%)

0

50

50

3

0

100

12

0

100

12.1

100

0

14

100

0

15

50

50

17

50

50

a

Supplemented with 0.1% v/v trifluoroacetic acid

ELSD configuration: Nebulizer temperature: 50
C, Evaporator temperature: 75
C, Nitrogen gas flow rate: 1.5 standard lit/min. Data acquisition and processing: Agilent ChemStation software. 13. The encapsulation efficiency and practical loading of lipidoid in the LPN dispersion are calculated using Eqs. 2 and 3. 3.4 In Vitro Testing of the LPNs

1. The in vitro gene silencing of the prepared LPNs loaded with EGFP-siRNA is determined in the following way: The human non-small lung carcinoma cell line H1299 stably transfected with EGFP (EGFP-H1299) is maintained in RPMI 1640 medium containing 10% (v/v) fetal bovine serum (FBS, Gibco, Grand Island, NY, USA) at 37
C and 95/5% O2/CO2. 2. The cells are seeded in 24-well tissue culture plates (Corning, Corning, NY, USA) at a density of 1  105 cells/well and are allowed to adhere overnight. 3. The culture medium is aspirated and replaced with new medium (900 μL) along with the test formulations (10, 100 μL) in a concentration range (e.g., using fivefold dilutions and 6–8 different concentrations), followed by a 24 h incubation period. 4. The cells are then washed with phosphate-buffered saline (PBS, pH 7.4, Sigma-Aldrich) and reincubated with new culture medium for additional 24 h. 5. The cell culture medium is again aspirated, the cells are washed with 1 mL PBS and trypsinized using 300 μL trypsin–EDTA solution (10).

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6. Gene silencing is measured by quantifying (1) the EGFP expression by flow cytometry and (2) the EGFP mRNA levels by real time polymerase chain reaction (RT-PCR). 7. For protein quantification, transfer the trypsinized cells into FACS tubes and centrifuge them at 1000  g for 5 min. Gently pour off the supernatant and place the remaining last drop present on the wall of the tube onto tissue paper while the tube is still placed upside down to ensure that all liquid is withdrawn. Resuspend the pellet in 300 μL PBS, pH 7.4. Vortex briefly and measure the EGFP fluorescence of the cells in the green channel of a flow cytometer (Gallios, Beckman Coulter, Brea, CA, USA). Data is analyzed using the FlowJo software version 10 (Three Star, Ashland, OR, USA). The concentration corresponding to 50% gene silencing is calculated using curve fitting algorithms with for example GraphPad Prism (GraphPad, La Jolla, CA, USA). Optionally, dead cells can be excluded by propidium iodide staining. 8. For quantification of mRNA expression, transfer the trypsinized cells into a microcentrifuge tube and spin down the cells at 1000  g for 5 min. Discard the supernatant and resuspend the pellet in 350 μL cell lysis buffer. Extract total RNA from the cell pellet employing the NucleoSpin® RNA Plus kit (Macherey-Nagel GmbH & Co., Du¨ren, DE) following the manufacturer’s protocol. 9. The concentration of total RNA is determined using a microvolume spectrophotometer (NanoDrop™ 2000c, Thermo Fisher Scientific, Wilmington, DE, USA, see Note 6 for additional details). 10. Reverse transcription of 1 μg total RNA is performed with a mix of oligo dT and random hexamer primers applying the iScript cDNA synthesis Kit (Bio-Rad Laboratories, Hercules, CA, USA).

Table 3 Sequences of primers used for PCR Gene

NCBI RefSeq

Forward primer

Reverse primer

Product length (bp)

GADPH

NM_002046.6

50 -AGGCTGGGGC TCATTTGCAGG-30

50 -CAGTTGGTGGT GCAGGAGGCA-30

148

EGFP

GQ404376.1

50 -TGCACGCCGT AGGTCAGGGT-30

50 -GACGGCGACGT AAACGGCCA-30

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11. Dilute the synthesized cDNA with PCR-grade water (1:10) and measure the amplification of diluted cDNA using the primers listed in Table 3. 12. Each PCR reaction comprises a total volume of 20 μL containing 5 μL of diluted cDNA, 0.5 μM forward and reverse primer, respectively, and 10 μL of LightCycler® 480 SYBR Green I Master (2, Roche, Basel, CH, see Note 7 for additional information). 13. The following PCR conditions can be applied, for example using a LightCycler® 480 (Roche): Initial denaturation at 95
C for 5 min. Number of cycles: 35. Denaturation at 95
C for 10 s. Annealing at 60
C for 10 s. Elongation at 72
C for 10 s. Melting curve analysis at 95
C for 5 s, 65
C for 1 min and continuous detection every 5
C until reaching a temperature of 97
C. 14. Perform crossing point (Cp) analysis using the LightCycler® 480 software v 1.5.0 (Roche) for each gene. 15. The specific knockdown of EGFP can be determined using the ΔΔCp method considering the Cp values of reference genes (here GAPDH) and that of the untreated samples [5]. It is preferable to use more than one reference gene for normalization purposes. 16. The effect of the LPNs on cell viability in vitro can be assessed as follows. 17. Seed the H1299 EGFP cells in 96-well plates at a density of 10,000 cells/well. On the following day, aspirate the cell culture medium from each well and add 180 μL of new medium. 18. Incubate the cells with a concentration range (e.g., using fivefold dilutions and 6–8 different concentrations) of LPNs for 24 h: A 10 concentration of the test dose of the formulation is prepared, and 20 μL is added to each well. 19. After 24 h, aspirate the medium containing the test samples, wash the cells once with PBS and add freshly prepared 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT, Sigma-Aldrich, 500 μg/mL in PBS, 150 μL/ well). 20. Reincubate the cells for 4 h at 37
C and 5% CO2 for formation of insoluble MTT formazan. 21. Excess MTT solution is carefully removed from each well using a pipette. Add 200 μL of DMSO to each well to solubilize the formazan.

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22. The cell viability is assessed by measuring the absorbance of dissolved formazan at 550 nm and normalized to the absorbance of formazan formed by control cells. 23. Experiments are performed in quadruplicate, and data is analyzed using GraphPad Prism. The concentration corresponding to 50% cell viability is calculated using curve fitting algorithms with for example GraphPad Prism.

4

Notes 1. DEPC is poorly miscible with water, and the mixture should be well shaken to fully mix DEPC. This must be followed by incubation for a longer period to inactivate RNases (37
C for at least 12 h). Subsequently, DEPC is removed by autoclaving (121
C, 1 h and 1 atm), which results in hydrolysis of DEPC to EtOH and CO2. DEPC is incompatible with certain types of plastic, and hence glass bottles (reagent bottle) should be used for preparing DEPC solutions. 2. The water solubility of PVA is approximately 50 mg/mL, which is almost four times lower than the concentration needed for LPN preparation. Hence, heating of the mixture is necessary. If large aggregates of PVA are formed in water, the mixture can also be bath-sonicated at elevated temperature (approximately 50–60
C). We have noticed that the filter material used for filtration of the PVA solution is important. For example, cellulose acetate adsorbs PVA during filtration, eventually resulting in rapid clogging of the filter and reduction of the final concentration of PVA in the solution. Polyethersulfonate-based membranes are preferred for filtration of PVA solutions. 3. Use the trehalose solution immediately after preparation. 4. Unencapsulated siRNA is removed from the formulation, and the amount of siRNA encapsulated in the LPNs is quantified. This serves the following purposes: (1) to understand the effect of formulation components on the siRNA encapsulation efficiency, (2) to clearly differentiate the effect of encapsulated siRNA from unencapsulated siRNA (e.g., in cell culture studies), (3) to establish a method for recovering the unencapsulated siRNA and reuse it in the perspective of large-scale industrial scale production, and (4) to ensure batch-to-batch reproducibility by using the encapsulation efficiency as a critical quality attribute of the formulation. 5. The concentration of PVA solution used for stabilization purposes is usually higher than minimum requisite in order ensure that excess stabilizer is always present to cope up with any small

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changes in process parameters or formulation components. However, PVA being surfactant can be toxic and hence any excess amounts of it should be removed from the final formulation. 6. The quality of the isolated RNA should also be determined. The ratio of A260 to A280 reflects the protein contamination in the sample. For an acceptable RNA purity, the ratio should be above 2. The ratio of A260 to A230 reflects the solvent contamination in the sample, and it should generally be in the range of 1.8–2.2. 7. The PCR reactions are usually performed as two technical and three or more biological replicates. The viscosity of the master mix differs a lot from the viscosity of the other reagents (primers, cDNA and PCR grade water). Hence, it is advisable to prepare n + 1 reactions in a separate tube (0.2 mL) and mix thoroughly, where n is the total number of technical replicates.

Acknowledgments We gratefully acknowledge the support from the Lundbeck Foundation—Denmark (Grant No. R219-2016-908 and R181-20143793) and the Novo Nordisk Foundation—Denmark (Grant No. NNF17OC0026526). Conflicts of interest: The authors declare no conflicts of interest. References 1. Uprichard SL (2005) The therapeutic potential of RNA interference. FEBS Lett 579 (26):5996–6007 2. Dizaj SM, Jafari S, Khosroushahi AY (2014) A sight on the current nanoparticle-based gene delivery vectors. Nanoscale Res Lett 9(1):252 3. Thanki K, Zeng X, Justesen S, Tejlmann S, Falkenberg E, Van Driessche E, Morck Nielsen H, Franzyk H, Foged C (2017) Engineering of small interfering RNA-loaded lipidoid-poly(DL-lactic-co-glycolic acid) hybrid nanoparticles for highly efficient and safe gene silencing: a quality by design-based approach. Eur J Pharm Biopharm 120:22–33 4. Akinc A, Zumbuehl A, Goldberg M, Leshchiner ES, Busini V, Hossain N, Bacallado SA, Nguyen

DN, Fuller J, Alvarez R, Borodovsky A, Borland T, Constien R, de Fougerolles A, Dorkin JR, Narayanannair Jayaprakash K, Jayaraman M, John M, Koteliansky V, Manoharan M, Nechev L, Qin J, Racie T, Raitcheva D, Rajeev KG, Sah DW, Soutschek J, Toudjarska I, Vornlocher HP, Zimmermann TS, Langer R, Anderson DG (2008) A combinatorial library of lipid-like materials for delivery of RNAi therapeutics. Nat Biotechnol 26 (5):561–569 5. Pfaffl MW (2001) A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Res 29(9):e45

Chapter 10 Layer-by-Layer Assembled Nanoparticles for siRNA Delivery Michaela Guter and Miriam Breunig Abstract Nanoparticles synthesized via layer-by-layer processes are promising candidates for successful drug and gene delivery. Widespread use of the layer-by-layer technique has resulted from its accessibility to every lab; to generate nanoscale structures, layer-by-layer processes require common lab equipment of only modest quality and do not involve the use of organic solvents. In addition, a wide range of different starting materials can be flexibly combined, enabling the production of a nearly unlimited number of different nanoparticles (NP) with various physicochemical properties. Here we describe the manufacturing of poly (lactic-co-glycolic acid) NPs coated with siRNA for gene silencing. Positively charged polyethyleneimine and negatively charged nucleic acids form the polyelectrolyte shell. Finally, the NPs are functionalized with hyaluronic acid, a polysaccharide which targets the CD44 receptor. Key words siRNA delivery, Layer-by-layer, PLGA nanoparticles, Polyethyleneimine, Hyaluronic acid

1

Introduction “Layer-by-layer” (LbL) is a term used to describe film formation by depositing oppositely charged materials on a surface [1]. Decher was the first to characterize the underlying mechanism in 1991, assembling a 35-layer film, 170 nm in thickness, composed of alternating anionic and cationic bipolar amphiphiles on a planar surface [2]. Since then, the field has developed immensely. LbL techniques are now routinely leveraged for applications in material science, physical chemistry, electrochemistry, and biomedical engineering [1]. In the fields of drug and gene delivery, LbL shell growth on nanoscale templates has emerged as an area of particular interest [3]. The structure of LbL-coated NPs allows for the inclusion of therapeutics (e.g., small molecules or macromolecules like proteins and nucleic acids) into either the multilayer shell or the NP core. The technique is even suitable for extremely sensitive or labile molecules like RNA, as the layering process is performed under mild conditions without the need for harsh pH or elevated temperatures. The surface properties of the fully assembled NP can be

Manfred Ogris and Haider Sami (eds.), Nanotechnology for Nucleic Acid Delivery: Methods and Protocols, Methods in Molecular Biology, vol. 1943, https://doi.org/10.1007/978-1-4939-9092-4_10, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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tailored on a molecular level without significantly altering the shape and size of the particle as a whole, simply by selecting a suitable charged substance for the final layer. Substances with targeting functions can readily be attached to the nanoparticle surface by similar means. Additionally, inclusion of pH-sheddable, redox-sensitive or enzymatically degradable materials enable spatially and temporally controlled or triggered release [4]. Despite these significant advantages, high-quality LbL assembly is quite difficult, impeding broad application of the technique [5]. Of utmost importance is the complete removal of unbound polyelectrolyte after each layering cycle. Any oppositely charged polymers remaining in solution after a cycle will form complexes which are difficult to separate from coated particles. Purification processes must be thorough but gentle, as irreversible aggregation of particles poses a serious issue. Commonly applied strategies are repeated centrifugation and ultrafiltration. We developed a LbL-based protocol for manufacturing poly (lactic-co-glycolic acid) (PLGA) NPs for siRNA delivery. Figure 1 summarizes the composition of the particles. The NP core is composed of PLGA, an FDA-approved biodegradable polymer that is commercially available in high quality and a wide range of molecular weights. The synthesis of PLGA NPs is well established in the literature [6]. During synthesis of the PLGA core particle, we utilize polyethyleneimine (PEI) instead of poly(vinyl alcohol) as a stabilizing agent. As a well-known and highly effective transfection agent [7], PEI is intended to enhance the particles’ effectiveness as gene delivery vehicles. Further, PEI provides the NPs with the

Fig. 1 The LbL-based NP consists of a positively charged PLGA/PEI core and three subsequently deposited polyelectrolyte layers. The first layer consists of negatively charged siRNA. This is coated and protected by a layer of polycationic PEI. A final layer of anionic HA is then deposited to enable targeting of CD44 receptors

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cationic surface charge necessary for adsorption of an initial layer of negatively charged siRNA. To protect the siRNA from degradation, another layer of PEI was adsorbed on top of the siRNA. The negatively charged polysaccharide hyaluronic acid (HA) was used as a final, outermost layer. HA enhances the NP colloidal stability [8], increases NP mobility in the extracellular space [9], and serves as a targeting moiety [10]. As the main ligand of the CD44receptor, HA directs the NP to cells with high receptor density (e.g., tumor cells) [11] and others known to strongly interact with their extracellular matrix as for example in the trabecular meshwork [12]. References [13, 14] are excellent review articles covering this topic. The protocol presented here may be used as a general guide for LbL-based NP synthesis. However, if other materials, particle sizes, or polymer molecular weights are used, experimental conditions must be adapted to each individual case. This is due to the sensitivity of small particles to coating and purification conditions, wherein opportunities for irreversible aggregation are most often introduced.

2

Materials Strictly adhere to RNAse-free working conditions. Use a separate work space that has been treated with RNAse-digesting substances and avoid working in locations with turbulent airflow. Wear a lab coat and gloves at all times and change them on a regular basis. Exclusively use barrier tips and other RNAse-free consumables from unopened boxes and bags. Bake all glassware for at least 8 h at 200
C prior to use. Have an extra set of chemicals and reagents and use only RNAse-free equipment to remove them from their containers (see Note 1). Make sure to exclusively use RNAse-free water (e.g., from a certified reverse osmosis facility). Keep all solutions containing siRNA or siRNA-coated NPs on ice if possible.

2.1 Synthesis of PLGA NPs (PLGAPEI)

l

Branched poly(ethyleneimine) (PEI) 25 kDa: 0.5% (w/v) solution in water (see Note 2).

l

Acid terminated 38–54 kDa poly(D,L-lactide-co-gylcolide) (PLGA), lactide–glycolide 50:50: 10 mg/mL solution in acetonitrile (see Note 3).

l

Sodium chloride: 10 mM in water.

l

Ultrapure water.

l

Snap-cap vials, stir plate, stir bars.

l

2 mL Eppendorf tubes and a centrifuge (see Note 4).

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2.2 Freeze Drying of PLGA NPs

l

Sucrose: 4% (w/v) in water.

l

Lyophilization vials with plugs.

2.3 Coating with siRNA

l

2.4

Coating with PEI

2.5 Coating with Hyaluronic Acid

3

siRNA: 20 μM in water or buffer (follow manufacturer’s instructions).

l

Ultrapure water.

l

Snap-cap vials, stir plate, stir bars.

l

2 mL Eppendorf tubes and a centrifuge.

l

PEI: 100 mg/mL solution in water. Make sure that PEI is dissolved completely.

l

Ultrapure water.

l

Snap-cap vials, stir plate and stir bars.

l

2 mL Eppendorf tubes and centrifuge.

l

Sodium hyaluronate (HA), 13 kDa: 10 mg/mL in water (see Note 5).

l

Ultrapure water.

l

Snap-cap vials, stir plate and stir bars.

l

2 mL Eppendorf tubes and centrifuge.

Methods A schematic overview on synthesis and LbL coating procedures for PLGA NPs is given in Fig. 2.

3.1 Synthesis of PLGA NPs

Stir 8 mL of 0.5% PEI in a snap-cap vial to create a vortex and slowly inject 2 mL PLGA solution. When the solution becomes turbid, the NPs have formed. Stir for at least 4 h to guarantee the complete curing of the particles and evaporation of the organic solvent (see Note 6).

3.2

Split the NP dispersion into several Eppendorf tubes and centrifuge for 7 min at 4
C with a speed of 5000  g (see Note 4). Remove the supernatant. Resuspend the pellet in ultrapure water and centrifuge the supernatant again for 7 min at 4
C with a velocity of 7000  g. To increase yield, repeat the last step with a centrifugation speed of 9000  g (see Note 7). Discard the supernatant of the last step and combine the resuspended NPs. Repeat the entire process (see Note 8). If the purified NPs are to be coated with siRNA, perform the final resuspension in 10 mM sodium chloride.

NP Purification

LbL NP for siRNA Delivery

157

Fig. 2 Schematic illustration of the synthesis and LbL coating procedures for PLGA NPs. PLGA NPs are prepared by nanoprecipitation and freeze dried for further storage. Resuspended NPs are coated with siRNA, PEI and HA, in that order. Each coating step is followed by purification of NPs from excess polyelectrolyte 3.3 Freeze Drying and Storage

Pool the purified NPs, mix them with an equal volume of sucrose solution and fill them into tared lyophilization vials. Freeze the NPs at 80
C and then dry first for 5 days at 20
C and then for 2 days at þ20
C under vacuum (see Note 9). Weigh the dried samples and calculate the NP mass in the lyophilizate by subtracting the mass of sucrose added initially. Resuspend the NPs in ultrapure water to a concentration of 1 mg/mL and purify them as described above prior to coating.

3.4 Coating with Polyelectrolytes

Place the respective amount of polyelectrolyte solution into a snapcap vial equipped with a stir bar. Slowly add purified NPs dropwise into the gently stirred solution of polyelectrolyte and continue stirring for approximately 30 min at room temperature. The concentration of the polymers during coating is 1 mg/mL in water or 4 μM in 10 mM NaCl for polymers (PEI and HA) and siRNA respectively (see Notes 10–13).

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3.5 Characterization by Dynamic Light Scattering and Electrophoretic Mobility

Next, characterize the as-prepared NPs by dynamic light scattering. If you use a Zetasizer Nano ZS (Malvern Instruments, Herrenberg, Germany), use 173
backward scatter in the general-purpose mode with automatic measurement position and attenuator selection at a temperature of 25
C. Conduct the zeta potential measurements in monomodal mode. An example for size and charge measurement results is given in Fig. 3.

3.6 Scanning Electron Microscopy

Add purified NPs dropwise to Thermanox Plastic Coverslips (13 mm diameter), allow the drops to air dry, and then sputter the coverslips with Au/Pd using a Polaron SC 515 SEM Sputter Coating System. An example for SEM of LbL-coated NPs is given in Fig. 4.

Fig. 3 (a) The size of fully assembled LbL-coated NPs was determined by dynamic light scattering. The average particle size was 229 nm. The absence of aggregates was shown by the low PDI of 0.065. No additional peaks appeared in the intensity distribution. (b) The zeta potential flips sign after successive deposition of oppositely charged polyelectrolytes

Fig. 4 SEM images of LbL-coated NPs as shown in Fig. 1. The particles are spherical in shape and similar in size. Scale bar: 2 μm

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Notes 1. Purification of substances before use is not required, but nuclease contamination must be rigorously avoided. 2. PEI at this molecular weight is a sticky, high viscosity liquid. Add a spatula full of PEI to the inner wall of a tared snap-cap vial. Next, add the necessary amount of water to obtain the desired concentration. As the dissolution of the polymer might take several hours, prepare the solution in advance and stir if necessary. Once the PEI container has been opened, store in a desiccator to avoid water retention; PEI is hygroscopic. 3. Minor variations in the molecular weight or the lactide-toglycolide ratio of PLGA might influence the characteristics and properties of the NPs. Before using another polymer batch, conduct preliminary studies to make sure that NPs from both batches are comparable. 4. Use 2 mL Eppendorf tubes for centrifugation. Due to varying geometry, tubes from other providers might increase the NP aggregation tendency. 5. HA is a highly hydrophilic polymer. Nevertheless, complete dissolution can take from several minutes to hours. Therefore, the solution should be prepared in advance. 6. The preparation described here results in particles of about 200 nm. However, the size can be adjusted by varying the ratio of PLGA to PEI and by using PLGA of different molecular weight. 7. Centrifugation conditions must be adapted to each individual NP formulation. Larger NPs sediment at lower speeds, whereas smaller particles need higher speeds and/or longer centrifugation times. Additionally, the material properties of the outermost coating layer determine aggregation tendency, which must also be considered in defining a purification protocol. 8. After centrifugation, NPs are easily resuspendable without the need for ultrasonication or vortexing. If this is not the case, the NPs have aggregated and must be discarded. 9. Lyophilization conditions must be adjusted to the respective freeze-drying facility. We used a Christ LMC-2 (Osterode am Harz, Germany). 10. For the preparation of 1 mL of coated particles we used, in order: (a) 200 μL siRNA 20 μM þ 800 μL purified NPs; (b) 10 μL PEI 100 mg/mL þ 990 μL purified NP; (c) 100 μL HA 10 mg/mL þ 900 μL purified NP.

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11. To monitor polyelectrolyte adsorption and NP aggregation, size and zeta potential measurements should be carried out after every coating step. 12. The procedure reported here is not a generally applicable protocol. Before using it for other particles or polymers, perform preliminary tests to determine the optimal coating and purification conditions (e.g., variation of the salt and polymer concentrations or the centrifugation speed). 13. Store the NPs for as short a time as possible to avoid the degradation of siRNA by nucleases. However, if siRNA is replaced by other polyelectrolytes, the NP might be able to be stored at 4
C for several weeks. References 1. Ariga K, Yamauchi Y, Rydzek G, Ji Q, Yonamine Y, Wu KC-W, Hill JP (2014) Layer-by-layer nanoarchitectonics: invention, innovation, and evolution. Chem Lett 43:36–68 2. Decher G, Hong J-D (1991) Buildup of ultrathin multilayer films by a self-assembly process, 1 consecutive adsorption of anionic and cationic bipolar amphiphiles on charged surfaces. Makromol Chem Macromol Symp 46:321–327 3. Yan Y, Such GK, Johnston APR, Lomas H, Caruso F (2011) Toward therapeutic delivery with layer-by-layer engineered particles. ACS Nano 5:4252–4257 4. Wohl BM, Engbersen JFJ (2012) Responsive layer-by-layer materials for drug delivery. J Control Release 158:2–14 5. Elbakry A, Zaky A, Liebl R, Rachel R, Goepferich A, Breunig M (2009) Layer-bylayer assembled gold nanoparticles for siRNA delivery. Nano Lett 9:2059–2064 6. Danhier F, Ansorena E, Silva JM, Coco R, Le Breton A, Pre´at V (2012) PLGA-based nanoparticles: an overview of biomedical applications. J Control Release 161:505–522 7. Neuberg P, Kichler A (2014) Recent developments in nucleic acid delivery with polyethylenimines. Adv Genet 88:263–288 8. Almeida PV, Shahbazi M-A, M€akil€a E, Kaasalainen M, Salonen J, Hirvonen J, Santos

HA (2014) Amine-modified hyaluronic acidfunctionalized porous silicon nanoparticles for targeting breast cancer tumors. Nanoscale 6:10377–10387 9. Martens TF, Remaut K, Demeester J, de Smedt SC, Braeckmans K (2014) Intracellular delivery of nanomaterials: How to catch endosomal escape in the act. Nano Today 9:344–364 10. Deng ZJ, Morton SW, Ben-Akiva E, Dreaden EC, Shopsowitz KE, Hammond PT (2013) Layer-by-layer nanoparticles for systemic codelivery of an anticancer drug and siRNA for potential triple-negative breast cancer treatment. ACS Nano 7:9571–9584 11. Orian-Rousseau V (2010) CD44, a therapeutic target for metastasising tumours. Eur J Cancer 46:1271–1277 12. Guter M, Dillinger Andrea, Scherl Franziska, Fuchshofer R, Breunig M Layer-by-layer assembled nanoparticles for glaucoma therapy. Manuscript in process, in press, https://doi. org/10.1002/smll.201803239 13. Platt VM, Szoka FC (2008) Anticancer therapeutics: targeting macromolecules and nanocarriers to hyaluronan or CD44, a hyaluronan receptor. Mol Pharm 5:474–486 14. Dosio F, Arpicco S, Stella B, Fattal E (2016) Hyaluronic acid for anticancer drug and nucleic acid delivery. Adv Drug Deliv Rev 97:204–236

Chapter 11 Layer-By-Layer Film Engineering for Sequential Gene Delivery Lingxiao Xie, Yi Zou, Sean Carroll, Maria Muniz, and Guangzhao Mao Abstract Layer-by-layer (LbL) films are assembled with poly(amido amine)s (PAAs), a type of polycations containing bioreducible disulfide bond, and DNA plasmids to enable LbL film degradation in physiologic conditions by reacting with glutathione or redox-active membrane proteins. The interior layer structure of the LbL films during assembly and disassembly is studied by atomic force microscopy (AFM), ellipsometry, dynamic light scattering (DLS), and fluorescence spectroscopy. Insertion of barrier layers in bioreducible LbL films is necessary to stabilize the interior layer structure and slow down the film degradation rate to achieve sequential gene delivery. Localized gene delivery from the LbL films is demonstrated using human embryonic kidney 293 (HEK 293) cells. Key words Atomic force microscopy, Bioreducible poly(amido amine), DNA delivery, Film degradation, Layer-by-layer (LbL) film, Polyelectrolyte multilayers, Sequential gene delivery

1

Introduction

1.1 LbL Film for Gene Delivery

Gene therapy, the delivery of exogenous and therapeutic genes to human body for the purpose of treating genetic diseases, has the potential to be the next revolution in modern medicine [1, 2]. In the recent decade, much effort has been focused on gene delivery vectors broadly divided into viral and nonviral types. The nonviral type includes liposome- and polymer-based systems. The majority of these products aim to improve transfection efficiency and safety, building a strong foundation for future clinical applications. However, transfection efficiency, targeting specificity, gene expression regulation vector safety, and stability remain be major challenges in the gene therapy field. LbL deposition of polycations and polyanions to build polyelectrolyte multilayers (PEMs) is an approach to prepare tunable and biologically active surfaces [3]. DNA is bioactive and anionic to be incorporated into the LbL films. On the other hand, LbL films

Manfred Ogris and Haider Sami (eds.), Nanotechnology for Nucleic Acid Delivery: Methods and Protocols, Methods in Molecular Biology, vol. 1943, https://doi.org/10.1007/978-1-4939-9092-4_11, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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can be deposited on various biomedical substrates such as Ti [4, 5], stainless steel stents [6–8], and microneedles [9–11]. LbL deposition has become one of the most promising coating methods that can mimic cellular microenvironments and release therapeutic agents from the surface of biomedical devices. The release dosage, rate, and sequence of therapeutic agents can be controlled by LbL film disassembly. The LbL film disassembly process is closely related to the film assembly process and film interior structure. Due to different polyelectrolyte property, LbL films have two different growth patterns when their thickness or mass is monitored as a function of the number of layers, linear and exponential growth [12, 13]. These two types of LbL films could exhibit different disassembly properties. Based on the relationship between degradable LbL film interior structure and degradation dynamics, it is possible to design LbL film layer structure to achieve sustained and sequential gene delivery. The selection of appropriate polycations as a co-component in LbL films is a key factor for DNA loading efficiency, its protection, film cytotoxicity, and film transfection efficiency [14]. Bioreducible polymers have attracted significant interest for systemic and localized gene delivery because of their lowered cytotoxicity and targeting specificity [1, 15]. Polymers containing the disulfide bond remain stable in extracellular environment to protect its DNA cargo and only degrade under reducing environment such as cell surface containing thiol proteins and intracellular compartments with higher concentrations of glutathione where they are degraded via the thiol–disulfide exchange reaction [16]. 1.2 LbL Film Assembly and Disassembly

LbL film deposition of polyelectrolytes of alternating positive and negative charges has been studied extensively [3]. Adopting a robotic or other types of automatic dipping systems improves the consistency and scalability of LbL film assembly [17, 18]. For example, as Fig. 1 shows, the automatic dipping machine allows the programming of the following LbL deposition cycles. First a charged substrate (e.g., a negative charged glass slide) is immersed in a polycation solution. After a fixed time of polycation adsorption, the substrate is taken out the solution and rinsed by water or buffer in a prefixed number of rinsing cycles before being submerged in the negatively charged DNA plasmid solution. The irreversible adsorption and charge overcompensation in each polyelectrolyte deposition ensures the buildup of the LbL films [19]. LbL film structure is ultimately dependent on the properties of the polyelectrolytes and deposition conditions than the properties of the substrate (e.g., the geometry and charge density of the substrate) [3, 20]. Some LbL films show a linear film growth behavior, that is, the film thickness increases linearly with the number of deposited layers. A typical example of the linearly grown LbL films is the extensively studied pair of poly

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Fig. 1 LbL film deposition process by dip coating. Step 1 and step 3 involve the adsorption of polycation and polyanion, respectively. Step 2 and step 4 are the washing steps to remove excess polyelectrolytes. The four steps are the basic buildup sequence for the LbL film architecture to make multilayers denoted as (polycation/polyanion)n

(styrenesulfonate) (PSS) and poly(allylamine hydrochloride) (PAH) [12]. Other LbL films show an exponential growth behavior in which the thickness increases nonlinearly with the number of deposited layers. Examples of LbL films with exponential growth tendency include films made by poly(L-lysine) (PLL) and alginate [21], hyaluronan (HA) [22], or poly(L-glutamic acid) [12]. The main reason for the two types of film growth modes is attributed to polyelectrolyte interlayer diffusion [12, 18]. When assembling the LbL films, if the polyelectrolyte deposited on the film surface only reacts with the surface layer with minimal diffusion to the interior of the film, the film is likely to display the linear growth mode. The linearly grown films tend to maintain a well-defined layer structure in the absence of significant interlayer diffusion. On the other hand, if the polyelectrolyte interacts extensively with neighboring species and diffuses “in” and “out” of the film, the LbL film shows an exponential growth mode. The interlayer diffusion is different from the ubiquitous interlayer penetration of polyelectrolyte segments into 2–3 adjacent layers in LbL films [23]. The films with interlayer diffusion do not exhibit well defined layer structure. The exponential growth usually appears in LbL films containing weakly charged, short chain polyelectrolytes or biologically derived molecules (e.g., polypeptides and polysaccharides) [13, 24].

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LbL film disassembly is dependent on the film polyelectrolyte constituents. Biodegradable polycations are necessary for localized gene delivery by LbL films because nonbiodegradable vectors such as polyethylenimine (PEI) are too stable in physiologic conditions to allow release of DNA from the LbL films. There are two main types of degradation mechanisms: hydrolytically degradation of the ester bond and reducible degradation of the disulfide bond. The poly(β-amino ester)s family is hydrolytically degradable by the ester bond in the main chain of the polymers in the physiologic environment [25, 26]. Polycation such as poly(amido amine)s (PAAs) containing the disulfide bond are bioreducible by the redox agents in subcellular compartments such as glutathione. For example, the long polymer chain with the disulfide bond can be cleaved into short oligocations by the disulfide–thiol exchange. The lower binding affinity of oligocations with DNA allows for the release of DNA. Based on this mechanism, DNA molecules built into the LbL films can be released into the biological environment [8, 27]. The interlayer structure of the LbL films also can influence the film disassembly dynamics. LbL films without interlayer diffusion are capable of maintaining the desirable layered structure. LbL films with a high degree of interlayer diffusion display a more homogeneous distribution of film components [12]. The stratified and diffused layer structure can give rise to “surface erosion” and “bulk erosion” disassembly type, respectively. While the bulk release capability provides an opportunity to release high concentrations of multiple agents simultaneously, surface erosion is necessary for applications requiring sequential and sustained release of individual film components from the LbL films [18, 24, 28]. 1.3 LbL Film for Sequential Gene Delivery

Our work focuses on the LbL films fabricated with bioreducible poly(amido amine) (PAA) and plasmid DNA. The positively charged biodegradable PAA contains the disulfide bond in its backbone as Fig. 2a shows [18]. PAA and plasmid DNA are alternatively deposited on the glass substrate by the solution dipping method to yield the PAA and DNA bilayer structure. Repeated this procedure, film can be built in multilayer structure. This LbL film fabricated by PAA and DNA with 16.5 bilayer named as LbL-1 (see Note 1). The thickness of LbL-1 film has an exponential film growth pattern which can be characterized by AFM and ellipsometry (Fig. 3) [18]. This implies significant interlayer diffusion during film assembly and this film is likely to degrade fast and in bulk when encountering the reducing environment. In order to acquire sustained and sequential release, one needs to change the film chemical composition and interior structure to limit the degree of interlayer diffusion [18]. Polyethylenimine (PEI) is a polycation with higher charge density and lower diffusion coefficient than PAA. 25 kDa branched PEI has already been widely studied and demonstrated to exhibit high transfection efficiency. The disadvantage of PEI, which limits

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Fig. 2 (a) Chemical structure of bioreducible PAA. Composition change of R1 represents the reducible disulfide bond content. R2 represents the hyperbranched. Reproduced from “Layer-by-layer films with bioreducible and nonbioreducible polycations for sequential DNA release” 2014 with permission from Biomacromolecules. Copyright (2014) American Chemical Society. (b) LbL layer composition of film PAA/DNA (LbL-1 film) and PAA/DNA with the PEI layer (LbL-2 film). Both films have the same total number of layers of 33

Fig. 3 LbL film thickness growth curve as a function of the number of bilayer deposited on the glass substrate as measured by AFM and ellipsometry. The line fit for LbL-2 film has a slope of 4.6 nm per bilayer. Reproduced from “Layer-bylayer films with bioreducible and nonbioreducible polycations for sequential DNA release” 2014 with permission from Biomacromolecules. Copyright (2014) American Chemical Society

its clinical translation, includes its toxicity and nonbiodegradable property. But PEI as a minor component of the bioreducible LbL films may benefit the film disassembly control by providing a separating and stabilizing layer between bioreducible layers to decrease the degree of LbL film interlayer diffusion. As Fig. 2b shows, LbL film with PEI barrier layer with linear growth pattern and relative lower thickness indicates that periodically insertion of the PEI layer into the PAA/DNA LbL film can effectively prevent interlayer diffusion. PEI with high charge density and small diffusion coefficient binds strongly to the film surface during the film assembly

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process. The PEI layer prevents the diffusion of PAA molecules and acts as a barrier layer. The PEI barrier layer inside the film can also screen the residue charges on the film, which has been found to be one of the main driving forces for interlayer diffusion [29] (see Note 2). Interlayer diffusion is the main reason for exponential growth pattern and bulk release property of LbL-1 film (see Notes 3–5). DNA plasmids can be considered as nondiffusing polyelectrolytes in the LbL film system because of their chain length, rigidity, and high charge density [29]. PAA is a type of polycations with interlayer diffusion property. In addition, PAA used here is a biodegradable polyelectrolyte, which can be cleaved into shorter chains in the reducing environment. The short PAA fragments can intensify the degree of interlayer diffusion since the diffusion coefficient is strongly dependent on the molecular weight [29, 30]. This high degree of interlayer diffusion results in a highly mixed LbL interior structure. The short PAA chains have a higher chain mobility, which is a factor leading to fast and bulk disassembly property in LbL-1 film [18]. In order to achieve sustained and sequential gene delivery, it is necessary to incorporate nondiffusible polyelectrolytes such as PEI. In LbL-2 film the second polycation, PEI, binds strongly on the film surface during the assembly process and acts as a barrier to prevent the deposited PAAs to cross the layer. When LbL-2 film is placed in a reducing environment, PEI with high charge density is capable of kinetically trapping the nearby PAA chains or cleaved PAA fragments. Chains in this kinetically trapped state have significantly reduced chain mobility, which is translated into significantly reduced film degradation rate. The PEI chains only can be released out when the entire PEI layer becomes soluble. It is likely that PEI and nearby DNA rearrange into polyplex-like particles from their extended chain conformations before they desorb from the film surface. This process contributes to slow degradation kinetics and small particle size being released from LbL-2 film. During the reducing process, only the surface DNA can be rearranged into the polyplex form due to the highest chain mobility at the film surface. DNA molecules in the film interior are bound strongly to PEI and need to overcome a high free energy to move freely. The differential chain mobility from film surface to bulk is responsible for the sequential release kinetics in LbL-2 film. Our study demonstrated that by inserting a nondiffusing polyelectrolyte such as PEI into bioreducible PAA/DNA LbL films, we can successfully prevent interlayer diffusion and prolong LbL film degradation period to achieve the sustained and sequential gene delivery outcome [18]. This chapter describes a method of engineering LbL films for sustained and sequential release of DNA to a localized environment. The biodegradable PAAs are synthesized by Michael addition copolymerization. The assembly and disassembly of the LbL films

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are characterized by AFM, ellipsometry, DLS, and fluorescence spectroscopy. The bioreducible property enables LbL films to release therapeutic genes in a specific target area and decrease toxicity through polymer degradation. The sustained and sequential delivery can prolong the therapeutic time window and deliver multiple therapeutic genes in a programmable sequence. This work also establishes relationship among LbL film interior structure, assembly sequence, and disassembly dynamics. Provide a mechanism of sustain and sequential DNA releasing in LbL film gene delivery. It provides a simple method to design bioreducible LbL films for sequential and long-time DNA release.

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Materials 1. 1-(2-Aminoethyl)piperazine (AEPZ). 2. 1-Methylpiperazine (MPZ). 3. N,N0 -methylenebis(acrylamide) (MBA). 4. N,N0 -Cystaminebis(acrylamide) (CBA). 5. 1,5-diiodopentane (DIP). 6. Dithiothreitol (DTT). 7. Fluorescein isothiocyanate (FITC). 8. Tetramethylrhodamine isothiocyanate (TRITC). 9. Branched poly(ethylenimine) (PEI): Mw 25 kDa. 10. Poly(2-hydroxyethyl methacrylate) (PolyHEMA). 11. Hyaluronic acid sodium salt from human umbilical cord (HA). 12. Fibronectin from human plasmid, 0.1%. 13. Green fluorescent protein reporter plasmid (pEGFP-N1): from Clontech, 4700 bp. 14. Phosphate-buffered saline, 0.01 M: NaCl, 0.138 M; KCl, 0.0027 M pH 7.4. 15. Acetate buffer, 30 mM, pH 5.5.

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3.1 Synthesis of Bioreducible Poly (Amido Amine) (PAA)

The synthesis of hyperbranched bioreducible PAA is through Michael addition copolymerization by amines and bisacrylamide under mild conditions based on the reported paper [18, 31]. 1. CBA (0.260 g, 1.0 mmol), MBA (0.308 g, 2.0 mmol), and AEPZ (0.193 g, 1.5 mmol) are added in small vial with methanol–water mixture (3 mL, 7:3 v/v) (see Note 6). The reaction with stirring and reacted under 60  C for 2 days. Then 0.019 g,

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0.15 mmol AEPZ is added and allowed to react for another 24 h to stop reaction. The products are acidified with 1 M HCl to Ph4 (see Note 7). 2. The product PAA is fractionated using a semipermeable membrane with a cutoff molecular weight between 30 and 3 kDa. The final products are obtained by freeze-drying and stored in 20  C. 3. The chemical composition of the PAA characterized by 1H and 13 C NMR using a Varian spectrometer (400 MHz) as reported earlier [32]. Number-average molecular weight (Mn), weightaverage (Mw) molecular weight, and polydispersity index (Mw/ Mn) are determined by size exclusion chromatography (SEC) in 0.03 M sodium acetate (pH 4.5) using Waters Ultrahydrogel 250 PKGD column on Waters Alliance 2695 HPLC as reported paper [32]. 3.2 Deposition of LbL Films

The LbL Films are fabricated by dip coating method on glass slide or silicon wafer substrate by using programmable Carl Zeiss HMS50 slide stainer with a Teflon substrate holder [18]. 1. The substrate is immersed into the PAA or PEI solution (0.5 g/L with 30 mM pH 5.5 acetate buffer and 0.1 M NaCl) for 150 s and then in the DNA solution (0.25 g/L with 30 mM pH 5.5 acetate buffer and 0.1 M NaCl) for 150 s. Each dipping is followed by three deionized water rinses, each lasting 60 s. The LbL film deposition procedure is repeated until a desired number of layers is obtained (see Note 8).

3.3

AFM Imaging

1. AFM imaging is conducted by Dimension 3100 AFM from VEECO. LbL film surface morphology and thickness are measured under tapping mode in air. 2. When measuring film thickness, a razor blade is used to scratch film surface to expose part of the substrate. The height difference between the substrate and film surface is taken as the film thickness. Each sample is measured at least at five spots and an average number is reported. 3. The real time LbL film degradation imaging is conducted in tapping mode in liquid environment (5 μL, 20 mM DTT in PBS buffer, pH 7.4). AFM imaging started immediately after injection of DTT solution and continued with an average scan rate of 1–2 Hz until the end of film degradation (see Note 9). The range of frequency, amplitude, integral, and proportional gains used are 7–9 kHz, 0.5–1 V, 0.5–2, and 0.75–3, respectively. 4. The AFM probe used in air is silicon probe with nominal frequency of 150 kHz (VEECO). The probe used in liquid is

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silicon nitride probe with nominal radius of 0.28 N/m (NP type, VEECO). All images are analyzed by Nanoscope software version 5.12 by VEECO (see Note 10). 3.4

Ellipsometry

1. The thickness of LbL films deposited on silicon wafer is measured by a phase-modulated ellipsometer (Beaglehole Instruments, New Zealand). 2. Samples are measured under a fixed incidence angle near the Brewster angle (θB  70 ) and each sample is measured at least in five spots to get average. The thickness is obtained using the Drude equation and the ellipticity, ρ ¼ Im(rp/rs)|θB, where rp and rs are the complex reflection amplitudes for p and s.

3.5 Fluorescence Spectroscopy

1. LbL films containing TRITC- or FITC-labeled PAA are degraded by DTT and monitored by a fluorometer (SpectraMax M5 Plate Reader, Molecular Devices). The TRITC and FITC labeling process is conducted as in a previous study [18]. 2. LbL films are immersed into 20 mM DTT in PBS buffer (2.5 mL, pH 7.4) at room temperature. The degradation solution is collected in each set time point and freeze-dried to concentrate to a total volume of 0.25 mL in order to obtain sufficient fluorescence signals. 3. The cumulative fluorescence intensity divided by the total fluorescence intensity, I/Itotal, is used to determine the amount of dye-labeled PAA released from the LbL film as a function of time.

3.6 Dynamic Light Scattering (DLS)

1. The size of degradation products released from the LbL films is analyzed by zetasizer (Nanosizer ZS, Malvern Instrument) at room temperature [18]. 2. The LbL films are cut into small pieces by 5 mm  5 mm to fit the microcuvette (ZEN0040, Malvern Instrument). A stainless-steel mesh is inserted to the microcuvette and fixed at the top part. 3. The LbL films are placed on the mesh and the whole microcuvette is filled with 20 mM DTT solution (pH 7.4) with the LbL films completely submerged. DLS measurements start immediately to measure the hydrodynamic diameter (DH) of the products released from the LbL films as a function of the degradation time. DH is a function of the diffusion coefficient (D), temperature (T), and viscosity (η) according to the Stokes–Einstein equation: DH ¼ (kT/3πηD). k is the Boltzmann constant. T is 25  C. D is obtained from the autocorrelation function via the cumulant fitting.

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3.7 LbL Film Transfection In Vitro

1. 24-well culture plates are coated with polyHEMA before use by evaporating 200 μL of 20 g/L polyHEMA in 95% ethanol solution under sterile conditions overnight. 2. LbL film transfection is conducted by human embryonic kidney 293 (HEK 293) cells (purchased from American Type Culture Collection). Transfection is indicated by GFP reporter plasmid. 3. Each LbL film is placed into the bottom of the well of the coated tissue culture plate. LbL films may receive additional top coating of fibronectin with 40 μL fibronectin (0.2 g/L) adding to the LbL film surface for 60 min and then sterilized under UV light for 1.5 h (see Note 11). 4. Cells seeded on the LbL film surface with cell number of 4  104 are incubated at 37  C with 1 mL/well DMEM culture medium. Cell medium is changed to a fresh one every 2 days. Cell morphology and fluorescence are detected every day by optical microscope or fluorescence microscope.

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Notes 1. LbL-1 film thickness is measured as a function of the number of layers by AFM and ellipsometry. The AFM and ellipsometry results agreed with each other. The total thickness of the 16.5 bilayer film is 104  9 nm and the film shows the exponential film growth pattern. 2. Based on the LbL-1 film structure PEI layers are inserted in every three bilayers to form the block structure while maintaining the total number of layers same as LbL-1 to yield the LbL-2 film. Using the same thickness measurement methods of AFM and ellipsometry, the film thickness of the LbL-2 film is measured as a function of the number of layers and it shows the linear film growth trend (Fig. 3). The total thickness, 75  4 nm, is lower than that of the LbL-1 film. 3. LbL film disassembly can be monitored by AFM imaging in the reducing environment, 20 mM dithiothreitol (DTT) in PBS buffer (pH 7.4). DTT reacts with the disulfide bond in PAA to degrade the LbL film. Figure 4a shows the time-lapse AFM images of LbL-1 film immersed in 20 mM DTT solution [18]. Those images were focused on an area with part of the film removed and the substrate exposed. The height difference between the substrate and film was monitored during the degradation process. At time zero, LbL-1 film thickness in the PBS buffer was 600 nm with surface roughness of 124 nm. During the degradation process, part of the film pieces started to be peeled off from the substrate in 1 h. For example,

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Fig. 4 (a) Time lapse real-time AFM height images of the LbL-1 film thickness and morphological change in 20 mM DTT in PBS buffer (pH 7.4). Patches left by released film parts were marked in the images. Scan size is 30 μm. Z-range is 2 μm. (b) Time lapse AFM height images of the LbL-2 film thickness and morphological change treated with 20 mM DTT in PBS buffer (pH 7.4). The scan size is 15 μm. Z-range is 150 nm. Film thickness change as a function of degradation time in 20 mM DTT measured by AFM for (c) LbL-1 film and (d) LbL-2 film. Reproduced from “Layer-by-layer films with bioreducible and nonbioreducible polycations for sequential DNA release” 2014 with permission from Biomacromolecules. Copyright (2014) American Chemical Society

in 64 min, the released pieces with size of 3 μm by 17 μm, in 72 min, 9 μm by 15 μm, and in 83 min, 6 μm by 19 μm were captured by AFM. The evidence points to a film degradation behavior of bulk degradation and not the often assumed layerby-layer peeling off dynamics. The film was completely removed from the substrate after 100 min. The degradation behavior of the LbL-2 film that exhibits the linear growth mode was studied in the same reducing environment by AFM. As show in Fig. 4b, LbL-2 film degradation process lasted for 38 h, much longer than LbL-1 film [18]. Due to the longer reducing time, the film was taken out from the DTT solution periodically and imaged by AFM in air instead of in situ imaging. During the degradation process, we did not observe any micrometer sized pieces being peeled off the substrate. Instead, we observed continuous formation of nanometer sized particles on the film surface followed by subsequent release from the film surface. The average diameter of the particle is around 300 nm according to zeta sizer measurements. LbL-2 film degrades from the surface and in a much longer time than LbL-1 film. Figure 4c, d show the thickness variation during film disassembly in 20 mM DTT measured by AFM [18]. LbL-1 film does not show obvious thickness change

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in the first hour and then a rapid decrease from 60 to 90 min. But for LbL-2 film, its film thickness decrease rate in the first 3 h is relatively higher than the rest of the time. After 3 h, the degradation rate slows down with much more gradual thickness decrease lasting 120 h. The AFM results of LbL film degradation demonstrates that periodically insertion of the PEI barrier layer prolongs the film degradation time, from 1.5 h in LbL-1 film to more than 120 h in LbL-2 film. The film morphological difference supports the conclusion that LbL-1 film undergoes bulk degradation while LbL-2 film undergoes surface degradation. Films undergoing surface degradation are capable of sequential release of DNA. 4. LbL film disassembly is further analyzed by fluorescence spectroscopy. PAA is labeled with fluorescence dye, FITC or TRITC. The rate of LbL film disassembly can thus be determined by analyzing the cumulative fluorescence intensity of the degradation solution as a function of degradation time. As shown in Fig. 5a, b [18], LbL-1 film shows increased fluorescence intensity in the first hour before reaching the plateau, which indicates that all the PAA molecules in the film were released into the DTT solution in 1 h. But for LbL-2 film, the cumulative fluorescence intensity increased gradually continuously for up to 140 h. When the PAA molecules in top half of the LbL film were labeled with FITC and in the bottom half labeled with TRITC (Fig. 5c, d), LbL-1 film and LbL-2 film showed different fluorescence intensity variation trends [18]. All the PAA molecules in LbL-1 film were released at the same time. But for LbL-2 film, the appearance of the TRITC signal was delayed than that of the FITC. It indicates that in lbL-2 film PAA molecules in the bottom half are released later than those in the top half. The fluorescence data demonstrate the sequential DNA release capability of bioreducible LbL films containing PEI barrier layers. 5. The size distribution of particles released from the bioreducible LbL films in 20 mM DTT is analyzed by dynamic light scattering (DLS). The hydrodynamic diameter (DH) of released particles in 20 mM DTT as a function of degradation time is shown in Fig. 6 [18]. LbL-1 film releases particles with an average DH around 700 nm in first 30 min and then the particle size decreased to around 100 nm after 2 h. Since LbL-1 film completely degrades within 1.5 h, we can surmise that the particles after 2 h are the secondary degradation products due to further degradation in DTT solution of the primary 700 nm film pieces directly coming out of the film [18]. The DH of the released particles from LbL-2 film maintains a constant size range between 300 and 400 nm. The particle size measurements by DLS are consistent with the AFM results in Fig. 4,

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and the total length of degradation time is similar to fluorescence results in Fig. 5. Based on these measurements, the PEI barrier layer in the PAA/DNA LbL film system can slow down the degradation rate and reduce the released particle size. The small size is necessary for endocytic uptake of DNA particles and subsequent DNA transfection [1, 18]. 6. The amine group in AEPZ has different reaction activity, secondary amine (original) > primary amine > > secondary amine (formed) [33], which allows synthesis of either linear or hyperbranched structure via adjusting feeding ration of AEPZ: (CBA + MBA). The feeding ratio of AEPZ: (CBA + MBA) is 1:2 yields hyperbranched structure and ratio of 1:1 yields linear structure. The disulfide bond contents in polymer chain also can be adjusted by the feeding ratio of monomer CBA: MBA. 7. Acidification of PAA in pH 4 can improve the solubility during dialyzing in deionized water via semipermeable membrane. It

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also can improve the solubility of PAA in 30 mM pH 5.5 acetate buffer to ensure successful LbL film dipping coating process. 8. Plasmid DNA structure is fragile. We recommend avoiding high speed vortexing and violently pipetting. During the dip coating process, each polymer and DNA solution should be refreshed in at least every 8-dipping cycle to avoid contamination or concentration variation. The LbL films can be stored in deionized water at 4  C. 9. Fresh DTT should be used because it can be easily oxidized. 10. During in situ tapping mode AFM imaging, first the frequency selection in manually tuning process is critical for liquid operation. The peak around 8 kHZ usually could provide high quality image. Since the in situ AFM imaging continues for several hours, it is necessary to track and adjust the amplitude set point in order to remain at the minimum force. The background noise can be minimized by optimized integral gain and proportional gain. 11. Before seeding cells on LbL films, the films can be pretreated by fibronectin to improve the cell attachment. One can mix 40 μL fibronectin with 40 μL PBS buffer in order to fully cover the film surface. After UV light sterilization, the films are first immersed in cell culture medium in order to saturate the films before cell seeding.

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Chapter 12 Surface- and Hydrogel-Mediated Delivery of Nucleic Acid Nanoparticles Angela K. Pannier, Tyler Kozisek, and Tatiana Segura Abstract Gene expression within a cell population can be directly altered through gene delivery approaches. Traditionally for nonviral delivery, plasmids or siRNA molecules, encoding or targeting the gene of interest, are packaged within nanoparticles. These nanoparticles are then delivered to the media surrounding cells seeded onto tissue culture plastic; this technique is termed bolus delivery. Although bolus delivery is widely utilized to screen for efficient delivery vehicles and to study gene function in vitro, this delivery strategy may not result in efficient gene transfer for all cell types or may not identify those delivery vehicles that will be efficient in vivo. Furthermore, bolus delivery cannot be used in applications where patterning of gene expression is needed. In this chapter, we describe methods that incorporate material surfaces (i.e., surfacemediated delivery) or hydrogel scaffolds (i.e., hydrogel-mediated delivery) to efficiently deliver genes. This chapter includes protocols for surface-mediated DNA delivery focusing on the simplest and most effective methods, which include nonspecific immobilization of DNA complexes (both polymer and lipid vectors) onto serum-coated cell culture polystyrene and self-assembled monolayers (SAMs) of alkanethiols on gold. Also, protocols for the encapsulation of DNA/cationic polymer nanoparticles into hydrogel scaffolds are described, including methods for the encapsulation of low amounts of DNA (0.2 μg/μl) since incorporation of high amounts of DNA pose significant challenges due to aggregation. Key words Gene delivery, Hydrogel, Surface-mediated, Transfection, Nonviral

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Introduction Gene expression within a cell population can be directly altered through gene delivery approaches, which have tremendous potential for therapeutic applications, such as gene therapy to correct genetic deficiencies, or tissue engineering scaffolds, where gene delivery can present chemical factors to guide tissue formation in regeneration matrices for treatment of organ loss and failure. Furthermore, gene delivery is often critical to research applications, such as functional genomics, cell culture studies, and biotechnological assays. Gene delivery can be performed with both viral and

Manfred Ogris and Haider Sami (eds.), Nanotechnology for Nucleic Acid Delivery: Methods and Protocols, Methods in Molecular Biology, vol. 1943, https://doi.org/10.1007/978-1-4939-9092-4_12, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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Fig. 1 Bolus, surface- and hydrogel-mediated gene delivery

nonviral vectors, the latter of which are the focus of the techniques described in this chapter. Traditionally for nonviral delivery, plasmids, mRNA, or RNAi (miRNA or siRNA) molecules, encoding or targeting the gene of interest, are packaged within nanoparticles. These nanoparticles are then delivered to the media surrounding cells seeded onto tissue culture plastic; this technique is termed bolus delivery (Fig. 1). Although bolus delivery is widely utilized to screen for efficient delivery vehicles and to study gene function in vitro, this delivery strategy may not result in efficient gene transfer for all cell types or may not identify those delivery vehicles that will be efficient in vivo. Furthermore, bolus delivery cannot be used in applications where patterning of gene expression is needed. In this chapter, we will describe methods that incorporate material surfaces (i.e., surface-mediated delivery) or hydrogel scaffolds (i.e., hydrogel-mediated delivery) to efficiently deliver genes (Fig. 1). Surface-mediated delivery, also termed solid-phase delivery or reverse transfection, refers to the delivery of nucleic acid nanoparticles from a surface or biomaterial that supports cell adhesion. Nucleic acid nanoparticles are immobilized to the surface or biomaterial and cells are plated directly on top of the immobilized nanoparticles (Fig. 1), which enhances the extent of transgene expression, along with increasing cell viability [1, 2]. This method of delivery was first described in 2002 using a specific avidin-biotin bond to tether nanoparticles to surfaces [3, 4] and was later extended to nonspecific absorption of the nanoparticles to a variety of biomaterial surfaces [1, 2, 5–18]. Nonspecific adsorption of nanoparticles is accomplished through noncovalent mechanisms [1, 5–11, 19–22], including hydrophobic, electrostatic, and van der Waals interactions. Nonspecific binding depends upon the molecular composition of the vector (e.g., lipid versus polymer) and the relative quantity of each (e.g., nitrogen to phosphate ratio or N/P), as well as properties of the surface (e.g., hydrophilicity, charge, presence of serum proteins). In surface-mediated delivery, nucleic acid nanoparticles are concentrated at the delivery site and targeted to cells adhered to the substrate [1, 3, 4, 23]. The major advantages of surface-

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mediated nucleic acid delivery are as follows: (1) decreased nucleic acid nanoparticle aggregation and degradation, since nanoparticles can be produced in the most favorable buffer for their formation and then immobilized to the surface, which preserves the size observed in solution and avoids exposure of the particles in solution to the cell culture media, which often induces aggregation and decomplexation; (2) cells cultured on the substrate are exposed to elevated nucleic acid concentrations within the local microenvironment (but typically overall concentration of nucleic acid nanoparticles is minimized when compared to bolus techniques), which enhances transfection and minimizes cytotoxicity [1, 3–11, 24]; (3) elimination of mass transport limitations for the nanoparticles to reach the cell; and (4) immobilization of nucleic acid complexes to substrates allows for the ability of pattern transfection [10, 16, 25], allowing for spatial control of gene delivery, which is critical for biotechnological and tissue engineering applications. Below the reader will find protocols for surface-mediated DNA delivery focusing on the simplest and most effective methods, which include nonspecific immobilization of DNA complexes (both polymer and lipid vectors) onto serum-coated cell culture polystyrene and self-assembled monolayers (SAMs) of alkanethiols on gold. 3-Dimensional (3D) matrix-mediated delivery offers many advantages over conventional, surface-mediated delivery as the 3D matrices are more biomimetic than their 2D counterparts, often leading to increased cell spreading and proliferation, which favor transfection [18]. 3D matrices, more specifically 3D hydrogel scaffolds, have been studied for their ability to mediate transfection for over two decades, primarily through the encapsulation of naked DNA during hydrogel formation. Naked DNA has been successfully incorporated inside collagen [26], hyaluronic acid [27, 28], PEG-poly(lactic acid)-PEG [29], alginate [30], oligo(polyethylene glycol) fumarate [31], engineered silk elastin [32], and various other natural and synthetic hydrogels [33]. Although naked DNA has shown gene expression and ability to guide regeneration in vivo [26, 34–36], limitations with low gene transfer efficiency, rapid diffusion of the DNA from the hydrogel scaffold, and rapid DNA degradation motivated the use of DNA nanoparticles instead of naked DNA. Nucleic acids condensed either with cationic peptides, lipids, polymers, or lipid–polymer hybrids, have been introduced into fibrin hydrogels [37–41], enzymatically degradable PEG hydrogels [42], PEG-hyaluronic acid hydrogels [38], and various other hydrogels and 3D matrices [43–47]. The delivery of genes from hydrogel scaffolds is becoming an attractive route to introduce transgenes to cells for several reasons. First, in the biotechnology area, where new delivery agents are being investigated for in vivo gene transfer, the delivery of genes inside a matrix may be a better model for in vivo gene transfer. For example, hydrogels can be used to recreate disease models in vitro, including multiple cell

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types and complex extracellular matrices (e.g., cancer and organoid models) [48–51]. Second, in tissue engineering and regenerative medicine applications, it is often desired to transfect cells either infiltrating the scaffold (endogenous) or that are transplanted. In the case of endogenous cell transfection, the hydrogel scaffold serves as a depot that contains genes or siRNA that can transfect cells as the cells infiltrate the scaffold or the matrix is degraded and the genes or siRNA is released. In the case of transplanted cell transfection, the matrix and cells are encapsulated together. The goal here is to transfect the transplanted cells at a later time to induce their differentiation or the differentiation of nearby cells. Below, the reader will find protocols for the encapsulation of DNA–cationic polymer nanoparticles into hydrogel scaffolds. We divide the encapsulation into low amounts of DNA (0.2 μg/μl) since incorporation of high amounts of DNA pose significant challenges due to aggregation.

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PBS (prepared as described above). Prepare sterilely and aliquots may be stored at 4  C for short term (