Peptide Conjugation: Methods and Protocols (Methods in Molecular Biology, 2355) 1071616161, 9781071616161

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Methods in Molecular Biology 2355

Waleed M. Hussein Rachel J. Stephenson · Istvan Toth Editors

Peptide Conjugation Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

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

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

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

Peptide Conjugation Methods and Protocols

Edited by

Waleed M. Hussein, Rachel J. Stephenson and Istvan Toth School of Chemistry and Molecular Biosciences, The University of Queensland, St. Lucia, QLD, Australia

Editors Waleed M. Hussein School of Chemistry and Molecular Biosciences The University of Queensland St. Lucia, QLD, Australia

Rachel J. Stephenson School of Chemistry and Molecular Biosciences The University of Queensland St. Lucia, QLD, Australia

Istvan Toth School of Chemistry and Molecular Biosciences The University of Queensland St. Lucia, QLD, Australia

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1616-1 ISBN 978-1-0716-1617-8 (eBook) https://doi.org/10.1007/978-1-0716-1617-8 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface Several synthetic strategies exist for the preparation of well-defined peptide conjugates. Since the introduction of chemoselective conjugation of unprotected peptides using native chemical ligation, many peptide ligation techniques have been developed, including coppercatalyzed azide-alkyne cycloaddition click reaction, peptide hydrazide ligation, 5-oxaproline ketoacid-hydroxylamine ligation, and others. In this book, a variety of diverse protocols for peptide conjugation are detailed. In order to be readily understandable and useful for trained researchers and undergraduate students, alike, techniques are described in a detailed, easy-to-follow, step-by-step manner. As an additional resource, troubleshooting notes and alternative approaches that could be used to fix common problems are included at the end of each protocol. For several critical processes, the book provides a comprehensive set of basic, but thoroughly tested and scientifically valid techniques that will allow equipped readers to prepare, purify, characterize, and use peptide conjugation techniques effectively for chemical, biochemical, and biological studies. To list a few examples, this book contains protocols for peptide conjugation to polymers, gold nanoparticles, proteins, pegylated lipids, and vitamins. Chapters also cover enzymatic ligation using sortase A, electrochemical C-terminal peptide modifications, oxime/hydrazone conjugation at histidine in unprotected peptides, chemoselective α-ketoacid-hydroxylamine ligation, construction of a phage-displayed cyclic-peptide library, quantum dot-peptide conjugates, preparation of lipopeptides by CLipPA chemistry, and other very interesting techniques that enrich the field of peptide conjugation. St. Lucia, QLD, Australia

Waleed H. Hussein Rachel J. Stephenson Istvan Toth

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

1 Peptide-Polymer Conjugation Via Copper-Catalyzed Alkyne-Azide 1,3-Dipolar Cycloaddition. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Waleed M. Hussein, Istvan Toth, and Mariusz Skwarczynski 2 Conjugation of Peptides to Gold Nanoparticles. . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pornsuda Maraming and James Chen Yong Kah 3 Peptide-Protein Conjugation and Characterization to Develop Vaccines for Group A Streptococcus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sharareh Eskandari, Michael F. Good, and Manisha Pandey 4 Oxime/Hydrazone Conjugation at Histidine: Late-Stage Functionalization Approach of Unprotected Peptides . . . . . . . . . . . . . . . . . . . . . . . Anaı¨s F. M. Noisier and Ranganath Gopalakrishnan 5 Modification of Nanoparticles with Transferrin for Targeting Brain Tissues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sara Aly Attia, Xiang Li, Nina Filipczak, Daniel F. Costa, and Vladimir P. Torchilin 6 Peptide-Pegylated Lipid Conjugation Via Copper-Catalyzed Alkyne-Azide 1,3-Dipolar Cycloaddition. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Waleed M. Hussein and Istvan Toth 7 Vitamin B12 – Peptide Nucleic Acid Conjugates . . . . . . . . . . . . . . . . . . . . . . . . . . . . Aleksandra J. Wierzba, Monika Wojciechowska, Joanna Trylska, and Dorota Gryko 8 Enzymatic Ligation of Disulfide-Rich Animal Venom Peptides: Using Sortase A to Form Double-Knotted Peptides . . . . . . . . . . . . . . . . . . . . . . . . . Poanna Tran and Christina I. Schroeder 9 Solid-State, Thermal Synthesis of Peptide/Protein–Boron Cluster Conjugates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Krzysztof Fink and Tomasz M. Goszczyn´ski 10 Bioconjugation of Peptides to Hybrid Gold Nanoparticles . . . . . . . . . . . . . . . . . . . Rosalba Moretta, Monica Terracciano, Ilaria Rea, and Luca De Stefano 11 Design and Synthesis of a Peptide-Based Glioma-Targeted Drug Delivery Vector gHope2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . € lo Langel Kaido Kurrikoff, Nikita Oskolkov, Elo Eriste, and U 12 Electrochemically Enabled C-Terminal Peptide Modifications . . . . . . . . . . . . . . . . Yutong Lin and Lara R. Malins 13 Double Conjugation Using Mercapto-Acryloyl and Alkyne-Azide Reactions for the Synthesis of Branched Multiantigenic Vaccine Candidates. . . . Waleed M. Hussein, Mariusz Skwarczynski, and Istvan Toth

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Contents

Chemical Protein Synthesis by Chemoselective α-Ketoacid–Hydroxylamine (KAHA) Ligations with 5-Oxaproline . . . . . . . . . . . . Jakob Farnung, Haewon Song, and Jeffrey W. Bode Site-Specific Modification of Single-Chain Affinity Ligands for Fluorescence Labeling, Radiolabeling, and Bioconjugation . . . . . . . . . . . . . . . Boya Zhang, Sachith M. Vidanapathirana, and Colin F. Greineder Preparation and Characterization of Quantum Dot-Peptide Conjugates Based on Polyhistidine Tags . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Katherine D. Krause, Hsin-Yun Tsai, Kelly Rees, Hyungki Kim, and W. Russ Algar The Construction of a Genetically Encoded, Phage-Displayed Cyclic-Peptide Library. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Peng-Hsun Chase Chen and Wenshe Ray Liu The Chemical Synthesis of Site-Specifically Modified Proteins Via Diselenide-Selenoester Ligation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rhys C. Griffiths and Nicholas J. Mitchell Synthesis of Lipopeptides by CLipPA Chemistry. . . . . . . . . . . . . . . . . . . . . . . . . . . . Victor Yim, Yann O. Hermant, Paul W. R. Harris, and Margaret A. Brimble Constraining TAT Peptide by γPNA Hairpin for Enhanced Cellular Delivery of Biomolecules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Siddhartha Thennakoon, Rick Postema, and Xiaohong Tan Preparation of mRNA Polyplexes with Post-conjugated Endosome-Disruptive Peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Bo Lou, Chun Yin Jerry Lau, Wim E. Hennink, and Enrico Mastrobattista Targeted Subcellular Protein Delivery Using Cleavable Cyclic Cell-Penetrating Peptide-Conjugates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Luise Franz, Anselm F. L. Schneider, and Christian P. R. Hackenberger Facile Chemoselective Modification of Thioethers Generates Chiral Center-Induced Helical Peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yinghuan Liu, Kuan Hu, Feng Yin, and Zigang Li

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

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301 323

Contributors W. RUSS ALGAR • Department of Chemistry, University of British Columbia, Vancouver, BC, Canada SARA ALY ATTIA • Center for Pharmaceutical Biotechnology and Nanomedicine, Northeastern University, Boston, MA, USA JEFFREY W. BODE • Laboratorium fu¨r Organische Chemie, Department of Chemistry and Applied Biosciences, ETH Zu¨rich, Zu¨rich, Switzerland; Institute of Transformative BioMolecules (WPI-ITbM), Nagoya University, Nagoya, Japan MARGARET A. BRIMBLE • School of Chemical Sciences, The University of Auckland, Auckland, New Zealand; School of Biological Sciences, The University of Auckland, Auckland, New Zealand; The Maurice Wilkins Centre for Molecular Biodiscovery, The University of Auckland, Auckland, New Zealand PENG-HSUN CHASE CHEN • Department of Chemistry, Texas A&M University, College Station, TX, USA DANIEL F. COSTA • Center for Pharmaceutical Biotechnology and Nanomedicine, Northeastern University, Boston, MA, USA LUCA DE STEFANO • Institute of Applied Sciences and Intelligent Systems, National Research Council, Naples, Italy ELO ERISTE • TBD-Biodiscovery, Tartu, Estonia SHARAREH ESKANDARI • Institute for Glycomics, Griffith University, Southport, QLD, Australia JAKOB FARNUNG • Laboratorium fu¨r Organische Chemie, Department of Chemistry and Applied Biosciences, ETH Zu¨rich, Zu¨rich, Switzerland NINA FILIPCZAK • Center for Pharmaceutical Biotechnology and Nanomedicine, Northeastern University, Boston, MA, USA KRZYSZTOF FINK • Laboratory of Biomedical Chemistry, Department of Experimental Oncology, Hirszfeld Institute of Immunology and Experimental Therapy, Polish Academy of Sciences, Wroclaw, Poland LUISE FRANZ • Leibniz-Forschungsinstitut fu¨r Molekulare Pharmakologie (FMP), Berlin, Germany; Institute of Chemistry and Biochemistry, Freie Universit€ at Berlin, Berlin, Germany MICHAEL F. GOOD • Institute for Glycomics, Griffith University, Southport, QLD, Australia RANGANATH GOPALAKRISHNAN • Medicinal Chemistry, Research and Early Development, Respiratory & Immunology, BioPharmaceuticals R&D, AstraZeneca, Gothenburg, Sweden TOMASZ M. GOSZCZYN´SKI • Laboratory of Biomedical Chemistry, Department of Experimental Oncology, Hirszfeld Institute of Immunology and Experimental Therapy, Polish Academy of Sciences, Wroclaw, Poland COLIN F. GREINEDER • Departments of Pharmacology and Emergency Medicine, University of Michigan, Ann Arbor, MI, USA; BioInterfaces Institute, University of Michigan, Ann Arbor, MI, USA RHYS C. GRIFFITHS • School of Chemistry, University of Nottingham, University Park, Nottingham, UK

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Contributors

DOROTA GRYKO • Institute of Organic Chemistry, Polish Academy of Sciences, Warsaw, Poland CHRISTIAN P. R. HACKENBERGER • Leibniz-Forschungsinstitut fu¨r Molekulare Pharmakologie (FMP), Berlin, Germany; Department of Chemistry, Humboldt-Universit€ a t zu Berlin, Berlin, Germany PAUL W. R. HARRIS • School of Chemical Sciences, The University of Auckland, Auckland, New Zealand; School of Biological Sciences, The University of Auckland, Auckland, New Zealand; The Maurice Wilkins Centre for Molecular Biodiscovery, The University of Auckland, Auckland, New Zealand WIM E. HENNINK • Department of Pharmaceutics, Utrecht Institute for Pharmaceutical Science (UIPS), Utrecht University, Utrecht, The Netherlands YANN O. HERMANT • School of Chemical Sciences, The University of Auckland, Auckland, New Zealand; School of Biological Sciences, The University of Auckland, Auckland, New Zealand KUAN HU • State Key Laboratory of Chemical Oncogenomics, School of Chemical Biology and Biotechnology, Shenzhen Graduate School, Peking University, Shenzhen, China WALEED M. HUSSEIN • School of Chemistry and Molecular Biosciences, The University of Queensland, St. Lucia, QLD, Australia JAMES CHEN YONG KAH • Department of Biomedical Engineering, National University of Singapore, Singapore, Singapore HYUNGKI KIM • Department of Chemistry, University of British Columbia, Vancouver, BC, Canada KATHERINE D. KRAUSE • Department of Chemistry, University of British Columbia, Vancouver, BC, Canada KAIDO KURRIKOFF • University of Tartu, Institute of Technology, Tartu, Estonia € LO LANGEL • University of Tartu, Institute of Technology, Tartu, Estonia; Department of U Biochemistry and Biophysics, Stockholm University, Stockholm, Sweden CHUN YIN JERRY LAU • Department of Pharmaceutics, Utrecht Institute for Pharmaceutical Science (UIPS), Utrecht University, Utrecht, The Netherlands XIANG LI • Center for Pharmaceutical Biotechnology and Nanomedicine, Northeastern University, Boston, MA, USA; State Key Laboratory of Innovative Drug and Efficient Energy-Saving Pharmaceutical Equipment, Jiangxi University of Traditional Chinese Medicine, Nanchang, China ZIGANG LI • State Key Laboratory of Chemical Oncogenomics, School of Chemical Biology and Biotechnology, Shenzhen Graduate School, Peking University, Shenzhen, China YUTONG LIN • Research School of Chemistry, Australian National University, Canberra, ACT, Australia WENSHE RAY LIU • Department of Chemistry, Texas A&M University, College Station, TX, USA YINGHUAN LIU • State Key Laboratory of Chemical Oncogenomics, School of Chemical Biology and Biotechnology, Shenzhen Graduate School, Peking University, Shenzhen, China BO LOU • Department of Pharmaceutics, Utrecht Institute for Pharmaceutical Science (UIPS), Utrecht University, Utrecht, The Netherlands LARA R. MALINS • Research School of Chemistry, Australian National University, Canberra, ACT, Australia PORNSUDA MARAMING • Faculty of Associated Medical Sciences, Centre for Research and Development of Medical Diagnostic Laboratories, Khon Kaen University, Khon Kaen, Thailand

Contributors

xi

ENRICO MASTROBATTISTA • Department of Pharmaceutics, Utrecht Institute for Pharmaceutical Science (UIPS), Utrecht University, Utrecht, The Netherlands NICHOLAS J. MITCHELL • School of Chemistry, University of Nottingham, University Park, Nottingham, UK ROSALBA MORETTA • Institute of Applied Sciences and Intelligent Systems, National Research Council, Naples, Italy ANAI¨S F. M. NOISIER • Medicinal Chemistry, Research and Early Development, Cardiovascular, Renal and Metabolism, BioPharmaceuticals R&D, AstraZeneca, Gothenburg, Sweden NIKITA OSKOLKOV • Lumiprobe Corporation, Hunt Valley, MD, USA; Department of Psychiatry and Behavioral Sciences, Johns Hopkins University School of Medicine, Baltimore, MD, USA MANISHA PANDEY • Institute for Glycomics, Griffith University, Southport, QLD, Australia RICK POSTEMA • Department of Chemistry and Center for Photochemical Sciences, Bowling Green State University, Bowling Green, OH, USA ILARIA REA • Institute of Applied Sciences and Intelligent Systems, National Research Council, Naples, Italy KELLY REES • Department of Chemistry, University of British Columbia, Vancouver, BC, Canada ANSELM F. L. SCHNEIDER • Leibniz-Forschungsinstitut fu¨r Molekulare Pharmakologie (FMP), Berlin, Germany; Institute of Chemistry and Biochemistry, Freie Universit€ at Berlin, Berlin, Germany CHRISTINA I. SCHROEDER • Institute for Molecular Bioscience, The University of Queensland, Brisbane, QLD, Australia; National Cancer Institute, National Institutes of Health, Frederick, MD, USA MARIUSZ SKWARCZYNSKI • School of Chemistry and Molecular Biosciences, The University of Queensland, St. Lucia, QLD, Australia HAEWON SONG • Laboratorium fu¨r Organische Chemie, Department of Chemistry and Applied Biosciences, ETH Zu¨rich, Zu¨rich, Switzerland XIAOHONG TAN • Department of Chemistry and Center for Photochemical Sciences, Bowling Green State University, Bowling Green, OH, USA MONICA TERRACCIANO • Department of Pharmacy, University of Naples Federico II, Naples, Italy SIDDHARTHA THENNAKOON • Department of Chemistry and Center for Photochemical Sciences, Bowling Green State University, Bowling Green, OH, USA VLADIMIR P. TORCHILIN • Center for Pharmaceutical Biotechnology and Nanomedicine, Northeastern University, Boston, MA, USA; Department of Oncology, Radiotherapy and Plastic Surgery, I.M. Sechenov First Moscow State Medical University (Sechenov University), Moscow, Russia ISTVAN TOTH • School of Chemistry and Molecular Biosciences, The University of Queensland, St. Lucia, QLD, Australia POANNA TRAN • Institute for Molecular Bioscience, The University of Queensland, Brisbane, QLD, Australia JOANNA TRYLSKA • Centre of New Technologies, University of Warsaw, Warsaw, Poland HSIN-YUN TSAI • Department of Chemistry, University of British Columbia, Vancouver, BC, Canada

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Contributors

SACHITH M. VIDANAPATHIRANA • Departments of Pharmacology and Emergency Medicine, University of Michigan, Ann Arbor, MI, USA; BioInterfaces Institute, University of Michigan, Ann Arbor, MI, USA ALEKSANDRA J. WIERZBA • Institute of Organic Chemistry, Polish Academy of Sciences, Warsaw, Poland MONIKA WOJCIECHOWSKA • Centre of New Technologies, University of Warsaw, Warsaw, Poland VICTOR YIM • School of Chemical Sciences, The University of Auckland, Auckland, New Zealand; School of Biological Sciences, The University of Auckland, Auckland, New Zealand FENG YIN • State Key Laboratory of Chemical Oncogenomics, School of Chemical Biology and Biotechnology, Shenzhen Graduate School, Peking University, Shenzhen, China BOYA ZHANG • Departments of Pharmacology and Emergency Medicine, University of Michigan, Ann Arbor, MI, USA; BioInterfaces Institute, University of Michigan, Ann Arbor, MI, USA

Chapter 1 Peptide-Polymer Conjugation Via Copper-Catalyzed Alkyne-Azide 1,3-Dipolar Cycloaddition Waleed M. Hussein, Istvan Toth, and Mariusz Skwarczynski Abstract Dendrimers are structurally well-defined, artificial polymers with physicochemical characteristics that often imitate biomacromolecules. Consequently, they are encouraging candidates for the delivery of peptidebased vaccines. We developed a synthetic protocol for conjugating a peptide antigen derived from human papillomavirus (HPV) E7 protein to a poly(t-butyl acrylate) dendrimer to construct a vaccine candidate. The synthetic pathway utilized copper-catalyzed alkyne-azide 1,3-dipolar cycloaddition (CuAAC) click reaction, and resulted in a 76% substitution ratio of the 8-arm dendrimer. The obtained peptide-polymer construct was self-assembled, dialyzed, and characterized by microanalysis and dynamic light scattering. Key words Peptide-polymer conjugate, Alkyne-azide cycloaddition, Click chemistry, Peptide vaccines

1

Introduction Dendrimers are structurally well-defined, synthetic polymers that can be engineered to mimic the size and physiochemical properties of biomolecules (e.g., proteins) [1]. Their branched design allows antigen molecules to be attached and presented at the dendrimer’s periphery. By mimicking natural pathogens (e.g., viral capsid), multiple copies of the antigen are displayed to the immune system [1, 2]. Moreover, such conjugates can be designed to have amphiphilic properties and, therefore, the ability to self-assemble into nanoparticles to further mimic natural infectious agents [3]. Consequently, it was hypothesized that these dendrimers could produce self-adjuvanting delivery systems, negating the need for toxic adjuvants (immune stimulants) in vaccine design. To fulfill the above criteria, hydrophobic polyacrylate ester was chosen because poly(acrylic acid) and its esters are simple to synthesize and known to have a good safety profile [4, 5]. Linear and branched polyacrylate-peptide conjugates were developed as self-adjuvanting delivery systems for peptide antigens. The

Waleed M. Hussein, Rachel J. Stephenson and Istvan Toth (eds.), Peptide Conjugation: Methods and Protocols, Methods in Molecular Biology, vol. 2355, https://doi.org/10.1007/978-1-0716-1617-8_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Synthesis of peptide-polymer conjugate 3

conjugates were effective in the generation of strong cellular and humoral immune responses without the need for external adjuvant [6–17]. For example, a polyacrylate-based delivery system was used to develop peptide-based vaccines for the treatment of E7-positive tumor cells. 8Qmin (E744–57, QAEPDRAHYNIVTF) is a cytotoxic T-lymphocyte (CD8+) epitope derived from human papillomavirus HPV16 E7 protein [18]. As a therapeutic anticancer vaccine, the 8Qmin-polyacrylate conjugate was able to eradicate E7-positive TC-1 tumors in mice with a 100% survival rate [16]. In this chapter, conjugation between the azide derivative of 8Qmin (1) [12] and poly(t-butyl acrylate) 8-arm dendrimer (2) [16] was performed using copper-catalyzed alkyne-azide 1,3-dipolar cycloaddition (CuAAC) reaction in the presence of copper wire [19]. The final polyacrylate-8Qmin conjugate (3) was produced with a 76% antigen substitution ratio [16] (Fig. 1).

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Materials

2.1 Synthesis of Peptide-Conjugate 3

1. N-terminus 8Qmin azide (1). 2. Dried round-bottom flask (2 mL). 3. Poly(t-butyl acrylate) (2). 4. N,N-Dimethylformamide (DMF). 5. Copper wire. 6. Nitrogen balloon. 7. Stopwatch. 8. Aluminum foil. 9. Hot plate magnetic stirrer. 10. Oil bath.

Peptide-Polymer Conjugation Via Copper-Catalyzed Alkyne-Azide 1,3-Dipolar. . .

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11. Thermometer. 12. Filter (0.45 μm). 2.2 Self-Assembly of Peptide-Conjugate 3

1. Scintillation vial (20 mL). 2. Scintillation vial cap with hole. 3. Magnet (1 cm). 4. Endotoxin-free ultrapure water. 5. Syringe pump. 6. Magnetic stirrer. 7. Long bent needle (~15 cm). 8. Syringe (5 mL).

2.3 Purification of Peptide-Conjugate 3

1. Pierce Snakeskin dialysis bag, MWCO 3 K (see Note 1). 2. Endotoxin-free ultrapure water. 3. 1 L beaker. 4. Two clamps. 5. Piece of sponge (~20 cm2). 6. Magnetic stirrer. 7. Magnet (~10 cm). 8. Aluminum foil. 9. 10
Phosphate-buffered saline (PBS).

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Methods

3.1 Synthesis of Peptide-Conjugate 3

1. In a dry, 2 mL round-bottom flask, dissolve a mixture of Nterminus 8Qmin azide (1) (9.0 mg, 4.8 μmol) and poly(t-butyl acrylate) (2) (5.7 mg, 0.3 μmol) in DMF (1 mL). 2. Add copper wires (60 mg) to the reaction mixture (see Note 2). 3. Partially remove the air in the reaction mixture by bubbling a nitrogen throw solution with the help of nitrogen-filled balloon for 15 s (see Note 3). 4. Cover the reaction mixture with aluminum foil to protect it from light and stir at 50  C under nitrogen atmosphere. 5. Stop the reaction after 12 h (see Note 4). 6. Filter off the wires from the warm solution using a 0.45 μm filter and wash with 1 mL of DMF.

3.2 Self-Assembly of Peptide-Conjugate 3

1. Load the DMF filtrate into a 5 mL syringe. 2. Attach a long bent needle to the syringe (Fig. 2) (see Note 5).

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Fig. 2 Instrument assembly to produce 3 in particle form using solvent exchange

3. Mount the syringe into a syringe pump and adjust the rate of addition to 0.5 mL/h (see Note 6). 4. Add 4 mL endotoxin-free water into a 20 mL scintillation vial charged with a 1 cm magnet; cover using a cap with a hole (see Note 7). 5. Insert the bent needle through the hole in the cap and adjust the needle so that its tip touches the surface of the water, close to the middle of the vial. 6. Add the DMF solution to the endotoxin-free water at a rate of 0.5 mL/h while stirring the mixture at 250 rpm (see Note 8). 3.3 Purification of Peptide-Conjugate 3

1. Transfer the above mixture to an 8 cm dialysis bag (see Note 9). 2. Dialyze the solution against 1 L endotoxin-free water (pH 6.8) to clear away copper salts and unreacted peptides (see Note 10). 3. Quantify the concentration of conjugate 3 by freeze-drying half of the purified solution and measuring the mass of the resulting dry solid. 4. Use the freeze-dried product to perform elemental analysis and calculate the substitution ratio (see Note 11). 5. Adjust the concentration of 3 to 1 mg per 900 μL of water by evaporating the water using a stream of nitrogen. 6. Add 10
PBS (100 μL) to the 900 μL 3/water solution to adjust the concentration of 3 to 1 mg/mL in 1
PBS. 7. Measure particle size using a laser particle size analyzer (e.g., Mastersizer 2000) (see Note 12). 8. Determine the copper content using inductively coupled plasma optical emission spectroscopy (see Note 13). 9. Store the particle 3 solution at room temperature (see Note 14).

Peptide-Polymer Conjugation Via Copper-Catalyzed Alkyne-Azide 1,3-Dipolar. . .

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Notes 1. The separation characteristic most commonly defined by the pore size range of a dialysis membrane is the membrane’s molecular weight cutoff (MWCO). Dialysis bag selection is, therefore, based on the molecular weight of the peptide in focus. The MWCO of the dialysis bag must be larger than the molecular weight of the peptide and needs to be changed when peptides with different molecular weights are used. 2. Ensure that the used copper wires are shiny and pinkishorange; otherwise, the wires need to be treated with concentrated sulfuric acid (1 min), washed with distilled water, then methanol, and dried under reduced pressure. 3. The complete removal of air can cause a substantial delay or cease the reaction as a small amount of oxygen is needed to oxidize copper metal into copper I; this is essential for catalyzing the CuAAC reaction. In contrast, an excessive amount of oxygen can trigger the undesired formation of large amounts of copper II. 4. After 12 h, the reaction mixture changes from colorless to green (or bluish green), indicating the occurrence of the reaction. If the reaction mixture remains transparent, extend the reaction time. 5. The needle has to be washed three times with chloroform and dried before use. 6. The rate of addition of DMF to water needs to be adjusted to complete self-assembly in 3–4 h. The needle should not touch the wall of vial. 7. The 20 mL scintillation vial and 1 cm magnet need to be washed three times with endotoxin-free water before use. 8. Ensure that the magnet is rotating in the middle of the solution and not touching the wall of the vial. 9. The dialysis bag needs to be soaked and rinsed using endotoxin-free water before use. The bag should then be folded from the bottom twice. After loading the self-assembled solution into the bag using a 1 mL tip, fold the top of the bag twice, clamp it from both ends, and attach it to a piece of sponge (to allow dialysis bag submerge in water but not sink to the bottom of the flask). 10. Change the dialysis water 3
per day and continue dialysis for 3 days. 11. Compare the obtained nitrogen/carbon ratio with theoretical values for compound 3 by drawing a theoretical nitrogen/ carbon ratio versus substitution rate curve. For example, 0%

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substitution (N/C ¼ 0.020, value for polymer 2 alone), 100% substitution (N/C ¼ 0.124) due to the existence of a nitrogenrich peptide 1 in the peptide-polymer conjugate 3. 12. A milky suspension is formed upon addition of PBS (pH 7.4). The polymer-peptide conjugate 3 formed large particles of 13 μm in diameter. The size distribution of conjugate 3 was narrow (span ¼ 1.7). Replacement of 2 with a more hydrophilic peptide will produce a clearer solution and smaller particle size, in which case, the use of a dynamic light scatteringbased technique (e.g., with the help of a Zetasizer) is recommended. Utilizing water (e.g., for intranasal vaccine delivery) as a solubilizing solvent (instead of PBS) will have a significant influence on particle size. 13. The copper content in the conjugate was below 0.1 ppm (the recommended health standard level of copper is 15 ppm). 14. The final formulation needs to be freshly prepared before biological examination. It should not be stored in the fridge, as the particles tend to precipitate at low temperature. If necessary, 3 can be lyophilized and encapsulated inside a liposomal formulation [7], then it can be freeze-dried and kept long-term at 20  C prior to encapsulation. References 1. Boas U, Heegaard PM (2004) Dendrimers in drug research. Chem Soc Rev 33:43–63 2. Heegaard PMH, Boas U, Sorensen NS (2010) Dendrimers for vaccine and immunostimulatory sses. A review. Bioconjug Chem 21:405–418 3. Zhao GZ, Chandrudu S, Skwarczynski M, Toth I (2017) The application of selfassembled nanostructures in peptide-based subunit vaccine development. Eur Polym J 93:670–681 4. Hilgers LA, Nicolas I, Lejeune G, Dewil E, Strebelle M, Boon B (1998) Alkyl-esters of polyacrylic acid as vaccine adjuvants. Vaccine 16:1575–1581 5. Hilgers LA, Ghenne L, Nicolas I, Fochesato M, Lejeune G, Boon B (2000) Alkyl-polyacrylate esters are strong mucosal adjuvants. Vaccine 18:3319–3325 6. Faruck MO, Zhao L, Hussein WM, Khalil ZG, Capon RJ, Skwarczynski M, Toth I (2020) Polyacrylate-peptide antigen conjugate as a single-dose oral vaccine against group A streptococcus. Vaccines (Basel) 8:23 7. Khongkow M, Liu T-Y, Bartlett S, Hussein WM, Nevagi R, Jia Z, Monteiro MJ, Wells J, Ruktanonchai UR, Skwarczynski M, Toth I

(2018) Liposomal formulation of polyacrylate-peptide conjugate as a new vaccine candidate against cervical cancer. Prec Nanomed 1:186–196 8. Truong HH, Hussein WM, Liu TY, Jia Z, Wells JW, Monteiro MJ, Skwarczynski M, Toth I (2019) Self-adjuvanting peptide vaccines against cervical cancer. Vaccin Res Open J 2:81–89 9. Chandrudu S, Bartlett S, Khalil ZG, Jia ZF, Hussein WM, Capon RJ, Batzloff MR, Good MF, Monteiro MJ, Skwarczynski M, Toth I (2016) Linear and branched polyacrylates as a delivery platform for peptide-based vaccines. Ther Deliv 7:601–609 10. Skwarczynski M, Zaman M, Urbani CN, Lin IC, Jia Z, Batzloff MR, Good MF, Monteiro MJ, Toth I (2010) Polyacrylate dendrimer nanoparticles: a self-adjuvanting vaccine delivery system. Angew Chem Int Ed Engl 49:5742–5745 11. Zaman M, Skwarczynski M, Malcolm JM, Urbani CN, Jia ZF, Batzloff MR, Good MF, Monteiro MJ, Toth I (2011) Self-adjuvanting polyacrylic nanoparticulate delivery system for group A streptococcus (GAS) vaccine. Nanomedicine 7:168–173

Peptide-Polymer Conjugation Via Copper-Catalyzed Alkyne-Azide 1,3-Dipolar. . . 12. Liu TY, Hussein WM, Jia Z, Ziora ZM, McMillan NA, Monteiro MJ, Toth I, Skwarczynski M (2013) Self-adjuvanting polymer-peptide conjugates as therapeutic vaccine candidates against cervical cancer. Biomacromolecules 14:2798–2806 13. Ahmad Fuaad AA, Jia Z, Zaman M, Hartas J, Ziora ZM, Lin IC, Moyle PM, Batzloff MR, Good MF, Monteiro MJ, Skwarczynski M, Toth I (2014) Polymer-peptide hybrids as a highly immunogenic single-dose nanovaccine. Nanomedicine (London) 9:35–43 14. Liu TY, Ahmad Fuaad AA, Toth I, Skwarczynski M (2014) Self-assembled peptide-polymer conjugates as vaccines. Chim Oggi 32:18–22 15. Liu TY, Giddam AK, Hussein WM, Jia ZF, McMillan NAJ, Monteiro MJ, Toth I, Skwarczynski M (2015) Self-adjuvanting therapeutic peptide-based vaccine induce CD8(+) cytotoxic T lymphocyte responses in a murine human papillomavirus tumor model. Curr Drug Deliv 12:3–8

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16. Liu TY, Hussein WM, Giddam AK, Jia Z, Reiman JM, Zaman M, McMillan NA, Good MF, Monteiro MJ, Toth I, Skwarczynski M (2015) Polyacrylate-based delivery system for self-adjuvanting anticancer peptide vaccine. J Med Chem 58:888–896 17. Hussein WM, Liu TY, Jia ZF, McMillan NAJ, Monteiro MJ, Toth I, Skwarczynski M (2016) Multiantigenic peptide-polymer conjugates as therapeutic vaccines against cervical cancer. Bioorg Med Chem 24:4372–4380 18. Purcell AW, McCluskey J, Rossjohn J (2007) More than one reason to rethink the use of peptides in vaccine design. Nat Rev Drug Discov 6:404–414 19. Skwarczynski M, Zaman M, Urbani CN, Lin IC, Jia Z, Batzloff MR, Good MF, Monteiro MJ, Toth I (2010) Polyacrylate dendrimer nanoparticles: a self-adjuvanting vaccine delivery system. Angew Chem Int Ed 49:5742–5745

Chapter 2 Conjugation of Peptides to Gold Nanoparticles Pornsuda Maraming and James Chen Yong Kah Abstract Peptides and proteins have played an important role in many biological processes, functioning as enzymes, hormones, ligands, receptors, cell mediators, and structural components of cells. Being intrinsic molecules in signaling pathways, peptides allow for therapeutic intervention that closely mimic natural signaling cascades. However, the short chain of amino acids in free peptides is susceptible to proteolysis in vivo. Conjugation of peptides onto nanoparticles has been used as a strategy to extend peptide half-life through conferring steric hindrance and a high packing density that prevents proteolytic enzymes to degrade them. Here, we describe a method to conjugate the anticancer p53 peptides as our model peptide onto 12 nm gold nanoparticles (AuNPs) to form the AuNP-p53 peptide conjugate. Conjugation of the p53 short-chain peptide of 25 amino acids occurs through a combination of electrostatic interactions and covalent bonds between cysteine residues at the N-terminal of the peptide and the surface of the AuNPs. The AuNPs and AuNP-p53 are characterized by UV-Vis spectroscopy for its optical absorbance and zetasizer for their hydrodynamic diameter and zeta potential. The semiquantitative analysis of the amount of conjugated peptides on the AuNPs and peptide stability under trypsin treatment is performed on sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Key words Gold nanoparticles, Peptide, Bioconjugation, Peptide stability

1

Introduction Peptides have been widely studied for various biomedical applications including drug delivery [1], peptide-based biosensors [2, 3], disease theranostics [4], and imaging [5]. Among the various types of peptides, the p53 peptide possesses anticancer properties by functioning largely as an antagonist to the p53 protein that is widely studied in cancer research due to its potent tumor suppressive function [6]. Cells normally maintain the steady-state expression of p53 at very low levels by the mouse double minute 2 homologue (MDM2) [7]. MDM2 has E3 ubiquitin protein ligase activity, interacts with the p53 protein, and mediates the ubiquitindegradation pathway for destabilization of p53 [8]. p53 stabilization is primarily obtained via the disruption of MDM2/p53

Waleed M. Hussein, Rachel J. Stephenson and Istvan Toth (eds.), Peptide Conjugation: Methods and Protocols, Methods in Molecular Biology, vol. 2355, https://doi.org/10.1007/978-1-0716-1617-8_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

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interaction. For example, p53 protein is activated by stressinducing signals such as DNA damage, oncogene activation, or hypoxia [9]. Subsequently, p53 is phosphorylated by various kinases at the amino terminus, which prevents MDM2 binding, leading to apoptosis, cell-cycle arrest, senescence, or modulation of autophagy [10]. Therapeutic peptides such as the p53-activating peptide can also disrupt MDM2-p53 protein-protein interactions, which stabilize and activate the p53 proteins for downstream antitumor signaling pathways. Specifically, the p53-activating peptide (residues 361–382 in the p53 C-terminus region) with a net charge of +10, isoelectric point of 11.88, and molecular weight of 2650 Da could interact antagonistically with MDM2 and prevents degradation of p53 [11]. While effective in vitro, these free peptide drugs usually suffer from rapid proteolysis in a biological environment, thus reducing their bioavailability at the target site [12]. Among the various strategies studied to improve the stability of these peptides, their conjugation to nanoparticles has shown promise to enhance their stability against proteolytic degradation. These nanoparticle-peptide conjugates have not only improved the peptide stability and functionality [11], but also facilitated targeted imaging [13], specific intracellular delivery [14], and diagnosis of disease [15]. Among the various types of nanoparticles, gold nanoparticles (AuNPs) are an effective carrier for therapeutic peptides. The gold surface presents a high affinity to thiol and amine functional groups commonly found in proteins and peptides to form strong coordinate bonds. With appropriate conjugation protocol that achieves a high packing density of peptides on the surface of AuNPs, these peptides could be protected from proteolytic digestion [16] by preventing the access of proteases to the peptide through the steric hindrance offered by the high packing density [11]. Such a strategy to offer ligand protection against degradation through conjugation to nanoparticles has also been demonstrated in protecting negatively charged DNA from degradation by nuclease [17]. This chapter describes a method of synthesizing AuNPs, the process of conjugating p53-activating peptides on AuNPs, as well as the characterization and quantification of the resulting AuNPp53 peptide conjugates. We also described the test to assess the improved stability of p53 peptides under proteolytic treatment by trypsin following conjugation to AuNPs.

2 2.1

Materials Chemicals

Prepare all solutions of analytical grade reagents using MilliQ water with a resistivity of 18.2 MΩ cm at 25
C. Otherwise stated, all reagents used in the preparation were used as received. All waste

Conjugation of Peptides to Gold Nanoparticles

11

disposal regulations were strictly adhered to when disposing waste materials. 1. Tetrachloroauric (III) acid trihydrate (HAuCl4 ·3H2O, Mw ¼ 393.83 g/mol). 2. 1% sodium citrate tribasic dihydrate: dissolve 0.2 g sodium citrate tribasic dihydrate in 20 mL ultrapure (UP) water. 3. p53 peptide (C-GG-GSRAHSSHLKSKKGQSTSRHKK). 4. 1 M hydrochloric acid (HCl): slowly add 8.292 mL of 37% (w/w) HCl solution to 25 mL UP water. Adjust the final volume of solution to 100 mL with UP water. 5. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). 6. Bio-Safe™ Coomassie blue. 7. Novex 4–12% tris-glycine mini protein gels. 8. 5 loading buffer. 9. 0.25% W/V trypsin-EDTA. 10. Tris-glycine running buffer. 11. 50 mM HAuCl4: 394 mg of HAuCl4·3H2O dissolved in 20 mL UP water in a 50 mL tube (see Note 1). Store the stock solution at 4
C until use. 12. 1 mM HAuCl4: 2 mL of 50 mM HAuCl4 dissolved into 98 mL UP water in the Erlenmeyer flask and stirring constantly at 220
C and 700 rpm on the hot plate. 2.2 Labware and Equipment

1. 200 mL Erlenmeyer flask. 2. Magnetic stir bar. 3. Hot plate. 4. 1.5 mL microcentrifuge tubes. 5. 96-microplate well. 6. Platform shaker. 7. Centrifuge. 8. UV-Vis spectrometry. 9. Zetasizer. 10. Transmission electron microscopy. 11. Vertical electrophoresis cell.

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Methods

3.1 Preparation of AuNPs

1. While 1 mM HAuCl4 is boiling, reduce temperature to 180
C and quickly add 15 mL of a 1% solution of trisodium citrate dihydrate and allow it to turn deep red for 15 min. 2. Remove from heat and cool down to room temperature without stirring. 3. Transfer 1 mL of AuNP solution to each 1.5 mL microcentrifuge tube. 4. Centrifuge at 6,500  g for 20 min to remove excess citrate. 5. Discard supernatant and resuspend each sample with 1 mL of UP water. 6. Repeat steps 6 and 7. 7. Store at 4
C before use.

3.2 Conjugation of p53 Peptides on AuNPs

1. Add 84 μL of UP water to a 1.5 mL microcentrifuge tube. 2. Add 16 μL of 1 M HCl and 100 μL of 200 μM p53 peptide to the above tube (see Note 2). 3. Add 200 μL of 10 nM AuNP to the above mixture while vortexing to allow self-assembly and conjugation of the p53 peptides on AuNPs. This provides a peptide/AuNP incubation ratio of 20,000. To find out optimal condition, vary the volumes of 1 M HCl (0 μL, 4 μL, 8 μL, 12 μL, and 16 μL) and the peptide concentrations (50 μM, 100 μM, and 200 μM) (see Note 3). 4. Mix continuously on a platform shaker at 180 rpm for 3 h at 4
C. 5. Wash the resulting AuNP-p53 by repeated centrifugation twice with UP water at 6,500  g for 20 min to remove unbound p53 peptide. 6. Reconstitute the pellet with UP water to its initial volume of 400 μL and store at 4
C before use.

3.3 Characterization of AuNPs

1. Measure absorbance spectra of AuNPs and AuNP-p53 using UV-Vis spectroscopy by transferring 300 μL of AuNPs and 300 μL of AuNP-p53 into 96-microplate wells. Measure the absorbance spectrum over a wavelength of 400–900 nm. Determine the concentration of AuNPs from their absorbance at 450 nm (see Note 4). 2. Measure the hydrodynamic diameter and zeta potential of AuNPs using Zetasizer using instructions and protocols provided by the device manufacturer (Fig. 1).

Conjugation of Peptides to Gold Nanoparticles

13

Fig. 1 Characterization of synthesized citrate-capped AuNPs and AuNP-p53. (a) Citrate-capped AuNPs were monodispersed as imaged under TEM and (b) had an average diameter of 12.12  0.13 nm. (c) UV-Vis absorption spectra showed a red-shifted absorbance peak of AuNPs from 520 nm to 525 nm after conjugation with p53 peptides to form AuNP-p53, indicating successful conjugation. (d) Dynamic light scattering analysis showed monomodal size distribution of AuNPs and AuNP-p53 with an increased size and no large aggregation upon conjugation of p53 peptides. (Figure and caption adapted with permissions from Chan et al. Bioconjugate Chemistry 2019; 30(3): 920–930)

3. Examine AuNPs under transmission electron microscopy (TEM) (Fig. 1) (see Note 5). 3.4 Semiquantitative Measurement of the Amount of Peptides Conjugated on the AuNPs

1. Heat a 10 μL of 30 nM AuNP-peptide in 10 μL of 5 loading buffer at 100
C for 5 min (see Note 6). 2. Insert a polyacrylamide gel into a vertical electrophoresis cell. 3. Load 10 μL of peptide samples and run on SDS-PAGE at 100 V for 120 min in tris-glycine running buffer (see Note 7). 4. Rinse the gel with 200 mL UP water three times for 5 min each to remove SDS and buffer, which will interfere with the staining. 5. Remove all the water from the staining container and stain with 50 mL Bio-Safe™ Coomassie blue (or enough to completely cover the gel) for 30 min on shaker. 6. Destain with 200 mL UP water by gently shaking until background is clear. Stained gel can be stored in water. 7. Image the gel and quantify each peptide band in the picture using ImageJ software (National Institutes of Health) based on the density of the profile blot. A darker signal indicates higher

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Pornsuda Maraming and James Chen Yong Kah

density of peptides. The density of peptide band was compared against calibrating standards of known peptide concentrations to calculate the number of conjugated peptides per AuNP. 3.5 Stability of Conjugated p53 Peptides on AuNPs

1. Prepare a half serial dilution of stock 0.25–0.0020% W/V trypsin-EDTA and a blank control without trypsin in 1.5 mL microcentrifuge tubes (see Note 8). 2. A 10 μL of 30 nM of AuNP-p53 or 15 μM of free p53 peptides at equivalent concentration (based on our determined 500 p53 peptide molecules conjugated on each AuNP, as determined in Subheading 3.4) is added to 10 μL of trypsin over a range of concentrations (0, 0.0020, 0.0039, 0.0078, 0.0156, 0.0313, 0.0625, and 0.1250%) and incubated at 37
C for 5 min to digest the p53 peptide. Here, 10 μL of UP water serves at our negative control. 3. Heat the mixture of trypsin with AuNP-p53 or free p53 in 10 μL of 5 loading buffer at 100
C for 5 min to inactivate the trypsin and release all bound peptides from AuNPs. 4. Insert a Novex 4–12% tris-glycine mini protein gel into a vertical electrophoresis cell. 5. Load 10 μL of peptide samples into the gel and run the gel at 100 V for 120 min in tris-glycine running buffer. 6. Stain the peptide band by adding 50 mL of Bio-safe™ Coomassie blue as described in Subheading 3.4. 7. Image the gel and analyze peptide band intensity by using ImageJ software as described in Subheading 3.4 above. Calculate the percentage of intact p53 protein according to the formula: (density of peptide band of trypsin treatment/density of peptide band of negative control)  100.

4

Notes 1. Tetrachloroauric (III) acid trihydrate is very hygroscopic, so weighing must be done quickly to minimize inaccuracy of weight due to the absorbing water. The compound should be stored tightly closed and protected from light. 2. Prepare 200 μM of p53 peptide by dissolving 0.53 mg of p53 peptide in 1 mL of UP water. 3. AuNPs lose their colloidal stability during conjugation with p53 peptides in UP water. When p53 peptides are at a pH below its peptide isoelectric point of 11.88, the peptides carry a net positive charge, which promotes peptide binding on negatively charged AuNP surfaces. Therefore, using a high peptide conjugation ratio and optimal HCl volume maintains

Conjugation of Peptides to Gold Nanoparticles

15

colloidally stability of AuNPs after peptide conjugation. The optimal condition for conjugation is the HCl and peptide concentration giving colloidal stability of AuNPs with narrow particle size distribution, polydispersity index 95%. H NMR (600 MHz, DMSO-d6): δ 8.93 (s, 1H, H2_His), 8.46 (d, J ¼ 7.3 Hz, 1H, NH_Ala5), 8.11–8.09 (m, 2H, NH_Gly + NH_His), 8.07 (d, J ¼ 7.0 Hz, 1H, NH_Ala2), 7.99 (d, J ¼ 8.0 Hz, 1H, NH_4-F-Phe), 7.29–7.27 (m, 3H, H5_His + Hδ_4-F-Phe), 7.07–7.04 (m, 2H, Hε_4-FPhe), 4.53–4.49 (m, 2H, Hα_4-F-Phe + Hα_His), 4.26–4.20 (m, 2H, Hα_Ala5 + Hα_ Ala2), 3.68 (d, J ¼ 5.7 Hz, 2H, Hα_Gly), 3.06–3.00 (m, 2H, Hβ_4-FPhe + Hβ_His), 2.90 (dd, J ¼ 15.3, 8.2 Hz, 1H, Hβ_His), 2.78 (dd, J ¼ 14.0, 9.4 Hz, 1H, Hβ_4-F-Phe), 1.85 (s, 3H, CH3CO_Ac), 1.30 (d, J ¼ 7.3 Hz, 3H, Hβ_Ala5), 1.13 (d, J ¼ 7.1 Hz, 3H, Hβ_Ala2).

1

C NMR (150 MHz, DMSO-d6): δ 173.9 (CO2H_Ala5), 172.2 (CONH_Ala2), 170.6, 169.9, 169.5 (CONH_X + CONH_4-F-Phe + COCH3_Ac), 169.0 (CONH_Gly), 161.0 (d, 1JCF ¼ 241.7 Hz, Cζ_4-F-Phe), 133.7 (C2_His), 133.6 (d, 4JCF ¼ 2.8 Hz, Cγ_4-F-Phe), 131.1 (d, 3JCF ¼ 8.0 Hz, Cδ_4-F-Phe), 129.1 (C4_His), 116.9 (C5_His), 114.7 (d, 2JCF ¼ 21.0 Hz, Cε_4-F-Phe),

13

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Anaı¨s F. M. Noisier and Ranganath Gopalakrishnan

53.6 (Cα_4-F-Phe), 51.4 (Cα_His), 48.2 (Cα_Ala2), 47.6 (Cα_Ala5), 42.0 (Cα_Gly), 36.6 (Cβ_4-F-Phe), 26.9 (Cβ_His), 22.5 (COCH3_Ac), 17.7 (Cβ_Ala2), 17.1 (Cβ_Ala5). F NMR (471 MHz, DMSO-d6): δ 116.85 (tt, 3 JHF ¼ 9.0 ppm, 4JHF ¼ 5.6 ppm).

19

HR-MS (ESI+) (m/z): [M + H]+ calculated C25H33FN7O7, 562.2425; found 562.2451.

for

12. Store the peptide at 20  C until further use. 3.2 LSF of Unprotected Peptide at Histidine

1. Place a magnetic stir bar in a conical microwave vial. Charge the crude peptide from Subheading 3.1 (0.05 mmol; 28 mg; 1 equiv.) and MeCF2SO2Na (0.6 mmol; 91 mg; 12 equiv.) in the vial. Add 500 μL of a solution of H2O/DMSO (1/1, v/v) and seal the vial with a pre-pierced septum (see Notes 3–6). 2. Prepare 600 μL of a solution of TBHP in H2O/DMSO: by mixing 168 μL of TBHP (70% TBHP in H2O), 216 μL of H2O, and 216 μL of DMSO in a vial. Draw up the solution into a male luer lock plastic syringe connected to a female luer lock PTFE tubing. Ensure to push air out of the syringe and tubing (see Notes 7 and 8). 3. Insert the PTFE tubing in the vial through the pierced septum and secure the syringe onto the syringe pump. Set the appropriate syringe diameter. 4. Slowly add the TBHP solution (1.0 mmol; 20 equiv.) at a flow rate of 125 μL/h for 4 h to the reaction mixture stirring at RT (see Notes 9–11). 5. Monitor the reaction by UPLC-MS (see Notes 12 and 13). 6. Upon completion, quench the reaction by slowly adding a 0.1 M NH4HCO3 solution (pH 7) until gas evolution ceased, dilute with H2O/MeCN (1/1, v/v) + 0.1% TFA, and freezedry (see Note 14). 7. Purify the crude peptide by reverse-phase HPLC (gradient: 5% purification solvent B for 1 min, 5–15% B in 3 min, 15–20% B in 15 min, Atlantis T3 column). Freeze-dry pure fractions to afford the desired peptide as a white fluffy solid. Yield (4.5 mg, 15%, over 11 steps from resin loading). 8. The product was analyzed by analytical UPLC-MS, 1H, 13C, 19 F NMR, and HR-MS. UPLC-MS Rt ¼ 3.5 min (3–60% analytical solvent B over 10 min, λ ¼ 210 nm), purity >95%. 1

H NMR (600 MHz, DMSO-d6): δ 8.29 (d, J ¼ 6.7 Hz, 1H, NH_Ala5), 8.08 (t, J ¼ 5.5 Hz, 1H, NH_Gly), 8.05 (d, J ¼ 7.1 Hz, 1H, NH_Ala2), 8.01 (d, J ¼ 7.9 Hz, 1H,

Oxime/Hydrazone Conjugation at Histidine: Late-Stage Functionalization. . .

43

NH_X), 7.84 (d, J ¼ 8.2 Hz, 1H, NH_4-F-Phe), 7.27–7.25 (m, 2H, Hδ_4-F-Phe), 7.06–7.03 (m, 2H, Hε_4-F-Phe), 6.83 (bs, 1H5_X), 4.49 (td, J ¼ 8.5, 4.6 Hz, 1H, Hα_4-F-Phe), 4.40–4.37 (m, 1H, Hα_X), 4.24–4.19 (m, 2H, Hα_Ala2 + Hα_Ala5), 3.72–3.64 (m, 2H, Hα_Gly), 3.03 (dd, J ¼ 13.9, 4.3 Hz, 1H, Hβ_4-FPhe), 2.89–2.87 (m, 1H, Hβ_X), 2.79–2.73 (m, 2H, Hβ_4-F-Phe + Hβ_X), 1.99 (t, 3JHF ¼ 18.7 Hz, 3H, CF2CH3_X), 1.84 (s, 3H, CH3CO_Ac), 1.28 (d, J ¼ 7.3 Hz, 3H, Hβ_Ala5), 1.13 (d, J ¼ 7.1 Hz, 3H, Hβ_Ala2). C NMR (150 MHz, DMSO-d6): δ 173.8 (CO2H_Ala5), 172.0 (CONH_Ala2), 170.4, 170.4 (CONH_X + CONH_4-F-Phe), 169.7 (COCH3_Ac), 168.8 (CONH_Gly), 160.9 (d, 1JCF ¼ 241.7 Hz, Cζ_4F-Phe), 133.5 (d, 4JCF ¼ 2.1 Hz, Cγ_4-F-Phe), 131.0 (d, 3 JCF ¼ 8.0 Hz, Cδ_4-F-Phe), 114.6 (d, 2JCF ¼ 20.9 Hz, Cε_4-F-Phe), 53.4 (Cα_4-F-Phe), 52.5 (Cα_X), 48.2 (Cα_Ala2), 47.5 (Cα_Ala5), 42.0 (Cα_Gly), 36.5 (Cβ_4-FPhe), 22.8 (t, 2JCF ¼ 26.1 Hz, CH3CF2_X), 22.3 (COCH3_Ac), 17.8 (Cβ_Ala2), 17.0 (Cβ_Ala5).

13

Signals too weak to be assigned by 13C NMR: C2_X visible at 140 ppm by HMBC and CF2_X visible as a triplet at 118 ppm by HMBC. Cβ_X, C5_X and C4_X are not seen. 83.15

(q,

HRMS (ESI+) (m/z): [M + H]+ calculated C27H35F3N7O7, 626.2550; found 626.2562.

for

19

F

3

NMR (471 MHz, DMSO-d6): δ JHF ¼ 19.1 Hz, 2F), 116.96 (bs, 1F).

9. Store the peptide at 20  C until further use. 3.3 Conversion of the Difluoroethyl Group into a Conjugation Handle

1. Place the peptide (7.2 μmol; 4.5 mg) from Subheading 3.2 in a vial equipped with a magnetic stir bar, dissolve it in 1 mL of a solution of MeCN/H2O (1/1, v/v). Add 1 mL of a saturated solution of NaHCO3 (pH 8–9) and stir at RT for 10 min (see Note 15). 2. Monitor the reaction by UPLC-MS. 3. Freeze-dry the solution. 4. Desalt the peptide using a C18 cartridge: Prepare the column by eluting MeOH (5 column volume), equilibrate the column by eluting H2O + 0.1% TFA (5 column volume), dissolve the peptide in minimal volume of H2O + 0.1% TFA and load it on the column, elute with H2O + 0.1% TFA (5 column volume), then elute with a linear gradient of 5–50% of MeCN in H2O + 0.1% TFA over 5 column volume. Analyze the fractions by UPLC-MS and freeze-dry the relevant fractions to

44

Anaı¨s F. M. Noisier and Ranganath Gopalakrishnan

afford the desired peptide as a white fluffy solid. Yield (4.3 mg, quantitative). 5. The product was analyzed by analytical UPLC-MS, 1H, 13C NMR, and HR-MS. UPLC-MS Rt 3.2 min (3–43% analytical solvent B over 10 min, λ ¼ 210 nm), purity >95%. 1

H NMR (600 MHz, DMSO-d6): δ 8.30 (d, J ¼ 6.5 Hz, 1H, NH_Ala5), 8.08 (t, J ¼ 5.7 Hz, 1H, NH_Gly), 8.05–8.01 (m, 2H, NH_Ala2 + NH_X), 7.84 (d, J ¼ 8.3 Hz, 1H, NH_4-F-Phe), 7.27–7.25 (m, 2H, Hδ_4-F-Phe), 7.06–7.03 (m, 2H, Hε_4-F-Phe), 6.99 (bs, 1H5_X), 4.49 (td, J ¼ 8.6, 4.5 Hz, 1H, Hα_4-F-Phe), 4.44–4.41 (m, 1H, Hα_X), 4.25–4.19 (m, 2H, Hα_Ala2 + Hα_Ala5), 3.71–3.64 (m, 2H, Hα_Gly), 3.02 (dd, J ¼ 13.9, 4.3 Hz, 1H, Hβ_4-F-Phe), 2.93 (dd, J ¼ 14.9, 4.2 Hz, 1H, Hβ_X), 2.81–2.76 (m, 2H, Hβ_4-F-Phe + Hβ_X), 2.46 (s, 3H, COCH3_X), 1.84 (s, 3H, CH3CO_Ac), 1.28 (d, J ¼ 7.3 Hz, 3H, Hβ_Ala5), 1.13 (d, J ¼ 7.1 Hz, 3H, Hβ_Ala2). C NMR (150 MHz, DMSO-d6): δ 188.2 (COCH3_X), 173.9 (CO2H_Ala5), 172.1 (CONH_Ala2), 170.4, 170.4 (CONH_X + CONH_4-F-Phe), 169.8 (COCH3_Ac), 168.9 (CONH_Gly), 161.0 (d, 1JCF ¼ 241.7 Hz, Cζ_4F-Phe), 144.3 (C2_X), 133.6 (d, 4JCF ¼ 2.8 Hz, Cγ_4-FPhe), 131.2 (d, 3JCF ¼ 8.0 Hz, Cδ_4-F-Phe), 114.7 (d, 2 JCF ¼ 20.9 Hz, Cε_4-F-Phe), 53.5 (Cα_4-F-Phe), 52.5 (Cα_X), 48.2, 47.6 (Cα_Ala2 + Cα_Ala5), 42.1 (Cα_Gly), 25.3 (COCH3_X), 22.5 36.7 (Cβ_4-F-Phe), (COCH3_Ac), 18.0 (Cβ_Ala2), 17.1 (Cβ_Ala5).

13

Signals too weak to be assigned by 13C NMR: Cβ_X, C4_X and C5_X. F NMR (471 MHz, DMSO-d6): δ 116.96 (tt, 3 JHF ¼ 8.8 ppm, 4JHF ¼ 5.7 ppm).

19

HRMS (ESI+) (m/z): [M + H]+ calculated for C27H35FN7O8, 604.2531; found 604.2536. 6. Store the peptide at 20  C until further use. 3.4 Hydrazone Ligation with a Fluorophore

1. Place the peptide (0.02 mmol; 15 mg, 1 equiv.) from Subheading 3.3 and fluorescein-5-thiosemicarbazide (0.02 mmol; 10 mg; 1 equiv.) in a round-bottom flask, dissolve with 670 μL of a solution of 0.1 M NH4OAc (pH 4.5)/EtOH (1/1, v/v). Stir the reaction mixture at RT overnight (see Notes 16 and 17). 2. Monitor the reaction by UPLC-MS.

Oxime/Hydrazone Conjugation at Histidine: Late-Stage Functionalization. . .

45

3. Upon completion, concentrate the EtOH in vacuo and freezedry the reaction mixture. 4. Purify the crude peptide by reverse-phase HPLC (gradient: 5% purification solvent B for 1 min, 5–26% B in 3 min, 26–31% B in 15 min, Atlantis T3 column). Freeze-dry pure fractions to afford the desired product as a white fluffy solid. Yield (4.7 mg, 19%, over 13 steps from resin loading). 5. The product was analyzed by analytical UPLC-MS and HR-MS. UPLC-MS Rt 2.3 min (20–60% analytical solvent B over 10 min, λ ¼ 210 nm), purity >95%. HR-MS (ESI+) (m/z): [M + H]+ calculated C48H48FN10O12S, 1007.3158; found 1007.3140.

4

for

Notes 1. Peptide synthesis using manual Fmoc-SPPS is described here; however, automated SPPS protocols may be utilized to expedite the synthesis of the peptide substrate. 2. A simple filtration step was carried out as the UPLC-MS analysis of the model peptide indicated that the crude purity was >95%, but reverse-phase HPLC purification could be carried out instead to yield the pure model peptide. 3. The LSF method can be employed directly on the crude peptide, however, it is recommended to carry out at least a filtration step as described in Subheading 3.1, step 10. 4. The LSF methodology can be applied to unprotected peptides of various complexity spanning over a wide range of molecular weight. While the reaction could be carried out at a final peptide concentration of [50 mM] for the short model pentapeptide described herein, reactions could be diluted up to 10 to ensure the solubility of higher MW peptides, thus leading to a final peptide concentration of [5 mM]. However, it is essential to retain a high concentration with respect to the radical precursor and TBHP in the reaction medium. Thus, the final sulfinate and TBHP concentrations were kept at [0.6 M] and [1 M], respectively, regardless of the peptide concentration or equivalent ratios to the peptide substrate. 5. Zinc sulfinate salts can equally be used without adjustment to the protocol. 6. The reaction can be carried out in H2O alone; if both the peptide and the radical precursor are readily soluble without addition of DMSO.

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7. Only 500 μL of the TBHP solution in H2O/DMSO (1/1, v/v) is actually added to the reaction mixture which corresponds to 1.0 mmol; 140 μL; 20 equiv. of TBHP (70% TBHP in H2O); however, 600 μL are prepared and drawn up into the syringe. The additional 20% (100 μL) account for the syringe and PTFE tubing dead volume. 8. PTFE tubing is used instead of metallic needles as TBHP might be decomposed by metal [9]. 9. Slow addition of TBHP results in greatly improved conversions and yields. Rapid addition of TBHP causes a nonproductive consumption of the sulfinate reagents [19]. 10. Among the sulfinate salts tested, only MeCF2SO2Na and (CHF2SO2)2Zn were found to readily react at room temperature. Reactions with other sulfinate reagents required gentle heating of the reaction mixture from 50–70  C. It was hypothesized that while fluoroalkyl radicals are readily generated under oxidative conditions through the loss of SO2 and undergo facile addition to the protonated imidazole ring at room temperature, non-fluorinated sulfinates might require higher temperatures to promote radical formation and/or Minisci-type reactions. For reactions which require heating, slow addition of the TBHP solution was necessary to avoid its decomposition to methyl radicals and the formation of methylated side-products. Caution. Pressure can build up in the heated vial and connected syringe fitted with PTFE tubing. To avoid hazardous splashes and undesirable leaks, it is essential to secure the tight connection of the PTFE tubings to the syringe via a luer lock. 11. To ensure that the correct amount of oxidant is added, it is important to remove air bubbles from the syringe/PTFE tubing, so that the first drop of TBHP is added as soon as the syringe pump is started to respect the duration and rate of slow addition. 12. Reaction analysis may be performed by removing a small aliquot (10–20 μL) of the reaction mixture with a micropipette equipped with plastic tips and diluting in a mixture of MeCN/ H2O (1/1, v/v). The use of metallic needle to sample the reaction should be avoided as TBHP might be decomposed by metal. 13. If the reaction is not complete after 4 h, the stirring and heating can be prolongated after the slow addition is over. For primary alkyl radicals, a second addition of sulfinate reagents can be necessary to reach a good yield. 14. It is possible to skip the isolation of the difluoroalkylsubstituted His-containing peptides and perform the LSF and

Oxime/Hydrazone Conjugation at Histidine: Late-Stage Functionalization. . .

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conversion to the ketone in a one-pot manner by substituting the 0.1 M NH4HCO3 (pH 7) quenching solution described in Subheading 3.2, step 6 with a saturated solution of NaHCO3 (pH 8–9); it is then recommended to carry out a desalting step as described in Subheading 3.3, step 4 prior to purification by RP-HPLC. 15. Formation of the ketone handle occurs through hydrolysis of the gem-difluoroalkyl-substituted His-containing peptides under basic pH. The gem-difluoromethyl-substituted His was not found to hydrolyze to the corresponding aldehyde. 16. Hydrazines can be replaced by alkoxyamines in an oxime ligation reaction. No further changes in the protocol are required. 17. Although the rate of reaction was found to be considerably enhanced at pH 4.5, other organic solvent/buffer mixtures can be used if the substrates are not soluble or may degrade at pH 4.5. References 1. Cernak T, Dykstra KD, Tyagarajan S, Vachal P, Krska SW (2016) The medicinal chemist’s toolbox for late stage functionalization of drug-like molecules. Chem Soc Rev 45:546–576 2. Valeur E, Gueret SM, Adihou H, Gopalakrishnan R, Lemurell M, Waldmann H, Gossmann TN, Plowright AT (2017) New modalities for challenging targets in drug discovery. Angew Chem Int Ed 56:10294–10323 3. Boutureira O, Bernardes GJL (2015) Advances in chemical protein modification. Chem Rev 115:2174–2195 4. Hoyt EA, Cal PMSD, Oliveira BL, Bernardes GJL (2019) Contemporary approaches to siteselective protein modification. Nat Rev Chem 3:147–171 5. Noisier AFM, Brimble MA (2014) C-H functionalization in the synthesis of amino acids and peptides. Chem Rev 114:8775–8806 6. Wang W, Lorion MM, Shah J, Kapdi AR, Ackermann L (2018) Late-stage peptide diversification by position-selective C-H activation. Angew Chem Int Ed 57:14700–14717 7. Sengupta S, Mehta G (2017) Late stage modification of peptides via C-H activation reactions. Tetrahedron Lett 58:1357–1372 ˜ o O (2018) 8. Brandhofer T, Garcı´a Manchen Site-selective C-H bond activation/functionalization of alpha-amino acids and peptide-like derivatives. Eur J Org Chem 2018:6050–6067

9. deGruyter JN, Malins LR, Baran PS (2017) Residue-specific peptide modification: a chemists guide. Biochemistry 56:3863–3873 10. Malins LR (2018) Peptide modification and cyclization via transition-metal catalysis. Curr Opin Chem Biol 46:25–32 11. Malins LR (2018) Decarboxylative couplings as versatile tools for late-stage peptide modifications. Pept Sci 110(3):e24049 12. Bottecchia C, Noel T (2019) Photocatalytic modification of amino acids, peptides, and proteins. Chem Eur J 25:26–42 13. Liu JQ, Shatskiy A, Matsuura BS, K€ark€as MD (2019) Recent advances in photoredox catalysis enabled functionalization of α-amino acids and peptides: concepts, strategies and mechanisms. Synthesis 51:2759–2791 14. Ji Y, Brueckl T, Baxter RD, Fujiwara Y, Seiple IB, Su S, Blackmond DG, Baran PS (2011) Innate C-H trifluoromethylation of heterocycles. Proc Natl Acad Sci U S A 108:14411–14415 15. Smith JM, Dixon JA, deGruyter JN, Baran PS (2019) Alkyl sulfinates: radical precursors enabling drug discovery. J Med Chem 62:2256–2264 16. Ichiishi N, Caldwell JP, Lin M, Zhong W, Zhu X, Streckfuss E, Kim HY, Parish CA, Krska SW (2018) Protecting group free radical C-H trifluoromethylation of peptides. Chem Sci 9:4168–4175 17. Imiolek M, Karunanithy G, Ng WL, Baldwin AJ, Gouverneur V, Davis BG (2018) Selective

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radical trifluoromethylation of native residues in proteins. J Am Chem Soc 140:1568–1571 18. Noisier AFM, Johansson MJ, Knerr L, Hayes MA, Drury WJ III, Valeur E, Malins LR, Gopalakrishnan R (2019) Late-stage functionalization of histidine in unprotected peptides. Angew Chem Int Ed 58:19096–19102

19. O’Brien AG, Maruyama A, Inokuma Y, Fujita M, Baran PS, Blackmond DG (2014) Radical C-H functionalization of heteroarenes under electrochemical control. Angew Chem Int Ed 53:11868–11871

Chapter 5 Modification of Nanoparticles with Transferrin for Targeting Brain Tissues Sara Aly Attia, Xiang Li, Nina Filipczak, Daniel F. Costa, and Vladimir P. Torchilin Abstract The delivery of therapeutics to brain tissues is one of the main challenges in neuropathology. For the past two decades, a variety of drug delivery systems has been designed to target components of the blood-brain barrier, including the transferrin receptor, a transmembrane glycoprotein highly expressed in the brain endothelium. In this protocol, we describe the use of transferrin protein to activate the surface of nanoparticles with the aim to direct their uptake in the brain. The molecule is bound by an amide linker to a PEGylated lipid commonly used in the preparation of lipid nanoparticles, micelles, and liposomes. Key words Transferrin, Micelles, Nanoparticles, Brain, Targeting

1

Introduction One of the major challenges for effective treatment of diseases in the central nervous system is the ability of molecules to be transported from the bloodstream into the brain. A layer of tightly interconnected cells called blood-brain barrier (BBB) is the main responsible for restricting the delivery of hydrophilic drugs into the brain parenchyma [1]. Since the discovery of the transferrin receptor 1 (TfR1) on the endothelium of brain capillaries in 1984, this transmembrane glycoprotein has become an important effector of targeted delivery of therapeutics to the brain [2]. It binds to an 80 kDa protein, named transferrin, capable of solubilizing ferric iron in the serum and transporting it through the BBB for the maintenance of brain functions, such as neural conductivity [1, 3]. In drug delivery, transferrin bound to the surface of micelles, liposomes, and lipid

Waleed M. Hussein, Rachel J. Stephenson and Istvan Toth (eds.), Peptide Conjugation: Methods and Protocols, Methods in Molecular Biology, vol. 2355, https://doi.org/10.1007/978-1-0716-1617-8_5, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Synthesis of DOPE-PEG-transferrin conjugate. In the first step of the synthesis (a), the terminal amine of the lipid molecule (DOPE) replaces one of the p-nitrophenol end groups of PEG. The same reaction is used to form an amide link between primary amines of transferrin and the remaining pNP terminal group of DOPE-PEG (b)

nanoparticles has been used as an active targeting moiety to improve the distribution of payloads to brain tissues [4–6]. This protocol details the steps for the covalent linkage of transferrin to 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N[amino(polyethylene glycol)-3400 (DOPE-PEG), one of the most common PEGylated phospholipids used in the fabrication of nanoparticles. The reproducibility and modification efficacy of the protocol is 95–96%. The synthesis requires a p-nitrophenol activator for the formation of an amide link between the C-terminus of the PEG chain and the N-terminus of the transferrin protein (Fig. 1). The effectiveness of transferrin-activated nanoparticles was previously confirmed in the treatment of glioblastoma [7, 8].

2

Materials All solutions are prepared using analytical grade reagents and deionized water at room temperature. Store reagents as recommended by the supplier. Waste disposal regulations must be followed to discard any material. 1. DOPE solution: DOPE 25 mg/mL; triethylamine 2% v/v in anhydrous CHCl3 (see Note 1).

Modification of Nanoparticles with Transferrin for Targeting Brain Tissues

2.1 Synthesis of DOPE-PEG3,400p-Nitrophenol (DOPE-PEG3,400-pNP)

51

2. Nitrophenyl Carbonate-PEG-Nitrophenyl Carbonate, MW 3400 (pNP-PEG3,400-pNP) solution: pNP-PEG3,400-pNP 145 mg/mL in anhydrous CHCl3. 3. HCl solution: HCl 0.01 M in water (see Note 2). 4. Thin-layer chromatography (TLC) mobile phase solution: CHCl3:MeOH 80:20 v/v.

2.2 Preparation of pNP-Activated Micelles

1. 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy (polyethylene glycol)-1000 (DSPE-PEG1,000)solution: DSPEPEG1,000 10 mg/mL in anhydrous CHCl3. 2. DOPE-PEG3,400-pNP solution: 2.5 mg/mL in anhydrous CHCl3.

DOPE-PEG3,400-pNP

3. Citrate buffer, pH 6: C6H8O7. H2O 0.24% m/v; C6H5O7Na3. 2 H2O 2.6% m/v in water. Adjust pH with NaOH 0.1 N or HCl 0.1 N. 2.3 Preparation of Transferrin-Activated Micelles

1. Phosphate-buffered saline (PBS) buffer, pH 8: Na2HPO4. 2 H2O 3.4% m/v; NaH2PO4. H2O 0.15% m/v in water. Adjust pH with NaOH 0.1 N or HCl 0.1 N. 2. Transferrin solution: Transferrin 1% m/v in PBS buffer pH 8. 3. pNP-activated micelles solution: product from Subheading 3.2.

3

Methods All methods are carried out at room temperature, unless otherwise specified. Waste disposal regulations must be followed to discard any material.

3.1 Synthesis of DOPE-PEG3,400-pNitrophenol (DOPE-PEG3,400-pNP)

1. Mix 1 mL of DOPE solution with 4 mL of pNP-PEG3,400-pNP solution in an 8 mL glass vial (see Note 3). 2. Pour N2 into the system before closing the vial (see Note 4). 3. Stir for 6 h. 4. Dry the reaction mixture under vacuum in a rotary evaporator for 1 h. 5. Hydrate the remaining lipid film with 2 mL of HCl solution (see Note 5). 6. Purify the product in a sepharose CL-4B size exclusion column. Use HCl solution as mobile phase (see Note 6). Collect 60 aliquots of 2 mL each (see Note 7).

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Fig. 2 Thin-layer chromatography (TLC) analysis of DOPE-PEG3,400-pNP. On the left side of the image, Dragendorff’s reagent was used to stain free pNP-PEG3,400-pNP in orange, and molybdenum blue reagent was used to stain free DOPE in blue. Both components showed distinct retention factors on the silica gel. On the right side of the image, purified DOPE-PEG3,400-pNP showed positive staining for PEG and DOPE with the same retention factor, which validates the conjugation

7. Analyze the aliquots in two silica gel preparative TLC plates. Use TLC mobile phase solution to separate the components of each aliquot. Use Dragendorff’s reagent spray solution to stain plate #1 (see Note 8). Use molybdenum blue reagent spray solution to stain plate #2 (see Note 9). 8. Collect all the aliquots where the orange and blue stains show the same retention factor (Fig. 2) in a 50 mL tube. 9. Freeze the sample at
80  C for 2 h. 10. Freeze-dry the sample overnight (or until complete removal of water). 11. Store the powder (DOPE-PEG3,400-pNP) at
20  C. 3.2 Preparation of pNP-Activated Micelles

1. Transfer 200 μL of DSPE-PEG1,000 and 100 μL of DOPEPEG3,400-pNP solution to a glass vial (see Note 10). 2. Dry the solution of lipids under N2 in a chemical fume hood. 3. Remove traces of chloroform for 2 h in a freeze dryer. 4. Hydrate the remaining lipid film with 2.25 mL of citrate buffer pH 6 (final lipid concentration ¼ 1 mg/mL). 5. Vortex for 5 min or until complete solubilization of the lipid film (see Note 11). 6. Filter the formulation through a 0.2 μm PES filter into a 5 mL sterile tube. 7. Let the formulation stabilize for 30 min.

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8. Check Z-average size and polydispersity index by dynamic light scattering in a Zetasizer Nano Z (see Note 12). 3.3 Preparation of Transferrin-Activated Micelles

1. Mix 1.5 mL of pNP-activated micelles solution with 1.5 mL of transferrin solution in an 8 mL amber glass vial (see Note 13). 2. Adjust the pH to 8 with NaOH 0.1 N, if necessary (see Note 14). 3. Stir for 6 h. 4. Transfer the solution to a pretreated dialysis tubing (MWCO 300,000) (see Note 15). 5. Purify the formulation by dialysis in a beaker filled with 4 L of water for 2 h at 4  C. 6. Replace the water and continue the dialysis overnight at 4  C (see Note 16). 7. Remove the formulation from the dialysis tubing. 8. Filter the formulation through a 0.2 μm PES filter into a 5 mL sterile tube. 9. Let the formulation stabilize for 30 min. 10. Check Z-average size and polydispersity index by dynamic light scattering in a Zetasizer Nano Z (see Note 17). 11. Perform a bicinchoninic acid assay (BCA) to quantify the amount of transferrin on the surface of the micelles (see Note 18).

4

Notes 1. Triethylamine (TEA) must be added to maintain a basic pH in which the reaction happens. 2. Acidic pH protects the p-nitrophenol group from hydrolysis in aqueous solution. 3. pNP-PEG3,400-pNP is added in a fivefold molar excess compared to DOPE. This ratio was optimized to avoid modification of both pNP groups in the molecule of PEG [9]. 4. Creating a N2 atmosphere in the vial avoids degradation of the lipid by O2. 5. The product of the reaction (DOPE-PEG3,400-pNP) is an amphiphilic molecule. It will form micelles with a size over 5 nm in aqueous solution.

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6. The micelles of DOPE-PEG3,400-pNP will elute faster than unreacted components of the mixture based on their size. 7. The number and volume of aliquots will vary depending on the size of the column. For maximum yield, keep collecting aliquots until the product is completely eluted from the column based on the TLC stains. 8. Dragendorff’s reagent stains PEG chains in orange. 9. Molybdenum blue reagent stains the phosphate group from DOPE in blue. 10. The molar ratio between DSPE-PEG1,000 and DOPEPEG3,400-pNP is 95– 5. 11. Both components of the formulation are amphiphilic molecules and will form micelles in aqueous solution. Ultrasound bath can be used to facilitate detachment of the lipid film from the walls of the glass vial. 12. Make sure to also check average size by “number of particles” and “volume.” 13. Transferrin is added in a fivefold molar excess compared to DOPE-PEG3,400-pNP. 14. An amide link between DOPE-PEG3,400-pNP and primary amines of transferrin is formed at slightly basic pH. 15. Unbound transferrin (80 kDa) will be cleared from the formulation through the pores of the dialysis tubing. 16. Dialysis under 4  C will avoid (or slow down) the release of any drug that might have been encapsulated in the core of the micelles. 17. The attachment of transferrin to the surface of the micelles will cause an increase in size (Fig. 3) compared to the starting formulation (from 10 nm to 180 nm). Zeta potential will also change from
7 mV to around
4 mV. Both changes are key to confirm the successful modification of the particle surface. 18. Follow the instructions provided with the BCA kit. Use the solution of pNP-activated micelles as a negative control.

Acknowledgments This work was supported by the NIH Grant (1R21CA179286) to V. P. Torchilin. The authors thank Dr. W. Hartner for helpful comments on the preparation of the manuscript.

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Size Distribution by Intensity Intensity (Percent)

20 15 10 5 0 0.1

1

10 100 Size (d.nm)

1000

10000

Size Distribution by Intensity Intensity (Percent)

15 10 5 0 0.1

1

10

100

1000

10000

Size (d.nm)

Fig. 3 Size distribution of transferrin-modified micelles. On the top side of the image, size distribution of nonmodified micelles is presented (pNP-PEG3,400pNP+ DSPE-PEG1,000). On the bottom side, size distribution of transferrinmodified micelles is presented

References 1. Johnsen KB, Burkhart A, Melander F et al (2017) Targeting transferrin receptors at the blood-brain barrier improves the uptake of immunoliposomes and subsequent cargo transport into the brain parenchyma. Sci Rep 7 (1):10396 2. Jefferies WA, Brandon MR, Hunt SV et al (1984) Transferrin receptor on endothelium of brain capillaries. Nature 312:162–163 3. Cheng Y, Zak O, Aisen P et al (2004) Structure of the human transferrin receptor-transferrin complex. Cell 116(4):565–576

4. Muddineti OS, Kumari P, Ghosh B et al (2018) Transferrin-modified vitamin-E/lipid based polymeric micelles for improved tumor targeting and anticancer effect of curcumin. Pharm Res 35 (5):97 5. Lakkadwala S, Dos Santos Rodrigues B, Sun C et al (2019) Dual functionalized liposomes for efficient co-delivery of anti-cancer chemotherapeutics for the treatment of glioblastoma. J Control Release 307:247–260 6. Pinheiro RGR, Granja A, Loureiro JA et al (2020) Quercetin lipid nanoparticles

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functionalized with transferrin for Alzheimer’s disease. Eur J Pharm Sci 148:105314 7. Jhaveri A, Luther E, Torchilin V (2019) The effect of transferrin-targeted, resveratrol-loaded liposomes on neurosphere cultures of glioblastoma: implications for targeting tumourinitiating cell. J Drug Target 27(5–6):601–613 8. Jhaveri A, Deshpande P, Pattni B et al (2018) Transferrin-targeted, resveratrol-loaded

liposomes for the treatment of glioblastoma. J Control Release 277:89–101 9. Deshpande P, Jhaveri A, Pattni B et al (2018) Transferrin and octaarginine modified dualfunctional liposomes with improved cancer cell targeting and enhanced intracellular delivery for the treatment of ovarian cancer. Drug Deliv 25 (1):517–532

Chapter 6 Peptide-Pegylated Lipid Conjugation Via Copper-Catalyzed Alkyne-Azide 1,3-Dipolar Cycloaddition Waleed M. Hussein and Istvan Toth Abstract Targeted drug delivery is an important strategy in the treatment of many diseases. However, cancer cells are very difficult to target, making this a substantial obstacle in chemotherapy treatment. Bombesin fragment (BBN(6–14)) has been found to target gastrin-releasing peptide receptor (GRPR), which is overexpressed in many cancer cells. In this chapter, BBN peptide was used as a targeting moiety on the surface of polymeric-based nanoparticles to deliver its payload into prostate cancer PC-3 cell lines. Copper-catalyzed alkyne-azide 1,3-dipolar cycloaddition (CuAAC) click reaction was utilized to link the BBN peptide with an alkyne derivative of Pegylated lipid. Key words Bombesin, Targeting peptide, Polyethylene glycol linker, Alkyne-azide cycloaddition, Click chemistry

1

Introduction The mammalian bombesin (BBN) family of receptors is comprised of three subclasses: neuromedin B receptor; gastrin-releasing peptide receptor (GRPR); and orphan receptor, BBN receptor subtype-3. The ligand, 14-amino acid BBN peptide (PEQRLGNQ WAVGHLM), was initially isolated from the skin of an amphibian. Gastrin-releasing peptide (GRP) is the mammalian equivalent of BBN peptide: it has 27 amino acids, seven of which (WAVGHLM) mirror the C-terminus region of BBN [1]. These seven amino acids are accountable for both binding and internalization through GRPR [1]. Both BBN and GRP are mitogenic and account for many human carcinomas as autocrine growth factors [2]. GRPR is overexpressed in many cancers, including glioblastomas, small cell lung, gastric, pancreatic, prostate, breast, cervical, and colon cancers [3, 4]. Because of their strong receptor-targeting capacities, both BBN and GRP have been employed as targeting moieties in tumor-targeted gene delivery systems and in vivo tumor imaging

Waleed M. Hussein, Rachel J. Stephenson and Istvan Toth (eds.), Peptide Conjugation: Methods and Protocols, Methods in Molecular Biology, vol. 2355, https://doi.org/10.1007/978-1-0716-1617-8_6, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Synthesis of DOPE-PEG3400-alkyne (4)

[5]. Several studies have assessed a truncated BBN peptide BBN (6–14, NQWAVGHLM) as a targeting ligand for the delivery of siRNA and antisense oligonucleotides. Ming et al. showed that BBN(6–14) peptide successfully transmitted a target-specific delivery of splice-shifting oligonucleotides (SSO) into prostate cancer PC-3 cell lines [6]. And recently, our group demonstrated the crucial role of BBN(6–14) on the surface of nanoparticle formulations for expressing high levels of enhanced green fluorescence protein (eGFP), and eGFP knockdowns through delivery of DNA [7, 8] and siRNA [8, 9], respectively. This chapter describes the synthesis of DOPE-PEG3400-BBN (6) through the conjugation of pegylated alkyne derivative 4 with the azidoacetyl derivative, BBN(6–14) peptide (5), using coppercatalyzed alkyne-azide 1,3-dipolar cycloaddition (CuAAC) reaction. In this process, pegylated alkyne derivative 4 is synthesized first through a two-step, one-pot reaction (Fig. 1). In the presence of triethylamine (TEA), dioleoyl phosphatidylethanolamine (DOPE, 1) is treated with an excess amount of nitrophenylcarbonyl-PEG3400-nitrophenylcarbonyl (NPC-PEG3400-NPC, 2) to give the mono NPC derivative 3. The replacement of NPC groups with propargyl moieties is performed through the addition of an excess amount of propargyl amine and TEA to the reaction mixture to obtain a mixture of compound 4, along with the side product propargyl-PEG3400-propargyl. Thin-layer chromatography (TLC) using 30% methanol in chloroform is used to monitor the reaction progress, and the products are detected by Dragendorff’s stain. The tendency of product 4, rather than the dipropargyl analogue of 2, to form micelles allows for the isolation of 4 by size exclusion sepharose column chromatography (CL-4B). Pure product 4 (87%) was obtained as a light brown solid and confirmed by nuclear magnetic resonance (NMR).

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Fig. 2 Application of CuAAC reaction to synthesize DOPE-PEG3400-bombesin (6)

Fig. 3 MALDI-TOF mass spectrometry of DOPE-PEG3400-alkyne (4) (top) and DOPE-PEG3400-bombesin (6) (bottom)

The N-terminal azidoacetyl derivative of BBN(6–14) peptide (5, N3CH2CO-NQWAVGHLM-NH2) was obtained using Fmoc solid-phase peptide synthesis (Fmoc-SPPS) [10] (Fig. 1). Conjugation of DOPE-PEG3400-alkyne (4) with 5 provided DOPE-PEG3400-BBN (6) (Fig. 2). The product 6 (m/z ¼ ~ 5300) was confirmed by MALDITOF mass spectrometry (Fig. 3).

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Materials

2.1 Synthesis of DOPE-PEG3400Alkyne (4)

1. DOPE (1), 25 mg/mL stock in chloroform. 2. TEA. 3. NPC-PEG3400-NPC (2). 4. Anhydrous chloroform. 5. Nitrogen gas. 6. Propargylamine. 7. Sepharose CL-4B. 8. 30% methanol in chloroform for TLC. 9. Dragendorff’s stain.

2.2 Synthesis of DOPE-PEG3400Bombesin (6)

1. N-terminal azidoacetyl derivative of BBN(6–14) peptide (5, N3CH2CO-NQWAVGHLM-NH2). 2. DOPE-PEG3400-alkyne 4. 3. N,N-Dimethyl formamide (DMF). 4. Copper wire. 5. Aluminum foil. 6. Nitrogen balloon. 7. Stopwatch.

2.3 Self-Assembly of DOPE-PEG3400Bombesin (6)

1. Scintillation vial (20 mL). 2. Scintillation vial cap with hole. 3. Magnet (1 cm). 4. Endotoxin-free ultrapure water. 5. Syringe pump. 6. Magnetic stirrer. 7. Long bent needle (~15 cm). 8. Syringe (5 mL).

2.4 Purification of DOPE-PEG3400Bombesin (6)

1. Pierce snakeskin dialysis bag, MWCO 3 K (see Note 1). 2. Endotoxin-free ultrapure water. 3. 1 L beaker. 4. Two clamps. 5. Piece of sponge (~20 cm2). 6. Magnetic stirrer. 7. Magnet (~10 cm).

Peptide-Pegylated Lipid Conjugation Via Copper-Catalyzed Alkyne-Azide 1,3. . .

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Methods

3.1 Synthesis of DOPE-PEG3400Alkyne (4)

1. In a 25 mL round-bottom flask, dissolve NPC-PEG3400-NPC (2, 250 mg, 73.5 μmol, 4.9 equiv.) in anhydrous chloroform (3 mL) (see Note 2). 2. Add a mixture of DOPE (1, 11.4 mg, 456 μL, 25 mg/mL stock in chloroform, 15 μmol, 1 equiv.) and TEA (11.6 mg, 16 μL, 114 μmol, 7.6 equiv.) dropwise to the solution. 3. Stir the mixture for 14 h at room temperature under nitrogen atmosphere. 4. Add a mixture of propargylamine (47 μL, 40.4 mg, 735 μmol, 49 equiv.) and TEA (307 μL, 223 mg, 2.205 mmol, 147 equiv.) to the reaction mixture. 5. Stir the mixture for another 14 h at room temperature under nitrogen atmosphere. 6. Evaporate the solvent in vacuo and dry it in a freeze dryer overnight. 7. Take up the residue with water (1 mL), vortex, then purify the solution by size-exclusion sepharose CL-4B (see Note 3). 8. Detect the fractions by TLC (using 30% methanol in chloroform) and stain with Dragendorff’s reagent (see Note 4). 9. Combine the fractions, including DOPE-PEG3400-alkyne (4), together and freeze-dry (see Note 5).

3.2 Synthesis of DOPE-PEG3400Bombesin (6)

1. In a dry 2 mL round-bottom flask, dissolve a mixture of BBN azide 5 (3.1 mg, 2.7 μmol, 2.5 equiv.) and DOPE-PEG3400alkyne 4 (4.6 mg, 1.1 μmol, 1 equiv.) in DMF (1 mL). 2. Add copper wire (60 mg) to the reaction mixture (see Note 6). 3. Partially remove the air from the reaction mixture by bubbling a nitrogen balloon for 30 s (see Note 7). 4. Cover the reaction mixture with aluminum foil to protect it from light, then stir at 50  C under nitrogen atmosphere. 5. Stop the reaction after 2 h (see Note 8).

3.3 Self-Assembly of DOPE-PEG3400Bombesin (6)

1. Load the DMF filtrate into a 5 mL syringe. 2. Attach a long bent needle to the syringe (see Note 9). 3. Adjust the rate of addition to 0.5 mL/h using a syringe pump (see Note 10). 4. Add 4 mL of endotoxin-free water into a 20 mL scintillation vial charged with a 1 cm magnet; cover using a scintillation cap with hole (see Note 11).

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5. Insert the bent needle through the hole in the cap and adjust the needle so that its tip touches the surface of the water close to the middle of the vial. 6. Add the DMF solution to the endotoxin-free water at a rate of 0.5 mL/h while stirring the mixture at 250 rpm. 3.4 Purification of DOPE-PEG3400Bombesin (6)

1. Transfer the above mixture to a 6 cm dialysis bag (see Note 12). 2. Dialyze the solution against 1 L endotoxin-free water (pH 6.8) to clear away copper salts and unreacted peptides (see Note 13). 3. Freeze-dry the purified solution and measure the mass of the resulting dry solid. 4. Detect the freeze-dried product by MALDI-TOF mass spectrometry (m/z ¼ ~5400) (Fig. 3) (see Note 14).

4

Notes 1. The molecular weight cutoff (MWCO) of the membrane is the separation feature most generally described by the pore size range of a dialysis membrane. Therefore, dialysis bag selection is based on the molecular weight of the peptide in focus. The MWCO of the dialysis bag must be greater than the peptide’s molecular weight and needs to be adjusted when using peptides of different molecular weights. 2. The 25 mL round-bottom flask and stir bar must be washed with anhydrous chloroform, and the 250 mg of PNP-PEGPNP must be weighed in a clean (chloroform washed) vial. 3. The column should be 50 cm high  2 cm wide. The sample was eluted with degassed MilliQ water and 2 mL/fraction was collected. 4. PEG-containing compounds were stained orange using Dragendorff’s reagent, whereas the phosphate group-containing compound from DOPE was stained blue using molybdenum blue reagent. 5. The product DOPE-PEG3400-alkyne (4) was obtained as a light brown solid (56 mg, 87%). 1H NMR (500 MHz, CDCl3) δ 6.70 (s, 1H), 5.37 (s, 1H), 5.35–5.26 (m, 3H), 5.18 (s, 1H), 4.39–4.30 (m, 1H), 4.26–4.16 (m, 2H), 4.16–4.10 (m, 1H), 3.96–3.91 (m, 2H), 3.77–3.73 (m, 1H), 3.73–3.53 (m, 152H), 3.50–3.44 (m, 1H), 3.34 (s, 1H), 2.31–2.23 (m, 6H), 2.22 (t, J ¼ 2.5 Hz, 1H), 1.97 (q, J ¼ 6.5 Hz, 6H), 1.61–1.51 (m, 3H), 1.33–1.18 (m, 40H), 0.85 (t, J ¼ 7.0 Hz, 6H). 13C NMR (125 MHz, CDCl3) δ 173.4, 130.0, 129.7, 79.8, 71.4, 70.8, 70.5, 69.9, 69.4, 64.3, 34.2, 34.0, 31.9, 30.7, 29.72, 29.70, 29.5, 29.3, 29.24,

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29.21, 29.13, 29.10, 29.08, 27.18, 27.16, 24.87, 24.82, 22.6, 14.1. 6. Prior to use, the copper wires must be bright and pink in color. If they are not, wash them in a small sintered glass funnel with concentrated sulfuric acid (three times), water (six times), and methanol (six times), then vacuum-dry. 7. Some air is required to oxidize copper metal into copper I (catalysis of the CuAAC reaction). The complete removal of air may cause a major delay or stop the reaction, but too much oxygen could induce the undesirable formation of large amounts of copper II. 8. The reaction mixture changed from colorless to green after 2 h, indicating the occurrence of the reaction. 9. It is necessary to wash the needle with chloroform three times, then dry prior to use. 10. The rate of addition of DMF to water needs to be adjusted to complete self-assembly in 3–4 h. The needle should not touch the wall of vial. 11. It is necessary to wash the scintillation vial and magnet three times with endotoxin-free water before use. 12. Prior to use, the dialysis bag needs to be soaked and rinsed using endotoxin-free water. The bag should then be folded from the bottom twice. After loading the self-assembled solution into the bag using a 1 mL tip, fold the top of the bag twice, clamp it from both ends, and attach it to a piece of sponge (to allow the dialysis bag to submerge in water but not sink to the bottom of the flask). 13. Change the dialysis water three times per day; continue dialysis for 3 days. 14. Pure DOPE-PEG3400-BBN (6) was obtained as an amorphous powder (4.8 mg, 82%). References 1. Mansi R, Fleischmann A, Macke HR, Reubi JC (2013) Targeting GRPR in urological cancers -from basic research to clinical application. Nat Rev Urol 10:235–244 2. Ma LX, Yu P, Veerendra B, Rold TL, Retzloff L, Prasanphanich A, Sieckman G, Hoffman TJ, Volkert WA, Smith CJ (2007) In vitro and in vivo evaluation of Alexa Fluor 680-bombesin[7-14]NH2 peptide conjugate, a high-affinity fluorescent probe with high selectivity for the gastrin-releasing peptide receptor. Mol Imaging 6:171–180

3. Povsic TJ, Vavalle JP, Alexander JH, Aberle LH, Zelenkofske SL, Becker RC, Buller CE, Cohen MG, Cornel JH, Kasprzak JD, Montalescot G, Fail PS, Sarembock IJ, Mehran R (2014) Use of the REG1 anticoagulation system in patients with acute coronary syndromes undergoing percutaneous coronary intervention: results from the phase II RADAR-PCI study. EuroIntervention 10:431–438 4. Yang H, Cai HW, Wan L, Liu S, Li SF, Cheng JQ, Lu XF (2013) Bombesin analoguemediated delivery preferentially enhances the

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cytotoxicity of a mitochondria-disrupting peptide in tumor cells. PLoS One 8:e57358 5. Abd-Elgaliel WR, Gallazzi F, Garrison JC, Rold TL, Sieckman GL, Figueroa SD, Hoffman TJ, Lever SZ (2008) Design, synthesis, and biological evaluation of an antagonistbombesin analogue as targeting vector. Bioconjug Chem 19:2040–2048 6. Ming X, Alam MR, Fisher M, Yan YJ, Chen XY, Juliano RL (2010) Intracellular delivery of an antisense oligonucleotide via endocytosis of a G protein-coupled receptor. Nucleic Acids Res 38:6567–6576 7. Begum AA, Wan Y, Toth I, Moyle PM (2018) Bombesin/oligoarginine fusion peptides for gastrin releasing peptide receptor (GRPR)

targeted gene delivery. Bioorg Med Chem 26:516–526 8. Begum AA, Toth I, Moyle PM (2019) Gastrinreleasing peptide receptor-targeted hybrid peptide/phospholipid pDNA/siRNA delivery systems. Nanomedicine 14:1153–1171 9. Hussein WM, Cheong YS, Liu C, Liu G, Begum AA, Attallah MA, Moyle PM, Torchilin VP, Smith R, Toth I (2019) Peptide-based targeted polymeric nanoparticles for siRNA delivery. Nanotechnology 30:415604 10. Begum AA, Moyle PM, Toth I (2016) Investigation of bombesin peptide as a targeting ligand for the gastrin releasing peptide (GRP) receptor. Bioorg Med Chem 24:5834–5841

Chapter 7 Vitamin B12 – Peptide Nucleic Acid Conjugates Aleksandra J. Wierzba, Monika Wojciechowska, Joanna Trylska, and Dorota Gryko Abstract Vitamin B12 (cobalamin, Cbl) is an essential nutrient for all mammals and some bacteria. From a chemical point of view, it is a highly functionalized molecule, which enables conjugation at multiple positions and attachment of various cargoes. Both mammalian and bacterial cells have developed a specific transport pathway for the uptake of vitamin B12, and as a consequence, cobalamin is an attractive candidate for the delivery of biologically relevant molecules into cells. Indeed, hybrid molecules containing vitamin B12 in their structure have found various applications in medicinal chemistry, diagnostics, and biological sciences. Herein, we describe synthetic strategies toward the synthesis of vitamin B12 conjugates with peptide nucleic acid (PNA) oligomers. Such short-modified oligonucleotides targeted at bacterial DNA or RNA can act as antibacterial agents if efficiently taken up by bacterial cells. The uptake of such oligonucleotides is hindered by the bacterial cell envelope, but vitamin B12 was found to efficiently deliver antisense PNA into Escherichia coli and Salmonella Typhimurium cells. This paves the way to the use of vitamin B12-PNA conjugates in antibacterial and diagnostic applications. Vitamin B12-PNA conjugates can be prepared via copper(I)-catalyzed azide-alkyne cycloaddition (CuAAC) that gives access to covalently linked hybrids or via connecting both building blocks by reduction-sensitive disulfide bridge. Both approaches require prior modification of vitamin B12 by incorporation of the azide moiety or via transformation of the native functional group into a moiety reactive toward thiols. Conjugation of vitamin B12 with PNA-tagged substrates efficiently furnishes designed conjugates. Key words Vitamin B12, Cobalamin, Peptide nucleic acids, PNA, Antisense oligonucleotides, Drug delivery, Antibacterial agents, Conjugates

1

Introduction Vitamin B12 (cobalamin, Cbl) is a natural organometallic compound and an essential nutrient cofactor for all mammals and bacteria, but its biosynthesis is restricted to some microorganisms [1]. The human body is not able to synthesize this vitamin; therefore, it must be included in a diet. As an exogenous compound, cobalamin reaches mammalian cells via a system of transport

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Fig. 1 The structure of cyanocobalamin (on the left) with positions available for conjugation (marked with blue circles) and selected examples of vitamin B12 conjugates (on the right; a [4], b [5], c [6])

proteins, and this fact makes it an attractive candidate for the delivery of cargoes into cells [2]. Indeed, vitamin B12 has been recognized as a delivery vehicle for various biologically relevant molecules (drugs, fluorescent dyes, radionuclides, Fig. 1) [3]. The synthesis of vitamin B12 conjugates requires suitably designed building blocks. Cobalamin possesses a number of functional groups susceptible to chemical modifications; however, the position of conjugation must be selected in such a way that the process of cobalamin recognition is not disturbed. Modifications at the e-propionamide chain, β-axial position, and R5’-OH group were found to have the least adverse effect on the recognition process in mammalian cells [5], while for bacteria, R5’ position was determined to be the most convenient for these purposes (Fig. 1) [7]. A variety of microorganisms are also capable to take up vitamin B12. Gram-negative bacteria actively transport vitamin B12 using a cascade of membrane and periplasmic proteins [8–10]. The ability of vitamin B12 to transport oligonucleotide analogues [11, 12] into bacterial cells represents a new approach to antibacterial agents [13]. Due to the widespread emergence of bacterial resistance to commonly overused antibiotics, it may be vital for the treatment of bacterial infections [14, 15]. Therefore, the use of natural vitamin B12 as a delivery vehicle seems an excellent choice for the administration of oligonucleotides, which cannot penetrate the bacterial cell envelope. We thus designed and synthesized vitamin B12-peptide nucleic acid (PNA) conjugates [13]. PNA is a DNA analogue possessing nucleobases attached to the repeating N-(2-aminoethyl) glycine units linked by amide bonds (Fig. 2) [16]. The neutral PNA backbone increases its affinity toward natural nucleic acids because

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Fig. 2 Schematic representation of vitamin B12-PNA conjugate mode of action in bacterial cells (left) and comparison of RNA and PNA structure (right)

of no interstrand electrostatic repulsion of the phosphate groups. Specifically designed PNA oligonucleotides suppress proper expression of bacterial genes by complementary binding to essential sequences of bacterial DNA or RNA (Fig. 2) [17]. Cobalamin transports antisense PNA into Escherichia coli cells more efficiently than the most widely used cell-penetrating peptide (KFF)3K [13]. In this chapter, we describe the synthesis of vitamin B12-PNA conjugates. In the presented examples the PNA strand is appended at the R5’ position of the ribose moiety in the structure of cobalamin as it was found not to interfere with the recognition process in bacterial cells [7]. The synthesis involves preparation of specifically designed cyanocobalamin derivatives and subsequent attachment of a suitably tailored PNA oligomer to the vitamin either directly or via a linker. Stable vitamin B12-PNA conjugates are prepared via copper(I)-catalyzed azide-alkyne cycloaddition that furnishes covalently linked hybrids. For this purpose, the azide moiety is introduced to the structure of cobalamin either via substitution of cobalamin mesylate 2 or via carbamate methodology (Fig. 3a, b) [18, 19]. The latter approach involves activation of the hydroxyl group with 1,10 -carbonyl-di-(1,2,4-triazole) (CDT) and subsequent reaction with an aminoazide of choice. For the preparation of cleavable conjugates, the hydroxyl group at R5’ position is selectively transformed into pyridyl disulfide—the moiety highly reactive toward thiols (Fig. 3c) [20]. PNA oligomers are tagged with the respective functional groups (alkyne moiety for CuAAC or sulfhydryl group for disulfide bond formation, Fig. 4) during the solid-phase synthesis [13]. The final conjugation of vitamin B12 with modified PNA strand is performed in solution, and designed conjugates are purified via reversed-phase high-performance liquid chromatography

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Fig. 3 Synthesis of vitamin B12 derivatives with azide moiety (a, b) and pyridyl disulfide group (c)

Fig. 4 Conjugation of vitamin B12 with peptide nucleic acids via CuAAC (a, b) and disulfide bond formation (c)

(RP-HPLC) techniques (Fig. 4). Other positions in the structure of cobalamin were also engaged for the preparation of conjugates with PNA, but they are not discussed in this chapter [7].

2

Materials The term “vitamin B12” refers to the vitamin B12 form with β-cyanide ligand (cyanocobalamin, (CN)Cbl). All reagents and solvents should be purchased at reagent grade and ultrapure Milli-Q grade H2O used unless specified otherwise.

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Anhydrous solvents (96% (detected by evaporative light scattering detector) (see Note 14). Yield: (3.2 mg, 87%). ESI-MS: m/z 1466.2 (calc 1466.1) [M + 3H]3+; 1100.0 (calc 1099.8) [M + 4H]4+; 880.2 (calc 880.0) [M + 5H]5+; MW 4395.1.

Double Conjugation Using Mercapto-Acryloyl and Alkyne-Azide Reactions. . .

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Notes 1. Store solution at 4  C for a maximum of 1 week after preparation. Do not use the solution if its color has changed to yellow. 2. Azidoacetic acid is sensitive to light and can explode if heated to a high temperature such as 50  C. 3. Each washing must be performed carefully to ensure that all reagents and unbound byproducts are removed from the resin. Improper washing could result in the formation of side products during synthesis. 4. Fmoc deprotection of Thr, Val, and Ile were performed with 2% of 1,8-diazabicycloundec-7-ene (DBU) in DMF (twice, at 5 and 10 min) instead of 20% piperidine in DMF; Fmoc deprotection for Asp was performed with 0.1 M HOBt in 20% piperidine/DMF. 5. Aluminum foil is used to protect the azidoacetic acid from light. 6. Methanol wash is used for shrinking resin. Extensive washing may result in the loss of resin if it becomes small enough to pass through the filter. 7. After adding the two peptides, ensure that the pH of the reaction mixture is adjusted to 7.3 by testing 0.3 μL of the solution using pH indicator strips. 8. After 48 h, analytical HPLC showed the complete consumption of acryloyl E643–57 (2). 9. Method A: 0.1% TFA/H2O as solvent A and 90% MeCN/0.1% TFA/H2O as solvent B on either a Vydac analytical C4 column (214TP54; 5 μm, 4.6 mm  250 mm) or a Vydac analytical C18 column (218TP54; 5 μm, 4.6 mm  250 mm). 10. Ensure that the used copper wires are shiny and pink in color. If not, wash the copper wires with concentrated sulfuric acid (three times), water (six times), and methanol (six times) in a small sintered glass funnel, then dry under vacuum. 11. The complete removal of air can cause a substantial delay, or cease the reaction, as a small amount of oxygen is needed to oxidize copper metal into copper I; this process is essential for catalyzing the CuAAC reaction. In contrast, too much oxygen can trigger undesirable formation of large amounts of copper II. 12. After 4 h, the reaction mixture changed from colorless to green, indicating the occurrence of the reaction. Analytical HPLC and ESI-MS also showed the complete consumption of peptide 3 after 4 h.

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13. Before injecting the DMF solution into the HPLC column, the solution must first be filtered through a 0.45 μm filter, and the filter should be washed using 0.5 mL DMF. 14. To provide proof of purity, both UV and ELSD detectors were used to establish the HPLC signal of lipopeptide 5. References 1. Liu TY, Hussein WM, Toth I, Skwarczynski M (2012) Advances in peptide-based human papillomavirus therapeutic vaccines. Curr Top Med Chem 12:1581–1592 2. Purcell AW, McCluskey J, Rossjohn J (2007) More than one reason to rethink the use of peptides in vaccine design. Nat Rev Drug Discov 6:404–414 3. Sarkar AK, Tortolero-Luna G, Nehete PN, Arlinghaus RB, Mitchell MF, Sastry KJ (1995) Studies on in vivo induction of cytotoxic T lymphocyte responses by synthetic peptides from E6 and E7 oncoproteins of human papillomavirus type 16. Viral Immunol 8:165–174 4. Manuri PR, Nehete B, Nehete PN, Reisenauer R, Wardell S, Courtney AN, Gambhira R, Lomada D, Chopra AK, Sastry KJ (2007) Intranasal immunization with synthetic peptides corresponding to the E6 and E7 oncoproteins of human papillomavirus type 16 induces systemic and mucosal cellular immune responses and tumor protection. Vaccine 25:3302–3310 5. Liu T-Y, Hussein WM, Jia Z, Ziora ZM, McMillan NAJ, Monteiro MJ, Toth I, Skwarczynski M (2013) Self-adjuvanting polymerpeptide conjugates as therapeutic vaccine candidates against cervical cancer. Biomacromolecules 14:2798–2806 6. Liu TY, Hussein WM, Giddam AK, Jia Z, Reiman JM, Zaman M, McMillan NA, Good MF, Monteiro MJ, Toth I et al (2015) Polyacrylate-based delivery system for selfadjuvanting anticancer peptide vaccine. J Med Chem 58:888–896 7. Liu TY, Giddam AK, Hussein WM, Jia Z, McMillan NA, Monteiro MJ, Toth I, Skwarczynski M (2015) Self-adjuvanting therapeutic peptide-based vaccine induce CD8+ cytotoxic T lymphocyte responses in a murine human papillomavirus tumor model. Curr Drug Deliv 12:3–8 8. Raibaut L, Ollivier N, Melnyk O (2012) Sequential native peptide ligation strategies for total chemical protein synthesis. Chem Soc Rev 41:7001–7015

9. Kent SBH (2009) Total chemical synthesis of proteins. Chem Soc Rev 38:338–351 10. Aimoto S (1999) Polypeptide synthesis by the thioester method. Biopolymers 51:247–265 11. Dawson PE, Muir TW, Clark-Lewis I, Kent SBH (1994) Synthesis of proteins by native chemical ligation. Science 266:776–779 12. Dawson PE, Kent SBH (2000) Synthesis of native proteins by chemical ligation. Annu Rev Biochem 69:923–960 13. Biernat M, Stefanowicz P, Zimecki M, Szewczuk Z (2006) Amino-terminal dimerization of peptides on the solid support. Synthesis and biological activity of the immunosuppressive HLA-DR fragments linked by poly(ethylene glycol)s. Bioconjug Chem 17:1116–1124 14. Johnson DL, Farrell FX, Barbone FP, McMahon FJ, Tullai J, Kroon D, Freedy J, Zivin RA, Mulcahy LS, Jolliffe LK (1997) Aminoterminal dimerization of an erythropoietin mimetic peptide results in increased erythropoietic activity. Chem Biol 4:939–950 15. Bracci L, Falciani C, Lelli B, Lozzi L, Runci Y, Pini A, De Montis MG, Tagliamonte A, Neri P (2003) Synthetic peptides in the form of dendrimers become resistant to protease activity. J Biol Chem 278:46590–46595 16. Tang X-D, Wang G-Z, Guo J, Lue M-H, Li C, Li N, Chao Y-L, Li C-Z, Wu Y-Y, Hu C-J et al (2012) Multiple antigenic peptides based on H-2Kb-restricted CTL epitopes from murine Heparanase induce a potent antitumor immune response in vivo. Mol Cancer Ther 11:1183–1192 17. Hussein WM, Liu TY, Maruthayanar P, Mukaida S, Moyle PM, Wells JW, Toth I, Skwarczynski M (2016) Double conjugation strategy to incorporate lipid adjuvants into multiantigenic vaccines. Chem Sci 7:2308–2321 18. Skwarczynski M, Zaman M, Urbani CN, Lin IC, Jia Z, Batzloff MR, Good MF, Monteiro MJ, Toth I (2010) Polyacrylate dendrimer nanoparticles: a self-adjuvanting vaccine delivery system. Angew Chem Int Ed 49:5742–5745

Chapter 14 Chemical Protein Synthesis by Chemoselective α-Ketoacid–Hydroxylamine (KAHA) Ligations with 5-Oxaproline Jakob Farnung, Haewon Song, and Jeffrey W. Bode Abstract Chemical protein synthesis enables the precise construction of proteins by employing solid-phase peptide synthesis and chemoselective ligations. One such chemoselective reaction suitable for protein synthesis is the α-Ketoacid-Hydroxylamine (KAHA) ligation. Fully unprotected peptides are ligated by a selective reaction between α-ketoacids and hydroxylamines to give native amide bonds. Herein, we describe the chemical synthesis of ubiquitin by a two-segment approach using the 5-oxaproline hydroxylamine. Key words KAHA ligation, 5-Oxaproline, Hydroxylamine, α-Ketoacid, Ubiquitin, SPPS

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Introduction Chemical protein synthesis has facilitated access to proteins carrying a variety of natural and unnatural modifications at discrete locations not accessible with recombinant techniques [1, 2]. Assembly of full-length proteins depends on the interplay of solid-phase peptide synthesis (SPPS) and peptide ligation techniques. SPPS, in general, affords peptides of up to about 50 amino acids, depending on the sequence. As the majority of proteins are significantly larger, techniques have been developed to ligate multiple peptide segments together to afford full-length proteins. The field of peptide ligation was revolutionized by Kent et al. with the development of native chemical ligation (NCL) [3]. In NCL, a peptide segment with a C-terminal thioester reacts selectively with a peptide segment containing an N-terminal cysteine [4]. At the outset, the requirement for cysteine at the ligation site restricted the number of potential ligation sites. This limitation was addressed by the development of desulfurization strategies, leading to the widespread adoption of NCL [5]. Alternative ligation approaches such as

Waleed M. Hussein, Rachel J. Stephenson and Istvan Toth (eds.), Peptide Conjugation: Methods and Protocols, Methods in Molecular Biology, vol. 2355, https://doi.org/10.1007/978-1-0716-1617-8_14, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

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Ser/Thr ligation and KAHA ligation were developed to increase the number of potential ligation sites and overcome the limitations of NCL [6]. In KAHA ligation, C-terminal α-ketoacids react chemoselectively with N-terminal hydroxylamines of a different segment to provide a native amide bond. Commonly, (S)-5-oxaproline (Opr) is used as the nucleophilic reaction partner, resulting in a non-native homoserine residue at the ligation site [7]. A variety of amino acids can be mutated to homoserine with negligible effects on protein folding or activity. Oxaproline ligations initially provide a depsipeptide as the product which, under basic aqueous conditions, readily undergoes O-to-N acyl shift to the amide [8]. α-Ketoacids of a variety of amino acids have been synthesized and readily attached to Fmoc-SPPS compatible Rink Amide resin via a simple acid-labile linker using standard coupling conditions (Fig. 1) [9]. Ligations are performed in acidic organic-aqueous media, which are well suited for solubilizing even hydrophobic proteins that commonly cause severe difficulties in assembly and ligation. KAHA ligation is also amenable to multi-segment ligation by utilizing orthogonally protected oxaprolines and/or α-ketoacid derivatives [10, 11]. The first step in designing the chemical synthesis of a protein by KAHA ligation is the selection of a suitable ligation site, which divides the protein into segments of acceptable lengths. The selection of a suitable ligation site is dictated by the available α-ketoacids, and by which amino acids can functionally be substituted by homoserine [12]. Ideally, such ligation sites should be located in regions of the protein where they are unlikely to alter the folding or function. A variety of amino acid-derived α-ketoacids are readily available; however, the most commonly used α-ketoacids are derived from leucine, phenylalanine, tyrosine, and valine. α-Ketoacids can also be accessed via a cyanosulfurylide-linker, which is converted to the α-ketoacid after resin cleavage and purification by treatment with aqueous acidic Oxone [13, 14]. 5Oxaproline results in the noncanonical amino acid homoserine at the ligation site and should, therefore, replace amino acids with functional or structural similarity to homoserine. These amino acids include, but are not limited to, serine, threonine, aspartic acid, asparagine, and methionine. Recent advancements enable access to native amino acids (Ser, Thr, Asp) or reactive handles (aspartyl aldehyde) [15–18]. Herein, we report the two-segment synthesis of ubiquitin by KAHA ligation [19, 20]. Ubiquitin segments are prepared by standard Fmoc-SPPS incorporating phenylalanine-ketoacid and 5-oxaproline. This approach can be adapted for the synthesis of other proteins by two-segment ligations or—with the assistant of orthogonally protected α-ketoacids and hydroxylamines—for longer proteins by multi-segment assembly.

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Fig. 1 Loading of α-ketoacid monomer on resin for Fmoc-SPPS [9]

2

Materials 1. Automated peptide synthesizer. 2. Fmoc-Rink Amide resin. 3. 2-Chlorotrityl chloride resin. 4. UV spectrometer. 5. Standard Fmoc-protected amino acids, including norleucine (Nle). 6. Boc-protected (S)-5-oxaproline (Opr). Prepare this monomer using the methods described in reference [21]. This monomer should be stored at 4
C.

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7. Acid-labile phenylalanine α-ketoacid. Prepare this monomer using the methods described in reference [9]. This monomer should be stored at 4
C. 8. Anhydrous N,N-dimethylformamide (DMF). 9. Piperidine. 10. 2-(6-Chloro-1-H-benzotriazole-1-yl)-1,1,3,3-tetramethylaminium hexa-fluorophosphate (HCTU). 11. 1-[Bis(dimethylamino)methylene]-1H-1,2,3-triazolo[4,5-b] pyridinium 3-oxid hexafluorophosphate (HATU). 12. N,N-Diisopropylethylamine (DIPEA). 13. N-Methyl-2-pyrrolidone (NMP). 14. Acetic anhydride (Ac2O). 15. N-Methylmorpholine (NMM). 16. Methanol. 17. 1,8-Diazabicyclo(5.4.0)undec-7-ene (DBU). 18. Resin cleavage cocktail: Trifluoroacetic acid (TFA)/H2O/2,20 -(Ethylene-dioxy)diethanethiol (DODT) ¼ 95:2.5:2.5 (v/v/ v). Prepare directly before use. 19. Rotary evaporator. 20. Diethyl ether (Et2O). 21. Dichloromethane (CH2Cl2). 22. Centrifuge. 23. Centrifuge tubes. 24. CH3CN/H2O (1:1) containing 0.1% (v/v) TFA. 25. Collection tubes. 26. Analytical HPLC (fitted with reverse-phase C18 column, 4.6  250 mm; pore size: 100 Å; particle size: 5 μm), monitoring at 220 nm, 254 nm, and 301 nm. 27. Preparative HPLC (fitted with reverse-phase C18 column, 20  250 mm; pore size: 100 Å; particle size: 5 μm), monitoring at 220 nm, 254 nm, and 301 nm. 28. Eluents: Deionized H2O containing 0.1% (v/v) TFA (eluent A) and CH3CN containing 0.1% (v/v) TFA (eluent B). 29. MALDI or equivalent mass spectrometer. 30. Lyophilizer. 31. Liquid N2. 32. 80% aqueous DMSO (v/v) containing 0.1 M oxalic acid. Dissolve oxalic acid (90 mg) in 8.0 mL of H2O, add 2.0 mL of DMSO. 33. Glass vial and cap.

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34. Constant temperature shaker. 35. Bench-top vortexer. 36. Centrifuge. 37. Rearrangement buffer: 0.2 M Na2CO3/NaHCO3, pH 9.5. Dissolve 212 mg of Na2CO3 and 168 mg of NaHCO3 in 10 mL of deionized water and adjust the pH to 9.5 with 1.0 M HCl and 1.0 M NaOH. This solution can be stored for up to a week at room temperature. 38. Folding buffer: 20 mM NaOAc, 50 mM Gn.HCl adjusted to pH 5.0. Dissolve 16.4 mg of NaOAc and 47.8 mg of Gn.HCl in 10 mL of deionized water and adjust the pH to 5.0 with 1.0 M HCl. This solution can be stored for up to a week at room temperature. 39. Centrifugal filters, 3 k molecular weight cutoff. 40. Deionized water.

3

Methods

3.1 Preparation of C-Terminal α-Ketoacid Resin

1. Prepare the acid-labile-protected phenylalanine α-ketoacid monomer for segment 1 as described in reference [9]. 2. Weigh out Fmoc-Rink Amide resin (2 g) into a SPPS vessel (see Note 1). 3. Wash the Fmoc-Rink Amide resin with CH2Cl2 (2  10 mL) and DMF (2  10 mL). 4. Swell the resin using DMF (10 mL) for 20 min then remove the solvent. 5. Remove Fmoc protecting group by agitating with 20% piperidine in DMF (v/v, 10 mL) for 20 mins at room temperature (see Note 2). 6. Remove remaining solution and wash the resin with CH2Cl2 (3  10 mL) and DMF (3  10 mL). 7. Dissolve the protected phenylalanine α-ketoacid monomer (364 mg, 1.00 equiv. for a target loading of 0.3 mmol/g) and HCTU (243 mg, 0.98 equiv) in DMF (5 mL). Upon complete dissolution, add NMM (132 μL, 2.00 equiv). Allow preactivation to occur for 30 s (see Notes 3 and 4). 8. Add preactivated reaction mixture from step 7 to resin. 9. Seal the SPPS vessel and agitate gently overnight (see Note 2). 10. Remove remaining solution and wash the resin with CH2Cl2 (3  10 mL) and DMF (3  10 mL). 11. Dry the resin and measure the loading of the resin using standard methods (see Notes 5 and 6) [22].

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12. Swell the resin using DMF (10 mL) for 20 min then remove the solvent. 13. Mix 20% Ac2O in DMF (v/v) and 2 M DIPEA/NMP in a 4:1 ratio for a total volume of 10 mL. 14. Add capping solution from step 13 to pre-swelled resin. 15. Agitate resin gently for 10 min. 16. Remove remaining solution and wash the resin with CH2Cl2 (3  10 mL) and DMF (3  10 mL). 17. Dry resin using a N2-stream and measure loading (see Notes 5 and 6). 18. Store resin under N2 gas or use immediately for Fmoc-SPPS on the automated peptide synthesizer. The resin can be stored at 4
C for several weeks. 3.2 Preparation of C-Terminal Carboxylic Acid Resin

1. Weigh out 2-chlorotrityl chloride resin (2 g) into the SPPS vessel. 2. Wash resin with CH2Cl2 (2  10 mL) and DMF (2  10 mL). 3. Swell resin in DMF (10 mL) for 20 min, then remove the solvent. 4. Dissolve Fmoc-glycine (178 mg, 1.00 equiv. for a target loading of 0.3 mmol/g) in DMF (5 mL). Upon complete dissolution, add NMM (132 μL, 2.00 equiv). 5. Add preactivated glycine solution from step 4 to pre-swelled resin. 6. Agitate the resin gently for 4 h (see Notes 2 and 7). 7. Remove remaining solution and wash resin with CH2Cl2 (3  10 mL) and DMF (3  10 mL). 8. Mix DMF, methanol, and NMM in a 17:1:2 ratio (v/v/v) for a total volume of 10 mL. 9. Add capping mixture from step 8 to the resin. 10. Agitate resin gently for 5 min. 11. Remove remaining solution and wash resin with CH2Cl2 (3  10 mL) and DMF (3  10 mL). 12. Dry resin using a N2-stream and measure loading (see Notes 5 and 6). 13. Store resin under N2 gas or use immediately for Fmoc-SPPS on the automated peptide synthesizer. The resin can be stored at 4
C for several weeks.

3.3 Preparation of α-Ketoacid Segment (1)

1. Weigh out the phenylalanine α-ketoacid-forming resin prepared above (357 mg, loading of 0.28 mmol/g, scale 0.1 mmol) and transfer resin to a reaction vessel of an automated peptide synthesizer.

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157 O

H 2N

OH

QIFVKTLTG10 KTITLEVEPS20 DTIENVKAKI30 QDKEGIPPDQ40 QRLIF O

segment 1 (45 aa) Me

O

H N

O

O

GKQL50 EDGRTLSDYN60 IQKESTLHLV70 LRLRGG

OH

segment 2 (31 aa)

Fig. 2 Two segments synthesized by Fmoc-SPPS

2. Perform Fmoc-SPPS for sequence of ketoacid segment (Fig. 2). Use standard Fmoc-SPPS conditions (see Note 8) to synthesize the sequence until position 1. Met1 was mutated to Nle to avoid oxidation. 3. Remove final Fmoc protecting group using standard Fmoc deprotection conditions. Wash the resin with DMF (3  2 mL). 4. Perform final microcleavage to assess retention time and purity of segment (see Note 9). 5. Dry resin containing the α-ketoacid segment using a N2-stream. 6. Transfer dry resin into a 40 mL glass vial and add the cleavage cocktail (TFA/H2O/DODT, 10 mL per 500 mg of resin). Gently agitate at room temperature for 2 h (see Note 10). 7. Filter the resin and evaporate TFA on a rotary evaporator at 40
C (see Note 11). 8. Add 40 mL of cooled (0
C) diethyl ether to the crude material and centrifuge using a centrifuge tube at 3000  g for 3 min at room temperature (see Note 12). 9. Decant the diethyl ether gently. 10. Repeat steps 8 and 9 two more times to remove excess impurities. 11. Dry the crude peptide gently using a N2 stream. The crude peptide can be stored under N2 atmosphere at 4
C for several weeks if needed. 12. Analyze and purify the peptides using reverse-phase HPLC and mass spectrometry (see Notes 13 and 14). Collect and lyophilize the purified fractions.

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3.4 Preparation of Carboxylic Acid Segment (2) with Opr

1. Weigh out glycine resin prepared above (400 mg, loading of 0.25 mmol/g, scale 0.1 mmol) and transfer resin to a reaction vessel of an automated peptide synthesizer. 2. Perform Fmoc-SPPS for sequence of carboxylic acid segment (Fig. 2). Use standard Fmoc-SPPS conditions (see Note 8) to synthesize the sequence until position G47. 3. Remove Fmoc protecting group using standard Fmoc deprotecting conditions. Wash the resin with DMF (3  2 mL). 4. Weigh out Boc-Opr (43.4 mg, 2.00 equiv. to the initial resin loading) and HATU (74.9 mg, 1.98 equiv.) in a glass vial and dissolve in DMF (2 mL). Add NMM (37.2 μL, 4.00 equiv.) to the mixture. Allow preactivation to occur for 30 s. 5. Add the preactivation solution from step 4 to the resin and agitate the resin gently for 2 h (see Notes 2). Wash the resin with DMF (3  2 mL). 6. Perform final microcleavage to assess retention time and purity of segment (see Note 9). 7. Dry resin containing carboxylic acid segment using a N2-stream (see Note 5). 8. Transfer dry resin into a 40 mL glass vial and add the cleavage cocktail (TFA/H2O/DODT, 10 mL per 500 mg of resin). Gently agitate at room temperature for 2 h. 9. Filter the resin and evaporate TFA on a rotary evaporator at 40
C (see Note 11). 10. Add 40 mL of cooled (0
C) diethyl ether to the crude material and centrifuge using a centrifuge tube at 3000  g for 3 min at room temperature (see Note 12). 11. Decant the diethyl ether gently. 12. Repeat steps 10 and 11 two more times to remove excess impurities. 13. Dry the crude peptide gently using a N2 stream. The crude peptide can be stored under N2 atmosphere at 4
C for several weeks if needed. 14. Analyze and purify the peptides using reverse-phase HPLC and mass spectrometry (see Notes 13 and 14). Collect and lyophilize the purified fractions.

3.5 KAHA Ligation and O-to-N Acyl Shift (Figs. 3 and 4)

1. Prepare 80% DMSO solution containing 0.1 M oxalic acid (see Note 15). 2. Add 32.2 μL reaction solution (final peptide concentration 20 mM) to segments (segment 1, 3.0 mg, 1.0 equiv. and segment 2, 2.3 mg, 1.1 equiv.) pre-weighed into a glass vial and leave to agitate gently at 60
C (see Notes 16 and 17). The reaction should be monitored for completion using an

Chemical Protein Synthesis by Chemoselective α-Ketoacid-. . .

O H 2N

segment 1 (44 aa)

N H

OH

O

159

O

H N

N H

O

segment 2 (30 aa)

OH

i) KAHA ligation DMSO/H2O + oxalic acid 60 ˚C, 16 h

O H 2N

segment 1 (44 aa)

O

N H

NH2

O

segment 2 (30 aa)

N H

OH

ii) O–>N rearrangement NaHCO3/Na2CO3 buffer pH 9.5 rt, 2 h

H 2N

segment 1 (44 aa)

N H

H N O

O N H

segment 2 (30 aa)

OH

OH Ubiquitin (Met1Nle, Ala46Hse)

Fig. 3 Schematic outline of two-segment synthesis of ubiquitin

Fig. 4 HPLC traces of KAHA ligation, O-to-N acyl shift, and purified ubiquitin. Note the minor shift in retention time upon O-to-N acyl shift

analytical HPLC and mass spectrometry (~16 h) (see Notes 13 and 18). 3. Upon full consumption of segment 1, dilute using 0.2 M Na2CO3/NaHCO3 solution to a final concentration of 2 mM. This will form a cloudy solution.

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4. Gently agitate on a shaker at room temperature. The completion of the O-to-N acyl shift should be monitored for completion using an analytical HPLC (~2 h) (see Note 19). 5. Acidify the mixture using CH3CN/H2O (1:1) containing 0.1% (v/v) TFA and purify on a preparative HPLC (see Note 14). 6. Collect and lyophilize the purified fractions (2.1 mg recovered, 38% yield). 7. Dissolve the lyophilized peptide in minimum amount of DMSO and dilute 20-fold using deionized water. Dilute further tenfold with 20 mM NaOAc, 50 mM Gn.HCl folding buffer adjusted to pH 5.0. 8. Remove the DMSO and concentrate the ubiquitin solution to the desired concentration by centrifugal filtration. The buffer can be exchanged to an alternative buffer as desired, using the same centrifugal technique or dialysis.

4

Notes 1. Use a fritted syringe for resin loading and SPPS. 2. Resin should be agitated by either bubbling N2 through the resin suspension or by shaking of the reaction vessel. 3. HCTU and HATU are sensitizers and should only be handled in a well-ventilated area. 4. Desired loading is 0.2–0.3 mmol/g. The loading may need to be optimized depending on the SPPS efficiency. Resins can be loaded more than once prior to capping to achieve the desired loading. 5. Resins should be dried thoroughly by washing multiple times with diethyl ether and drying with a gentle flow of N2. This is essential for long-term storage, loading measurement and final TFA cleavage. The resin should be free-flowing without any clumps at the end of the drying. 6. Resin loading can be measured using UV spectrometer, measuring the absorbance of Fmoc group deprotected at the characteristic wavelength of 304 nm. Deprotection was carried out using 2% DBU solution in DMF as outlined in reference [22]. 7. The resin may turn purple during loading. 8. Detailed step-by-step procedures for automated Fmoc-SPPS are beyond the scope of this protocol. The conditions used for ubiquitin synthesis described herein are outlined briefly in Table 1. 9. Perform a microcleavage during the synthesis to check its success. For a microcleavage, remove a small amount of resin from the reaction (ca. 10 mg), add cleavage cocktail (1 mL), and shake at room temperature for 2 h. Remove TFA using a

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Table 1 Conditions for automated Fmoc-SPPS Step

Reagents

Time

Fmoc deprotection

20% piperidine in DMF (v/v)

2  8 min

Peptide coupling

4.00 equiv. Fmoc-amino acid 3.98 equiv. HCTU 8.00 equiv. NMM

2  45 min

Capping

20% Ac2O in DMF (v/v) 2 M DIPEA in NMP

1  5 min

rotary evaporator and add diethyl ether to precipitate peptide. Pellet the peptide by centrifugation. Dissolve the pellet in CH3CN/H2O (1:1, v/v, 0.1% TFA) and analyze by analytical HPLC and mass spectrometry. 10. Triisopropyl silane (TIPS) is a commonly used scavenger, but this should be avoided for α-ketoacid-containing segments to avoid reduction of the α-ketoacid. 11. TFA is corrosive and toxic. The evaporation should be carried out inside a fume hood. 12. Use a mixture of diethyl ether/pentane (1:1, v/v) for hydrophobic peptides. 13. Analytical RP-HPLC conditions: isocratic 20% eluent B for 3 min then 20–95% buffer B over 22 min with flow rate 1 mL/min at 60
C. 14. Preparative RP-HPLC conditions: isocratic 25% eluent B for 10 min then 25–70% buffer B over 25 min with flow rate 10 mL/min at 60
C. 15. KAHA ligation can be carried out in other solvent mixtures depending on the solubility, such as 1:1 AcOH/HFIP +1% H2O at 45
C [20] or 7:3 NMP/H2O with 0.1 M oxalic acid at 60
C [23]. 16. To aid solubilization, the reaction mixture may be centrifuged and vortexed. Heating of the reaction mixture to 60
C facilitates solubilization. During the course of the reaction, the reaction mixture may turn into a gel. This does not affect the progress of the reaction. 17. The glass vial should be sealed tightly to prevent solvent evaporation. 18. A small aliquot (0.5 μL) from the reaction mixture can be diluted with CH3CN/H2O (30 μL) and stored at 4
C for later analysis. 19. The product peak will undergo a shift to later retention time upon O-to-N acyl shift. Optimization of HPLC conditions may be required to observe this shift in retention time, depending on the amino acid sequence.

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References 1. Kent SBH (2009) Total chemical synthesis of proteins. Chem Soc Rev 38:338–351 2. Bondalapati S, Jbara M, Brik A (2016) Expanding the chemical toolbox for the synthesis of large and uniquely modified proteins. Nat Chem 8:407–418 3. Dawson PE, Muir TW, Clarklewis I et al (1994) Synthesis of proteins by native chemical ligation. Science 266:776–779 4. Agouridas V, El Mahdi O, Diemer V et al (2019) Native chemical ligation and extended methods: mechanisms, catalysis, scope, and limitations. Chem Rev 119:7328–7443 5. Yan LZ, Dawson PE (2001) Synthesis of peptides and proteins without cysteine residues by native chemical ligation combined with desulfurization. J Am Chem Soc 123:526–533 6. Fang GM, Li YM, Shen F et al (2011) Protein chemical synthesis by ligation of peptide hydrazides. Angew Chem Int Edit 50:7645–7649 7. Pattabiraman VR, Ogunkoya AO, Bode JW (2012) Chemical protein synthesis by chemoselective alpha-ketoacid-hydroxylamine (KAHA) ligations with 5-oxaproline. Angew Chem Int Edit 51:5114–5118 8. Wucherpfennig TG, Rohrbacher F, Pattabiraman VR et al (2014) Formation and rearrangement of homoserine depsipeptides and depsiproteins in the alpha-ketoacid-hydroxylamine ligation with 5-oxaproline. Angew Chem Int Edit 53:12244–12247 9. Thuaud F, Rohrbacher F, Zwicky A et al (2016) Incorporation of acid-labile masking groups for the traceless synthesis of C-terminal peptide alpha-Ketoacids. Org Lett 18:3670–3673 10. Thuaud F, Rohrbacher F, Zwicky A et al (2016) Photoprotected peptide alphaketoacids and hydroxylamines for iterative and one-pot KAHA ligations: synthesis of NEDD8. Helv Chim Acta 99:868–894 11. Ogunkoya AO, Pattabiraman VR, Bode JW (2012) Sequential a-ketoacid-hydroxylamine (KAHA) ligations: synthesis of C-terminal variants of the modifier protein UFM1. Angew Chem Int Edit 51:9693–9697 12. Harmand TJ, Murar CE, Bode JW (2016) Protein chemical synthesis by alpha-ketoacidhydroxylamine ligation. Nat Protoc 11:1130–1147 13. Ju L, Lippert AR, Bode JW (2008) Stereoretentive synthesis and chemoselective amide-

forming ligations of C-terminal peptide alphaketoacids. J Am Chem Soc 130:4253–4255 14. Ju L, Bode JW (2009) A general strategy for the preparation of C-terminal peptide alphaketoacids by solid phase peptide synthesis. Org Biomol Chem 7:2259–2264 15. Pusterla I, Bode JW (2015) An oxazetidine amino acid for chemical protein synthesis by rapid, serine-forming ligations. Nat Chem 7:668–672 16. Baldauf S, Schauenburg D, Bode JW (2019) A threonine-forming oxazetidine amino acid for the chemical synthesis of proteins through KAHA ligation. Angew Chem Int Edit 58:12599–12603 17. Baldauf S, Ogunkoya AO, Boross GN et al (2020) Aspartic acid forming alpha-ketoacidhydroxylamine (KAHA) ligations with (S)-4,4difluoro-5-oxaproline. J Org Chem 85:1352–1364 18. Murar CE, Thuaud F, Bode JW (2014) KAHA ligations that form aspartyl aldehyde residues as synthetic handles for protein modification and purification. J Am Chem Soc 136:18140–18148 19. Bang D, Makhatadze GI, Tereshko V et al (2005) Total chemical synthesis and X-ray crystal structure of a protein diastereomer: [D-Gln 35]ubiquitin. Angew Chem Int Edit 44:3852–3856 20. Neumann K, Farnung J, Baldauf S et al (2020) Prevention of aspartimide formation during peptide synthesis using cyanosulfurylides as carboxylic acid-protecting groups. Nat Commun 11:982 21. Murar CE, Harmand TJ, Bode JW (2017) Improved synthesis of (S)-N-Boc-5-oxaproline for protein synthesis with the α-ketoacidhydroxylamine (KAHA) ligation. Bioorg Med Chem 25:4996–5001 22. Gude M, Ryf J, White PD (2002) An accurate method for the quantitation of Fmocderivatized solid phase supports. Lett Pept Sci 9:203–206 23. Wucherpfennig TG, Pattabiraman VR, Limberg FRP et al (2014) Traceless preparation of C-terminal α-ketoacids for chemical protein synthesis by α-ketoacid–hydroxylamine ligation: synthesis of SUMO2/3. Angew Chem Int Ed 53:12248–12252

Chapter 15 Site-Specific Modification of Single-Chain Affinity Ligands for Fluorescence Labeling, Radiolabeling, and Bioconjugation Boya Zhang, Sachith M. Vidanapathirana, and Colin F. Greineder Abstract Single-chain protein affinity ligands are recombinant polypeptides that recreate the antigen-binding site of parental, monoclonal antibodies (mAbs) or present unique binding surfaces derived from display technologies, computational design, or other approaches. These diverse ligands have several advantages over fulllength mAbs as agents for delivery of small molecule, protein, and nanoparticle cargoes to desired sites in the body. However, they present unique challenges for modification and bioconjugation. Fusion of a LPXTGG motif, or “sortag,” and a 5-amino acid, flexible linker to the C-terminus of these affinity ligands enables high-efficiency transpeptidation by the bacterial enzyme, Sortase A, and site-specific addition of fluorophores, radiolabels, or functional groups for oriented and stoichiometrically controlled bioconjugation. We describe in detail this method and address several challenges and pitfalls in the purification and characterization of modified single-chain affinity ligands. Key words Affinity ligand, Sortase, Site-specific modification, Bioconjugation, Labeling

1

Introduction Affinity interactions are the driving force behind the biologic effect of nearly all drugs, including therapeutic monoclonal antibodies (mAbs) [1]. The latter is often referred to as “targeted therapies,” but used alone, their mechanisms of action are not fundamentally different from most small molecule therapeutics—e.g., receptor agonism, antagonism, and allosteric modulation [2–4]. In contrast, when antibodies are conjugated to therapeutic cargo, they act in a distinct manner first conceptualized by Paul Ehrlich in his “magic bullet” hypothesis [5]. In these applications, affinity is no longer directly linked to therapeutic action but rather used to drive accumulation at an intended site of action, reduce off-target side effects, or cross biological barriers to reach otherwise inaccessible drug

Waleed M. Hussein, Rachel J. Stephenson and Istvan Toth (eds.), Peptide Conjugation: Methods and Protocols, Methods in Molecular Biology, vol. 2355, https://doi.org/10.1007/978-1-0716-1617-8_15, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

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targets [6, 7]. The development of anti-neoplastic antibody-drug conjugates (ADCs) and their recent success in the clinic have brought these concepts back into the limelight and focused attention on the bioconjugation of antibodies to cargo [8, 9]. Simultaneously, advances in recombinant DNA technology, display techniques for in vitro evolution, and computational modeling have expanded the library of available affinity ligands beyond traditional hybridoma-derived or recombinant mAbs [10–12]. Most of the newer agents in the drug targeting armamentarium are so-called “single-chain affinity ligands,” synthesized as a single polypeptide by prokaryotic or eukaryotic cell factories. The best known are single-chain variable fragments (scFv), derived from the variable heavy (VH) and light (VL) chains of mammalian immunoglobulins. More recent additions include camelid and cartilaginous fish-derived single domain antibodies (sdAb) and combinatorially engineered proteins like affibodies, DARPins, and centyrins [13– 15]. Each of these affinity ligands differs substantially in structure and in vivo behavior from full-length immunoglobulins. Even scFv, which closely resembles the mAb antigen-binding domain, is only ~1/5th the size and lacks a fragment crystallizable (Fc) domain, resulting in marked differences in pharmacokinetics, tissue penetration, and accessibility to sterically hindered epitopes [16, 17]. Furthermore, monovalent interaction with target antigens results in distinct binding kinetics and slower rates of cellular internalization than those induced by bivalent mAbs. Overall, each class of singlechain affinity ligand has distinct properties and in vivo behavior that lends themselves to specific drug delivery applications, including molecular imaging, cell surface anchoring, intracellular siRNA delivery, bispecific t-cell engagement, and others. For all of these advantages, single-chain affinity ligands present a number of challenges with regard to modification for radiotracing, fluorescent imaging, and bioconjugation of therapeutic cargo [18]. Small size and lack of Fc domain result in greater sensitivity to modification of key amino acid side chains like primary amines and free thiols. Likewise, the lack of a flexible hinge region makes these affinity ligands less tolerant of covalent attachment to other proteins or nanoparticles [19]. Non-selective N-hydroxysuccinimide esters, imidoesters, and maleimides, which remain the primary means of antibody modification and conjugation, are more likely to compromise the structure and function of single-chain affinity ligands, which typically require oriented and stoichiometrically controlled bioconjugation [20]. One solution to these challenges is that of enzymatic protein labeling, in which a recombinant affinity ligand is tagged with a specific amino acid sequence at the desired site of modification [21, 22]. The tag is recognized by an enzyme which then makes a site-specific and typically monomolecular modification, preserving antigen binding and enabling controlled, oriented bioconjugation.

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Amongst the enzymes developed for this purpose, sortase A (SrtA), a calcium-dependent transpeptidase from Staphylococcus aureus, stands out as perhaps the most widely employed and versatile, capable of both N- and C-terminal modification and attachment of nearly any functional group or label [23–25]. SrtA recognizes the sequence LPXTGG, referred to as a “sortag,” and cleaves the peptide bond between threonine and glycine, forming an acylenzyme intermediate. SrtA reforms the peptide bond and recycles itself using either the original C-terminal fragment or any other available peptide or protein bearing an N-terminal glycine (2–3 glycines ensure maximal incorporation) [26]. By using an excess of the peptide, the reversible reaction is driven towards transpeptidation and the desired C-terminal modification [27]. In this chapter, we describe the application of this technique to a wide variety of single-chain affinity ligands, including scFv, sdAb, and affibodies, via genetic fusion of a C-terminal sortag with a short, flexible linker to ensure accessibility to the sortase enzyme and robust site-specific modification [18]. Our group and others have validated this approach for the attachment of both fluorescent and radioactive labels with minimal effect on affinity [16, 18, 20, 25, 28]. Moreover, we have modified affinity ligands with functional groups and shown covalent, oriented, and stoichiometrically controlled bioconjugation to both protein and nanoparticle cargoes [16, 18, 29]. We review not only the basic procedure but key considerations in setting up the SrtA reaction and purifying the desired reaction product. In particular, we highlight a critical side reaction, which can adversely affect yield.

2

Materials 1. Single-chain affinity ligand with a 5-amino acid linker and C-terminal sortag and purification tag, e.g., scFv-linkerLPETGG-FLAGx3 or VHH-linker-LPETGG-Hisx6 (Fig. 1a and Note 1). 2. GG-peptide with desired C-terminal modifications (Fig. 1a and Note 2). 3. Sortase enzyme, typically A5 mutant (see Note 3). 4. Tris-buffered saline (TBS): 50 mM Tris–HCl, 150 mM NaCl, pH 7.5. 5. CaCl2: 100 mM stock solution in DI water. 6. Size-exclusion High Performance Liquid Chromatography (SEC-HPLC) or SDS-page gel electrophoresis for reaction mixture analysis (see Note 4). 7. Nickel-charged affinity resin, e.g., Ni-NTA agarose. 8. Centrifugal filters, e.g., Amicon® Ultra.

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Fig. 1 (a) Schematic showing reaction of various sortagged single-chain affinity ligands (scFv, nanobody, affibody) with different functionalized -GG peptides to achieve site-specific modification with fluorescent tags, radiolabels, and functional groups for controlled bioconjugation. (b) SEC-HPLC analysis of an unpurified sortase reaction mixture (left panel). Appearance of a fluorescent peak at the elution time of the affinity ligand (*) indicates successful transpeptidation. While useful for confirming a successful reaction, the relative area

Site-Specific Modification of Single-Chain Affinity Ligands for. . .

3 3.1

167

Methods Reaction Setup

1. Thaw reagents as necessary. For long-term usage, we store single-chain affinity ligands, -GG peptide, and SrtA enzyme in aliquots at
20  C or
80  C. Once thawed, aliquots are kept at 4  C for up to 6 months. 2. Calculate the required volume of each component based on the stock solution and the desired concentration of each component (see Note 5). Table 1 may be used for convenience. Remember to add SrtA enzyme after adding the -GG peptide to minimize the sortase side reaction (see Note 6). 3. Allow the reaction to come to equilibrium. We typically incubate reactions overnight at room temperature, although shorter times (e.g., 4 h) may be adequate for some affinity ligands and peptides. 37  C is also possible and will reach equilibrium faster but may result in lower yields by favoring the sortase side reaction (see Note 6).

3.2 Reaction Mixture Analysis

1. Reaction progression can be monitored over time or confirmed at a chosen endpoint using one of two methods: (1) Sizeexclusion HPLC (especially helpful if a fluorescent peptide is used and a fluorescence detector is available), (2) Gel electrophoresis. To analyze, take a small volume from the reaction mixture and dilute in HPLC or sample buffer (see Note 4). 2. For SEC-HPLC with a fluorescence detector, 1 μL of a standard reaction mixture (i.e., 10 μM affinity ligand) is enough to visualize a shift in the fluorescent signal from the peptide elution peak(s) to that of the affinity ligand (Fig. 1b). Alternatively, if the affinity ligand has a large-sized tag on the C-terminus (i.e., after the sortag), as in the case of scFvlinker-LPETGG-FLAGx3, then modification may be observed as a shift of the size of the protein (Fig. 1b). If HPLC is not available, SDS-PAGE can be used to analyze reaction mixtures (Fig. 1c). In some cases, depending on the size of the affinity ligand and the peptide being added, gel electrophoresis may even be capable of distinguishing transpeptidation from truncation (i.e., the sortase side reaction) (Fig. 2b).

ä Fig. 1 (continued) under the curve (AUC) of the peaks typically underestimates reaction efficiency as compared to SDS-PAGE or the final yield obtained upon purification. Right panel shows the purified reaction product (green line) shifted to the right relative to the unmodified affinity ligand (black line). (c) SDS-PAGE analysis: Lane 1—ladder, 2—unmodified affinity ligand (scFv-linker-LPETGG-FLAGx3), 3—Reaction mixture without SrtA enzyme, 4—Reaction mixture with SrtA, 5—purified reaction product. The latter shows the desired C-terminal modified scFv-FAM-azide on fluorescence imaging, while Coomassie staining reveals a mix of desired product, unmodified affinity ligand, and truncated form (scFv-LPET)

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Table 1 Table used for calculations for setup of sortase reactions Component

Stock concentration

Final concentration

Dilution

Volume (μL)

scFv-LPET

20 μM

10 μM

2

100

Sortase A5

100 μM

1 μM

100

2

Ca2+

50 mM

1 mM

50

4

GGG peptide

10 mM

50 μM

200

1

–

–

–

93

TBS Total volume

200

Fig. 2 (a) Schematic showing transpeptidation and the sortase side reaction. (b) Optimization of a sortase reaction analyzed by SDS-PAGE. Increasing concentrations of sortase initially result in greater reaction product, with an optimal concentration of 1.25 nM. Under these conditions, all of the affinity ligand is reacted, but no truncated form can be seen on Coomassie staining. At higher concentrations of sortase, however, the side reaction prevails with complete loss of C-terminal modified reaction product at 20 nM 3.3 Reaction Mixture Purification

1. At equilibrium, the sortase reaction mixture contains as many as six distinct proteins/polypeptides (Fig. 2a): (1) desired reaction product (i.e., C-terminal modified affinity ligand), (2) unreacted affinity ligand, (3) excess peptide, (4) C-terminal polypeptide removed from the original affinity ligand (e.g., GG-FLAGx3), (5) sortase A enzyme, and (6) side reaction product. Whenever possible, the first step should always be to remove sortase from the reaction mixture, as premature removal of peptide will result in immediate shunting of product

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to the side reaction (see Note 6). To accomplish this, we typically use 100 μL of nickel-charged affinity resin per mL of reaction mixture, although the amount required will vary based on the concentration of SrtA enzyme used. Of note, if the affinity ligand has a C-terminal Hisx6 tag after the sortag (see Note 1), this step will also remove any unreacted affinity ligand. In our experience, this results in a sample of scfv with a degree of modification of nearly 100%. 2. Wash nickel-affinity ligand twice with TBS, then add to the reaction mixture and incubate at room temperature for 30 min while rotating. Spin down the affinity resin at 2000  g for 5 min and gently remove the supernatant to a clean tube, while not disturbing the agarose (see Note 7). 3. Nickel resin can be washed with TBS to recover additional products. Spin to pellet affinity resin and add the supernatant from the TBS wash to that from step 2. 4. Once sortase has been completely removed, the excess peptide can be removed via desalting column or centrifugal filter. For larger single-chain affinity ligands (e.g., scFv), we use a molecular weight cutoff (MWCO) of 10 kDa, whereas, for smaller proteins like nanobodies or affibodies, we have successfully used 3 kDa MWCO. Spin repeatedly at 4–8000  g until free peptide has been completely removed. The final purified product should be analyzed by SEC-HPLC and SDS-PAGE (see Note 8).

4

Notes 1. Single-chain affinity ligands are expressed with a C-terminal sortag (LPETGG) separated from the protein by a 5-amino acid linker—both GGGGS and GSSSG have been used with good results. Sortagged affinity ligands have been successfully expressed in multiple protein production systems, including bacterial cytoplasm and periplasm and the secretory pathway of yeast, insect, and mammalian cells. A purification tag—e.g., FLAGx3 or Hisx6—is typically incorporated after the C-terminal sortag, so that it is removed by transpeptidation. The Hisx6 tag is particularly useful, as the unreacted affinity ligand is then removed by the nickel-charged affinity resin used to capture and remove the sortase enzyme. 2. We typically use -GG or -GGG peptides (i.e., 2 or 3 N-terminal glycines) synthesized by standard solid-phase synthesis. A fluorescent amino acid is typically incorporated into each peptide— even if fluorescent imaging is not the intended application—to facilitate analysis of the reaction mixture, optimization of

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reaction conditions, and quantification of yield and degree of modification (Figs. 1 and 2 and Note 4). Other desired modifications and functional groups are typically introduced on the final (i.e., C-terminal) amino acid of the peptide. For example, a single-chain affinity ligand intended for bioconjugation via click chemistry may be modified with an azide via reaction with the peptide GGG[K]GGSK-N3, where [K] ¼ Lys(5-FAM) and K-N3 ¼ azidolysine. 3. Single-chain affinity ligands can be modified efficiently by wildtype Ca2+-dependent sortase A from S. aureus or a number of mutants. These include: (1) Δ59, a truncated form, in which 59 N-terminal amino acids (the transmembrane domain) have been replaced by a Hisx6 tag [27], (2) Sortase A5 (alternately referred to as “5 M” sortase), a pentamutant of Δ59 that is significantly more active [30], or (3) “7 M” sortase, which adds two additional amino acid substitutions to the pentamutant to create a variant that is both highly active and relatively calciumindependent [31]. The reactions shown here were performed using sortase A5 produced in E. coli as previously described [20]. 4. Analysis of sortase reaction mixtures is critical and can be accomplished using SEC-HPLC or gel electrophoresis. Our size-exclusion column is used with a fluorescence detector to measure the transfer of fluorescent signal from the peptide elution peak to that of the modified single-chain affinity ligand, allowing us to track the kinetics and/or overall efficiency of a given reaction (Fig. 1b). The same can be done with SDS-PAGE (Fig. 1c), although quantification of reaction progression by these two methods does not always agree. 5. Whenever possible, sortagged single-chain affinity ligands should be modified at 10 μM concentration or higher. The efficiency of the SrtA reaction decreases at lower substrate concentrations, especially below 5 μM. To maximize yield, a 5:1 ratio of peptide:affinity ligand is optimal (e.g., 50 μM peptide, 10 μM affinity ligand), although lower ratios can be used if peptide is limited. The optimal sortase concentration depends on many factors, including: (1) the type of sortase used (see Note 3), (2) the affinity ligand being modified, and (3) the concentrations of affinity ligand and peptide. As such, we recommend setting up a series of test reactions (Fig. 2) and use SEC-HPLC or SDS-PAGE to quantify both transpeptidation and the sortase side reaction (see Note 6). 6. What our group refers to as the “sortase side reaction” (Fig. 2a) is a key consideration that has been discussed in some [18, 32, 33], but not all reports of sortase technology. In the absence of -GG peptide or other substrates (e.g., a protein containing an

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N-terminal glycine), the acyl-enzyme intermediate formed between SrtA and sortagged affinity ligand hydrolyzes, leaving a truncated form of the latter (Fig. 2b). Critically, this truncated protein lacks a threonine-glycine bond and, therefore, cannot be regenerated for transpeptidation [32]. Several rules-of-thumb can help to minimize the impact of the side reaction. First, sortase enzyme should always be added to reaction mixtures after -GG peptide. Likewise, SrtA must be removed from reaction mixtures first. Second, we favor running reactions at room temperature, rather than 37  C, as the latter decreases the ratio of modified:truncated affinity ligand [18]. Finally, as discussed in Note 5, the concentration of SrtA should be optimized for any given reaction. In our experience, it is the amount of enzyme added relative to the concentrations of affinity ligand and peptide which determines the relative rate of transpeptidation vs. side reaction (Fig. 2b). Sortase mutants with higher activity, like A5 and 7 M, demonstrate not only faster transpeptidation but also side reaction, so optimization must be repeated when changing from one type of sortase to another. 7. Residual sortase A, even at very low concentration, will slowly destroy modified affinity ligand via the sortase side reaction if it is not fully eliminated before removal of the -GG peptide. As such, care should be taken not to carry over any nickel-charged affinity resin, and modified proteins that are stored for prolonged periods are intermittently monitored to ensure that C-terminal modification is not being slowly removed. If the calcium-dependent enzyme is used, purified proteins can be stored in 1 mM EDTA to inhibit any residual SrtA. 8. Note that all sortase reaction products typically contain all three forms of the protein—unreacted, modified, and truncated (Fig. 1b)—unless the affinity ligand contains a Hisx6 tag after the sortag, in which case the unreacted protein can be selectively eliminated. Careful analysis with SEC-HPLC and SDS-PAGE can help determine the relative amount of each protein species. References 1. Yamada T (2011) Therapeutic monoclonal antibodies. Keio J Med 60(2):37–46. https:// doi.org/10.2302/kjm.60.37 2. Bernard-Marty C, Lebrun F, Awada A, Piccart MJ (2006) Monoclonal antibody-based targeted therapy in breast cancer: current status and future directions. Drugs 66 (12):1577–1591. https://doi.org/10.2165/ 00003495-200666120-00004

3. Suzuki M, Kato C, Kato A (2015) Therapeutic antibodies: their mechanisms of action and the pathological findings they induce in toxicity studies. J Toxicol Pathol 28(3):133–139. https://doi.org/10.1293/tox.2015-0031 4. Castelli MS, McGonigle P, Hornby PJ (2019) The pharmacology and therapeutic applications of monoclonal antibodies. Pharmacol Res Perspect 7(6):e00535. https://doi.org/ 10.1002/prp2.535

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5. Witkop B (1999) Paul Ehrlich and his magic bullets--revisited. Proc Am Philos Soc 143 (4):540–557 6. Kiseleva RY, Glassman PM, Greineder CF, Hood ED, Shuvaev VV, Muzykantov VR (2018) Targeting therapeutics to endothelium: are we there yet? Drug Deliv Transl Res 8 (4):883–902. https://doi.org/10.1007/ s13346-017-0464-6 7. Greineder CF, Howard MD, Carnemolla R, Cines DB, Muzykantov VR (2013) Advanced drug delivery systems for antithrombotic agents. Blood 122(9):1565–1575. https:// doi.org/10.1182/blood-2013-03-453498 8. Agarwal P, Bertozzi CR (2015) Site-specific antibody-drug conjugates: the nexus of bioorthogonal chemistry, protein engineering, and drug development. Bioconjug Chem 26 (2):176–192. https://doi.org/10.1021/ bc5004982 9. Mullard A (2013) Maturing antibody-drug conjugate pipeline hits 30. Nat Rev Drug Discov 12(5):329–332. https://doi.org/10. 1038/nrd4009 10. Greineder CF, Hood ED, Yao A, Khoshnejad M, Brenner JS, Johnston IH, Poncz M, Gottstein C, Muzykantov VR (2016) Molecular engineering of high affinity single-chain antibody fragment for endothelial targeting of proteins and nanocarriers in rodents and humans. J Control Release 226:229–237. https://doi.org/10.1016/j. jconrel.2016.02.006 11. Hu Y, Liu C, Muyldermans S (2017) Nanobody-based delivery systems for diagnosis and targeted tumor therapy. Front Immunol 8:1442. https://doi.org/10.3389/fimmu. 2017.01442 12. Altai M, Liu H, Ding H, Mitran B, Edqvist PH, Tolmachev V, Orlova A, Gr€aslund T (2018) Affibody-derived drug conjugates: potent cytotoxic molecules for treatment of HER2 overexpressing tumors. J Control Release 288:84–95. https://doi.org/10.1016/j. jconrel.2018.08.040 13. Plu¨ckthun A (2015) Designed ankyrin repeat proteins (DARPins): binding proteins for research, diagnostics, and therapy. Annu Rev Pharmacol Toxicol 55:489–511. https://doi. org/10.1146/annurev-pharmtox-010611134654 14. Goldberg SD, Cardoso RMF, Lin T, SpinkaDoms T, Klein D, Jacobs SA, Dudkin V, Gilliland G, O’Neil KT (2016) Engineering a targeted delivery platform using Centyrins. Protein Eng Des Sel 29(12):563–572. https://doi.org/10.1093/protein/gzw054

15. Ko¨nning D, Zielonka S, Grzeschik J, Empting M, Valldorf B, Krah S, Schro¨ter C, Sellmann C, Hock B, Kolmar H (2017) Camelid and shark single domain antibodies: structural features and therapeutic potential. Curr Opin Struct Biol 45:10–16. https://doi.org/ 10.1016/j.sbi.2016.10.019 16. Kiseleva RY, Glassman PG, LeForte KM, Walsh LR, Villa CH, Shuvaev VV, Myerson JW, Aprelev PA, Marcos-Contreras OA, Muzykantov VR, Greineder CF (2020) Bivalent engagement of endothelial surface antigens is critical to prolonged surface targeting and protein delivery in vivo. FASEB J 34 (9):11577–11593. https://doi.org/10.1096/ fj.201902515RR 17. Tsai WK, Wu AM (2018) Aligning physics and physiology: engineering antibodies for radionuclide delivery. J Labelled Comp Radiopharm 61(9):693–714. https://doi.org/10.1002/ jlcr.3622 18. Greineder CF, Villa CH, Walsh LR, Kiseleva RY, Hood ED, Khoshnejad M, WardenRothman R, Tsourkas A, Muzykantov VR (2018) Site-specific modification of singlechain antibody fragments for bioconjugation and vascular immunotargeting. Bioconjug Chem 29(1):56–66. https://doi.org/10. 1021/acs.bioconjchem.7b00592 19. Albrecht H, Burke PA, Natarajan A, Xiong CY, Kalicinsky M, DeNardo GL, DeNardo SJ (2004) Production of soluble ScFvs with Cterminal-free thiol for site-specific conjugation or stable dimeric ScFvs on demand. Bioconjug Chem 15(1):16–26. https://doi.org/10. 1021/bc030018+ 20. Glassman PM, Walsh LR, Villa CH, MarcosContreras OA, Hood ED, Muzykantov VR, Greineder CF (2020) Molecularly engineered nanobodies for tunable pharmacokinetics and drug delivery. Bioconjug Chem 31 (4):1144–1155. https://doi.org/10.1021/ acs.bioconjchem.0c00003 21. Rabuka D (2010) Chemoenzymatic methods for site-specific protein modification. Curr Opin Chem Biol 14(6):790–796. https://doi. org/10.1016/j.cbpa.2010.09.020 22. Zhang Y, Park K-Y, Suazo KF, Distefano MD (2018) Recent progress in enzymatic protein labelling techniques and their applications. Chem Soc Rev 47(24):9106–9136. https:// doi.org/10.1039/C8CS00537K 23. Popp MW, Antos JM, Ploegh HL (2009) Sitespecific protein labeling via sortase-mediated transpeptidation. Curr Protoc Protein Sci Chapter 15:Unit 15.13. https://doi.org/10. 1002/0471140864.ps1503s56

Site-Specific Modification of Single-Chain Affinity Ligands for. . . 24. Theile CS, Witte MD, Blom AEM, Kundrat L, Ploegh HL, Guimaraes CP (2013) Site-specific N-terminal labeling of proteins using sortasemediated reactions. Nat Protoc 8 (9):1800–1807. https://doi.org/10.1038/ nprot.2013.102 25. Warden-Rothman R, Caturegli I, Popik V, Tsourkas A (2013) Sortase-tag expressed protein ligation: combining protein purification and site-specific bioconjugation into a single step. Anal Chem 85(22):11090–11097. https://doi.org/10.1021/ac402871k 26. Antos JM, Popp MW, Ernst R, Chew GL, Spooner E, Ploegh HL (2009) A straight path to circular proteins. J Biol Chem 284 (23):16028–16036. https://doi.org/10. 1074/jbc.M901752200 27. Guimaraes CP, Witte MD, Theile CS, Bozkurt G, Kundrat L, Blom AEM, Ploegh HL (2013) Site-specific C-terminal and internal loop labeling of proteins using sortasemediated reactions. Nat Protoc 8 (9):1787–1799. https://doi.org/10.1038/ nprot.2013.101 28. Wang HH, Altun B, Nwe K, Tsourkas A (2017) Proximity-based sortase-mediated ligation. Angew Chem Int Ed Engl 56 (19):5349–5352. https://doi.org/10.1002/ anie.201701419 29. Hood ED, Greineder CF, Shuvaeva T, Walsh L, Villa CH, Muzykantov VR (2018)

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Vascular targeting of radiolabeled liposomes with bio-orthogonally conjugated ligands: single chain fragments provide higher specificity than antibodies. Bioconjug Chem 29 (11):3626–3637. https://doi.org/10.1021/ acs.bioconjchem.8b00564 30. Chen I, Dorr BM, Liu DR (2011) A general strategy for the evolution of bond-forming enzymes using yeast display. Proc Natl Acad Sci U S A 108(28):11399–11404. https:// doi.org/10.1073/pnas.1101046108 31. Hirakawa H, Ishikawa S, Nagamune T (2015) Ca2+ -independent sortase-A exhibits high selective protein ligation activity in the cytoplasm of Escherichia coli. Biotechnol J 10 (9):1487–1492. https://doi.org/10.1002/ biot.201500012 32. Mo¨hlmann S, Mahlert C, Greven S, Scholz P, Harrenga A (2011) In vitro sortagging of an antibody fab fragment: overcoming unproductive reactions of sortase with water and lysine side chains. Chembiochem 12 (11):1774–1780. https://doi.org/10.1002/ cbic.201100002 33. Heck T, Pham P-H, Yerlikaya A, Tho¨nyMeyer L, Richter M (2014) Sortase A catalyzed reaction pathways: a comparative study with six SrtA variants. Cat Sci Technol 4 (9):2946–2956. https://doi.org/10.1039/ C4CY00347K

Chapter 16 Preparation and Characterization of Quantum Dot-Peptide Conjugates Based on Polyhistidine Tags Katherine D. Krause, Hsin-Yun Tsai, Kelly Rees, Hyungki Kim, and W. Russ Algar Abstract Quantum dots (QDs) offer bright and robust photoluminescence among several other advantages in comparison to fluorescent dyes. In order to leverage the advantageous properties of QDs for applications in bioanalysis and imaging, simple and reliable methods for bioconjugation are required. One such method for conjugating peptides to QDs is the use of polyhistidine tags, which spontaneously bind to the surface of QDs. We describe protocols for assembling polyhistidine-tagged peptides to QDs and for characterizing the resultant QD-peptide conjugates. The latter include both electrophoretic and FRET-based protocols for confirming successful peptide assembly, estimating the maximum peptide loading capacity, and measuring the assembly kinetics. Sensors for protease activity and intracellular delivery are briefly noted as prospective applications of QD-peptide conjugates. Key words Quantum dots, Peptides, Polyhistidine, Fo¨rster resonance energy transfer (FRET), Agarose gel electrophoresis, Polyacrylamide gel electrophoresis

1

Introduction The advent of accurate, low-cost synthetic methods for peptides has opened the door for the myriad applications of peptides in use today. At least 60 peptide therapeutics have been approved for use by the United States Food and Drug Administration (FDA), with hundreds more in clinical trials [1, 2]. Peptides are also of growing interest as vaccines [3–5], antimicrobial agents [6–8], and targeting and biorecognition agents [9–12]. Protocols to label peptides with fluorophores, radiolabels, mass tags, and electrochemically active labels are essential to the bioanalytical and imaging applications of peptides. Although dye-labeled peptides have been used extensively for a variety of applications, dyes have numerous shortcomings as fluorescent labels. The susceptibility of dyes to photobleaching can

Waleed M. Hussein, Rachel J. Stephenson and Istvan Toth (eds.), Peptide Conjugation: Methods and Protocols, Methods in Molecular Biology, vol. 2355, https://doi.org/10.1007/978-1-0716-1617-8_16, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

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limit their use for long-term measurements; relative to other materials, the broad and asymmetric emission spectra and narrow absorption spectra of dyes may impede multicolor analysis and imaging; and dyes are only moderately bright. These limitations can be addressed by using alternative fluorophores, such as colloidal semiconductor nanocrystals. Better known as “quantum dots” (QDs), these materials are brighter than dyes, have superior resistance to photobleaching, and, in many cases, more desirable spectral properties [13–15]. The popularity of QDs as luminescent labels can, in part, be attributed to their advantageous optical properties; however, another significant advantage of QDs is their nanoscale structure, which offers opportunities for assay and imaging probe formats and applications that are not possible with molecular fluorophores [14, 16, 17]. The utility of peptides, in combination with the properties of QDs, is a powerful tool for bioanalysis and imaging. 1.1

Quantum Dots

QDs are typically 10 nm or less in diameter and comprise hundreds to thousands of atoms [15, 17]. An electron micrograph of QDs is shown in Fig. 1a and this small size imbues QDs with bright photoluminescence (PL, Note 1). The most widely used QDs are cadmium chalcogenide (CdX, X ¼ S, Se, Te) QDs and their core/ shell and core/shell/shell analogs (e.g., CdSe/ZnS, CdSe/CdS/ ZnS). The use of an epitaxially grown semiconductor shell around the core nanocrystal serves several purposes: the shell passivates any surface defects on the core, thereby improving the PL quantum yield (QY); it protects the core from oxidative or other environment-induced damage; and it minimizes cytotoxicity by preventing degradation of the core and seepage of heavy metal ions (see Note 2) [14, 15, 19–24]. The range of accessible PL wavelengths of a QD is determined by its constituent material. CdX QDs, for example, have PL wavelengths that span the visible spectrum [17], facilitating their use in conjunction with or in lieu of common fluorescent dyes. Other QD materials (e.g., PbX) offer near-infrared (NIR) emission, with brighter and longer-wavelength emission than NIR dyes, which is advantageous for in vivo imaging. QDs of a given composition are characterized by continuously tunable, size-dependent PL, where increasing nanocrystal diameter yields longer-wavelength emission, as illustrated in Fig. 1b. QDs typically have larger molar absorptivity than dyes (104– 107 M1 cm1 versus 104–105 M1 cm1 for dyes) [13–15] and similar PL QYs, making them 1–2 orders of magnitude brighter [13–15]. This brightness, coupled with excellent resistance to photobleaching, makes QDs ideal for applications where highsensitivity detection or long-term imaging is required. QDs are also characterized by broad absorbance spectra and narrow, symmetric PL emission spectra (typical full-width-at-half-maximum of ~30 nm for CdX in the visible region or ~90 nm for PbX in the

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Fig. 1 (a) TEM images of CdSe/CdS/ZnS QD605. The scale bar is 30 nm. (Reproduced with permission from Ref. (18). Copyright 2018 American Chemical Society). (b) Size dependence of CdSe QD absorbance and PL emission. (Reproduced with permission from Ref. (19). Copyright 2010 American Chemical Society). (c) Cartoon schematic of an aqueous-phase QD, including its core, shell, and hydrophilic coating. The binding of hexahistidine-tagged peptide to the shell is also illustrated. (Not drawn to scale)

NIR) [13]. The narrow PL spectra facilitate multicolor detection by minimizing spectral crosstalk between different colors of emitters, while the broad absorbance spectra enable concurrent excitation of all colors of QD at a wavelength well separated from their emissions. QD PL is also typically offset from the wavelength range of strongest absorbance by at least 100 nm [15], enabling facile use of both the optimal excitation and optimal emission wavelength ranges for imaging purposes. 1.2 Applications of QD-Peptide Conjugates

QD-peptide conjugates, such as those depicted in Fig. 1c, are used for a variety of research applications, including but not limited to targeted imaging [25–29], drug delivery [30–33], and Fo¨rster resonance energy transfer (FRET)-based assays and probes for proteases and kinases (see Note 3) [17, 34–36].

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In the case of imaging and drug delivery, QDs conjugated with cell-penetrating peptides (e.g., p160 [31], HIV-TAT peptide derivatives [32, 37], Hph-1 derivatives [33]) and targeting peptides (e.g., F3 tumor-homing peptide [30], RGD-containing peptides for tumor targeting [26], vascular cell adhesion molecule-1 binding peptide [25]) have been common motifs. Peptides have been used to deliver QDs injected intravenously into mice to the lungs and to the blood and lymphatic vessels of tumors [38], adipose tissue [39], brain tumors [40], and squamous cell carcinoma tumors [41] for imaging purposes. The co-conjugation of QDs with cell-penetrating peptides and either siRNA [30, 33] or antisense oligonucleotides [31] has been used to downregulate the expression of target proteins. Targeted subcellular delivery of QD-peptide conjugates has also been achieved via nuclear localization sequence peptides [42] and the co-conjugation of cellpenetrating peptides and an antibody for a target protein within cells [43]. Figure 2a shows examples of cellular uptake of

Fig. 2 Examples of applications of QD-peptide conjugates. (a) Intracellular delivery of QD-peptide conjugates: (i) cartoon representation of the modular cell-delivery peptide assembled on a QD; (ii) targeted cytosolic delivery of QD550; and (iii) images from the injection of QD-peptide conjugates into a developing embryonic chick spinal column, including the distribution of QD PL and a merged image of the QD PL and fluorescence from the nuclear and extranuclear RNA stains. Scale bars are 50 μm. All images were acquired 48 h postinjection. (Reproduced with permission from Ref. (44). Copyright 2013 American Chemical Society). (b) Cartoon representation (left) of a QD-peptide conjugate and FRET for detection of protease activity. FRET pathways are lost with hydrolysis of the dye-labeled peptide. Protease activity was tracked (right) by monitoring the dye/QD PL ratio as a function of time. Increasing enzyme concentrations (0.1–200 nM in direction of arrow) yielded increasing rates of proteolysis. (Reproduced with permission from Ref. (18). Copyright 2018 American Chemical Society)

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QD-peptide conjugates, where the peptide sequences were developed to facilitate endosomal uptake of the QDs by cells and their subsequent escape into the cellular cytosol [44]. Assays for protease activity frequently utilize QDs conjugated with dye-labeled peptides. As illustrated in Fig. 2b, proteolysis of the peptides and diffusion of the dye away from the QD turns off FRET (see Note 4) between the QD and dye, and protease activity is measured by the decrease in dye PL, increase in QD PL, or, more commonly, the dye/QD emission ratio. Such configurations have been used to measure the activity of HIV-1 protease [45, 46], caspases 1 and 3 [47–49], various matrix metalloproteinases [48, 50–52], botulinum neurotoxin protease [53], thrombin [54–56], and others. We have previously described protocols for quantifying protease activity using QD-peptide conjugates [57] and have reviewed these systems in detail [58]. Similar assays for kinase activity most commonly utilize dye-labeled anti-phosphoamino acid antibodies that bind to the QD-bound peptide upon its phosphorylation by a kinase, turning on FRET [59–61]. The kinase activity is measured by the increase in FRET-sensitized dye emission or dye/QD emission ratio. Alternative configurations have used dye-labeled adenosine triphosphate [61] or streptavidincoated QDs in combination with dye-labeled peptide and biotinylated adenosine triphosphate [62]. In each of the foregoing applications, and many others, the successful and reproducible conjugation of peptides to a QD is essential. 1.3 Surface Chemistry and Bioconjugation of QDs

Unlike dyes, QDs have a nontrivial surface area, and this surface area plays a critical role in determining their properties. QDs are typically synthesized via methods that yield nanocrystals coated with hydrophobic ligands. These hydrophobic QDs can be transferred into aqueous media using either a ligand exchange or polymer encapsulation process [14, 15, 17]. In a ligand exchange reaction, the initial hydrophobic ligands are replaced with new ligands that have a QD-binding moiety (e.g., thiol or imidazole group) and a hydrophilic moiety (e.g., carboxylate, zwitterion, polyethylene glycol) that enables aqueous dispersion. When QDs are encapsulated within amphiphilic polymers, the hydrophobic moiety of the polymer interacts with the hydrophobic ligands on the QD and the hydrophilic moiety of the polymer enables dispersion in water. QD surface chemistry should be selected judiciously for a given application, as the choice of ligand or polymer may affect the PL properties (see Note 5), colloidal stability (see Note 6), and compatible bioconjugation strategies of the QDs (see Notes 7 and 8). A notable difference versus dyes is the “nanoscaffold” character of QDs: whereas one or more dye molecules are typically conjugated to a single biomolecule [13], the QD surface can

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accommodate as many as 50 peptides or 30 proteins [63], 3–8 antibodies [64, 65], 15–20 antibody fragments [65, 66], or 25–30 oligonucleotides [67, 68]. This capacity confers advantages for the development of assays with increased avidity and sensitivity [14, 17, 69] and the design of single-vector multifunctional materials [16, 31, 37, 70, 71]. General strategies for coupling biomolecules to QDs include covalent chemistries (e.g., heterobifunctional crosslinkers, click chemistries, carbodiimide coupling); direct coordination of a biomolecule to the QD nanocrystal through thiols or polyhistidine (Hisn) tags; secondary interactions (e.g., biotin-streptavidin); and electrostatic binding of charged biomolecules to oppositely charged QDs. Table 1 lists representative examples of these strategies applied to the preparation of QD-peptide conjugates (see Note 9) [25–28, 31, 42, 72–74]. Readers may refer to a number of excellent reviews on the bioconjugation of QDs for more information [17, 36, 83, 84]. 1.4 Bioconjugation of QDs via Polyhistidine Tags

Among the array of available bioconjugate chemistries, polyhistidine tags are widely used for assembling peptides and other biomolecules onto QDs. Hexahistidine tags are most common, but tetrahistidine tags [63, 85, 86], octahistidine tags [63, 86, 87], and peptides with multiple polyhistidine tags [88] have also been used. The imidazole rings of polyhistidine bind to Zn2+ sites at the surface of ZnS-shelled QDs. Binding is spontaneous at room temperature and equilibrium is usually reached within minutes [86]. The polyhistidine tag has high affinity for the QD surface across a broad range of pH values [17], with reported dissociation constants (Kd) of 0.1–100 nM [86]. This high affinity affords simple control over the number of biomolecules attached per QD because the assembly proceeds essentially quantitatively. The average number of biomolecules per QD is thus controlled by the stoichiometry (see Note 10) [89]. Post-conjugation purification steps also tend to be unnecessary. Further advantages of polyhistidine tags include control over biomolecule orientation and no undesired crosslinking because there is typically only one polyhistidine tag per biomolecule, good reproducibility from the absence of hydrolyzable crosslinking reagents, and compatibility with many peptidelabeling chemistries and affinity chromatography for postlabeling cleanup. Polyhistidine tags are also versatile: peptides, proteins, antibody fragments, and oligonucleotides have all been assembled to QDs using polyhistidine tags [36]. Table 2 lists representative examples of polyhistidine-mediated conjugation of peptides onto QDs, noting the wide range of ligand-based QD surface chemistries that are compatible with polyhistidine tags [18, 29, 32, 47, 54–56, 60, 75–82]. Both examples in Fig. 2 also used polyhistidine-tagged peptides.

Hisn-Ni-NTA

Direct Thiol coordination with coordination cysteine residue to QD surface Hisn tags Refer to Table 2

CdSe/ZnS (545)

b

RGDC

CdSeTe/ZnS coated with Vascular cell adhesion amine-terminated molecule-1 binding PEGd (696) peptide PEG2000-NH2QD (705) Thiolated cyclic arginineglycine-aspartic acid (RGD) peptide Heterobifunctional Amine-terminated PEG Thiolated chlorotoxin and crosslinkers (SIAf) QD (525, 655) dendrotoxin, peptidic neurotoxins Click chemistry (Cu(I)Azide-terminated CdSe/ Alkyne-terminated nuclear catalyzed azide-alkyne ZnS QD (620) localization sequence cycloaddition) peptide Click chemistry (thiol-ene) Alkene-terminated Si QD RGDC (440) Carbodiimide coupling Carboxylate-terminated Matrix metalloprotease (EDCg/sulfo-NHSh) PbS/CdS/ZnS (1200) substrate

Heterobifunctional Covalent crosslinker (sulfoattachment to SMCCc) ligand Heterobifunctional crosslinker (GMBSe)

Peak PL wavelength 5-Carboxytetramethylrhodamine c Sulfosuccinimidyl 4-(N-maleimidomethyl)cyclohexane-1-carboxylate d Polyethylene glycol, denoted by molecular weight (MW) when available e 4-Maleimidobutyric acid N-hydroxysuccinimide ester f N-hydroxysuccinimidyl iodoacetate g 1-Ethyl-3-(3-dimethylaminopropyl)carbodiimide h N-hydroxysulfosuccinimide

a

CdSe/ZnS with thiolated H6GG NTA ligands (512)

Biotin-streptavidin

2 interaction Biotinylated p160 (targets breast cancer cells)

Streptavidin-coated CdSe/ZnS (620)

Conjugation method

Category

QD Composition (Colora/ nm) Assembled peptide

Table 1 Representative examples of QD-peptide bioconjugation strategies

In vivo and in vitro imaging of inflamed endothelial tissue NIR tumor imaging

Fluorescence imaging of cancer cells Targeted delivery of QDs to nuclei of live cells Proof-of-concept trypsin detection with Si QDs NIR imaging of tumors

–

–

–

Dy485

[28]

[73]

[42]

[27]

[26]

[25]

Rhodamine Protease sensor for imaging [74] red-X cancerous and healthy cell lines

Methylene blue

Ref

[31] Delivery of antisense oligonucleotide to breast cancer cells Proof-of-concept FRET assay for [72] His6-Ni-NTA

Purpose or application

–

TAMRAb

–

Peptide label

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a

Ligand MW (g mol1)

CdSe/CdS/ZnS (602) CdSe/ZnS (550), CdSe/ CdS/CdZnS/ZnS (615) CdSe/ZnS (625)

Carboxybetaine 300

CL1b, CL2c, CL3d, CL4e CL4

400

300–400

CdSeS/ZnS (524, 624) CdSe/CdS/ZnS (630)

100 400

3000

3100

2600–3800

4200 3100–4300

CdSe/CdS/ZnS (520–650), 1800–4300 CdSeS/ZnS (525–540)

Cysteine Sulfobetaine

3300–4300

CdSe/CdS/ZnS (605), CdSe/CdS/CdZnS/ZnS (625) CdxZn1-xSe/CdyZn1-yS/ ZnS, ZnSe/CdyZn1-yS/ZnS (410–530) 3100–3200

2700

CdSe/ZnS (520, 540)

300

800

1000

2100–2500

CdSe/ZnS (540)

Protease detection

Purpose or application

–

Alexa Fluor 680, Alexa Fluor 647 Alexa Fluor 647 Alexa Fluor 488, 647 dyes Alexa Fluor 647, 680 dyes –

–

[18]

[77] [54]

[55, 56]

[76]

[29]

[32]

[75]

Ref

Intracellular QD uptake for [78] imaging Mapping embryonic chick [79] neurological development

Protease detection

Protease detection Protease detection

Smartphone-based protease detection

Extracellular pH monitoring

Doxorubicin, via Drug delivery cleavable linker C60 Imaging of cell membrane potential

Ru complexes, ferrocene

Peptide + label QD Composition (Color/nm) MW (g mol1) Peptide label

Zwitterionic Glutathione ligands

DHLAPEG750OMe DHLAPEG600NH2

800 PEG DHLA derivatives PEG600-OH 1200 DHLAPEG1000OH 1000 DHLAPEG750OMe

Ligand

Table 2 Representative examples of polyhistidine conjugation of peptides onto QDs with different ligand coatings

182 Katherine D. Krause et al.

1400 1900 3100

3000

ZnxHg1-xSe (670–907) CdSe/ZnS (605) CdSe/ZnS (580)

CdSe/ZnS (510, 550)

6000–10,000 CdSe/ZnS, CdSe/CdS/ZnS 4000–4200 (605)

200

DHLA

Dextran

100

MPAf

b

Dihydrolipoic acid (6,8-dimercaptooctanoic acid) 4-[2-[(6,8-dimercapto-1-oxooctyl)amino]ethyl]-1-piperazinepropanoic acid c N-[2-[(6,8-dimercapto-1-oxooctyl)amino]ethyl]-N-(2-hydroxyethyl)-β-alanine d N-[2-[bis(2-hydroxyethyl)amino]ethyl]-6,8-dimercapto-octanamide e N-(2-carboxyethyl)-N-[2-[(6,8-dimercapto-1-oxooctyl)amino]ethyl]-β-alanine f 3-Mercaptopropionic acid

a

Other ligands

Anionic ligands

Alexa Fluor 647, 680 dyes

– CalciumRubyCl, Ca2+-sensitive indicator dye –

–

Protease detection

[82]

Intracellular QD uptake for [81] imaging

Intracellular QD uptake for [80] imaging Kinase detection [60] Ratiometric Ca2+ detection [47]

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Because the polyhistidine tag coordinates directly to the QD surface, the tag must be able to access this surface through the hydrophilic coating on the QD, as illustrated in Fig. 1c. Polyhistidine tags are therefore not compatible with all pairings of biomolecules and QD surface coatings. For example, bulky ligand or polymer coatings may occlude the QD surface and restrict access for large biomolecules such as globular proteins. Alternative QD coatings have thus been developed to mitigate steric effects, including ligands [90] and amphiphilic polymers [91] that provide interfacial nitrilotriacetic acid (NTA) or carboxylate groups [92, 93] for Ni2+-mediated binding of polyhistidine tags. In addition, helical peptidyl linkers have been useful as coating-penetrating spacers between a polyhistidine tag and a globular protein [94]. Readers are referred to several reviews for more information on different methods for polyhistidine conjugation to QDs [17, 36, 89]. Here, we provide detailed procedures for preparing QD-peptide conjugates through polyhistidine tags, and for characterizing the conjugates via agarose and polyacrylamide gel electrophoresis and FRET. The procedures include methods for confirming peptide assembly, estimating the maximum peptide loading per QD, and determination of binding kinetics.

2 2.1

Materials Reagents

1. Stock solution (1–10 μM) of aqueous QDs with a ZnS shell. Examples of compatible QD surface coatings are listed in Table 2. 2. Peptides with a Hisn tag at one terminus. Some experiments (vide infra) require these peptides to be labeled with a fluorescent dye. Protocols for fluorescently labelingpeptides have been reported elsewhere [95]. 3. Agarose, molecular biology grade, low electroendosmosis. 4. Ultrapure water with specific resistance 18 MΩ cm. Unless otherwise stated, “water” will refer to ultrapure water hereafter. 5. Tris-borate-EDTA (TBE) buffer (1): pH 8.3, 89 mM Tris, 89 mM borate, 2 mM EDTA. Prepare by diluting commercially available 10 TBE buffer in a 1:9 ratio with water. The EDTA can optionally be omitted (i.e., 1 TB buffer; Note 11). This buffer is referred to as running buffer for gels. 6. Borate buffer: pH 9.2, 50–100 mM. 7. Ethanol (EtOH), 95%. 8. Glycerol, biotechnology grade.

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9. 3-(Trimethoxysilyl)propyl methacrylate (TMSM), 98% (see Note 12). 10. Acetic acid, glacial. 11. 37.5:1 Acrylamide:bis-acrylamide solution: 40% w/w in water. This solution can be purchased pre-made or be prepared by dissolving 37.81 g of acrylamide and 2.19 g of bis-acrylamide per 100 mL of water. (Unpolymerized acrylamide is a neurotoxin and must be handled with appropriate precautions.) 12. Tetramethylethylenediamine (TEMED). 13. Ammonium persulfate (APS) solution: 25% w/v in water. This solution should be prepared fresh by dissolving 25 mg in 100 μL of water. 14. Sigmacote siliconizing reagent (Sigma-Aldrich, St. Louis, MO, USA). 15. Fluorescein. 2.2

Equipment

1. Instrument for fluorescence measurements. It should be capable of excitation at 300–450 nm and measurements of emission between 450 and 750 nm. Emission should be measurable with spectral resolution or with multiple color channels. We use an Infinite M1000 Pro multifunction plate reader (Tecan US, Inc., Morrisville, NC, USA). 2. Agarose gel electrophoresis system, including power supply, gel casting unit, tray, tank, and combs. 3. Gel imaging system. For high-quality images, we use a Gel Doc XR+ System (Bio-Rad Laboratories, Inc., Hercules, CA, USA) and UV illumination. Gels can also be imaged at lower resolution using a handheld longwave UV lamp, UV-blocking filter, and a smartphone camera. 4. Borosilicate glass capillaries, 75 mm length, 0.4 mm inner diameter (Drummond Scientific Co., Broomall, PA, USA). 5. Inverted fluorescence microscope (see Note 13). 6. Capillary gel electrophoresis setup that is compatible with the fluorescence microscope and the capillaries. We use a custombuilt clear Plexiglas housing with a magnetic safety interlock in the lid. A schematic of the housing is shown in Fig. 3 [96]. It measures 16.8 cm  8.6 cm  3.0 cm and sits within a recess on our microscope stage. 7. High-voltage (HV) power supply or sequencer. We use an ER230 HV sequencer (eDAQ, Colorado Springs, CO, USA). 8. Glass microscope slide drilled with a 1.3 cm diameter hole (see Note 14) and patterned buffer reservoirs (see Note 15).

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Fig. 3 (a) Schematic of the custom-built PAGE apparatus. The Pt wires are attached to a high-voltage sequencer. (b) Photograph of the PAGE apparatus mounted on a microscope stage. (c) Schematic of the optical path. The PAGE apparatus is simplified to show only the capillary and the fluorescent materials migrating through its interior. (Reproduced with permission from Ref. (96). Copyright 2018 Royal Society of Chemistry)

9. Portable charge-coupled device (CCD) spectrometer (see Note 16) and compatible optical fiber connection to the fluorescence microscope. We use a GreenWave 16 VIS-50 (StellarNet, Tampa, FL, USA) with a round-to-linear fiber bundle of 7  200 μm diameter core fibers (Thorlabs Inc., Newton, NJ, USA). Light output from a camera port on the microscope is coupled into the fiber bundle that leads to the CCD spectrometer (see Note 17).

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10. Software for measuring PL spectra (or intensity) as a function of time. We use software custom-written in LabVIEW (National Instruments, Austin, TX, USA) with our microscope (see Note 18) and the normal operating software for our plate reader. 11. Plasma Cleaner (Harrick Plasma PDC-32G; Ithaca, NY, USA).

3

Methods Throughout this section, a QD with an average of N conjugated copies (see Note 10) of a peptide sequence is denoted by QD-[pep]N or, if dye-labeled, QD-[pep(dye)]N. PL measurements at particular wavelengths (or with particular filters) that correspond to predominantly QD emission and predominantly dye emission are denoted by λQD and λDye, respectively.

3.1 Peptide Assembly

The binding of Hisn-tagged peptides to QDs is robust and compatible with wide ranges of buffers with pH  7 (see Note 19), QD concentrations (50 nM–10 μM), and overall scales (2–200 pmol). The following steps are to assemble N copies of a Hisn-tagged peptide to QDs to yield QD-[pep]Nconjugates (or QD-[pep(dye)]Nconjugates if assembling a dye-labeled peptide). The protocol is adaptable to the assembly of multiple peptide sequences by adding the desired equivalents of each of those sequences at step 2. 1. Determine the desired volume and concentration for sample preparation (see Note 20). 2. Add N equivalents microcentrifuge tube.

of

Hisn-peptide

to

a

1.7-mL

3. Dilute with buffer (see Note 21). 4. Add the required amount of QD stock solution. 5. Mix immediately and let stand at room temperature for 1 h (see Note 22) and protect from light (see Note 23). 6. Store the QD-[pep]N conjugates in the dark at 4  C until needed. 3.2 Agarose Gel Electrophoresis 3.2.1 Confirmation of Peptide Binding

Agarose gel electrophoresis is a useful method for confirming the binding of Hisn-tagged peptides to QDs. Conjugated peptides alter the hydrodynamic size and net charge of a QD. The magnitude of the resulting change in the electrophoretic mobility of the QDs depends on the amino acid sequence and number of peptides per QD (see Notes 24 and 25). 1. Prepare a 1% w/v agarose gel using running buffer. 2. Prepare or obtain QD-[pep]N sample(s) for analysis (see Subheading 3.1).

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3. Prepare a QD-only control sample using the same batch of QDs that was used to prepare the QD-[pep]Nconjugates (see Note 26). 4. If the peptide is dye-labeled, prepare a pep(dye)-only control sample. 5. Dilute 0.6–6.0 pmol of each QD-containing sample to 10 μL in buffer (see Note 27). Each sample should have the same final concentration. 6. If analyzing a pep(dye)-only sample, dilute the sample to 10 μL and a final concentration N-fold higher than that of the QD-only sample. 7. To each sample, add 2 μL of a 60% v/v glycerol (aq) and mix (see Note 28). 8. Using a micropipette, transfer 10 μL of each sample into a gel well. Each sample should contain 0.5–5.0 pmol of QDs and 10% v/v glycerol. 9. Run the gel at a field strength of 6.67 V cm1 for 20–40 min (see Note 29). This field strength is equivalent to 100 V for electrodes 15 cm apart. 10. Image the gel using photoluminescence. This step should be done as soon as possible after the run is complete to minimize diffusion of the bands. Examples of gels that confirm successful assembly of peptides to QDs through changes in electrophoretic mobility are shown in Fig. 4 (see Note 30) [18, 63, 82]. In some cases, the above method can be adapted to confirm the assembly of two different peptide sequences by running suitable control samples (see Notes 31 and 32). Another strategy in this context is to determine the peptide loading capacity of a QD and assume (usually validly) that the multiple peptide sequences will successfully assemble as long as the total equivalents are less than the maximum peptide loading. 3.2.2 Estimation of Peptide Loading Capacity

The agarose gel electrophoresis method of the preceding section can be extended to estimate the maximum number of Hisn-tagged peptides that can assemble per QD. 1. Obtain dye-labeled peptides, pep(dye). The dye should have fluorescence that is visible using the gel imaging system, and, ideally, a different color than the QD PL (see Note 33). 2. Prepare a series of QD-[pep(dye)]Nconjugates (see Subheading 3.1). These samples should have increasing values of N and include multiple conjugates where N exceeds either the desired loading or the anticipated maximum loading (see Note 34). The sample concentrations should be sufficiently high that the dye fluorescence can be imaged directly.

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Fig. 4 Examples of agarose gels showing electrophoretic mobility shifts induced by successful peptide assembly. Red and blue arrows indicate the positions of QD and QD-peptide conjugate bands, respectively. Dashed lines indicate sample well positions. Equivalents of peptide are denoted for each gel. (a) The assembly of increasing equivalents of His6-tagged peptide induces a progressive decrease in the anodic mobility of anionic QD605. This trend is not observed with peptide without a Hisn tag. (Reproduced with permission from Ref. (18). Copyright 2018 American Chemical Society). (b) The assembly of increasing equivalents of anionic His6-tagged, dye-labeled peptide to uncharged QD645 induces a progressive increase in anodic mobility. QD PL is also progressively decreased by the addition of the Alexa Fluor 680 (A680) dye-labeled peptide. (Reproduced with permission from Ref. (82). Copyright 2020 American Chemical Society). (c) The assembly of increasing equivalents of His8-tagged, Cy5 dye-labeled peptide to anionic QD550 induces a decrease in anodic mobility. At 60 equivalents of peptide, a new band is seen for unbound peptide (green arrow and dotted outline). (Reproduced with permission from Ref. (63). Copyright 2010 Wiley-VCH Verlag GmbH)

3. Prepare a 1% w/v agarose gel using running buffer. 4. Prepare the samples of QD-[pep(dye)]Nconjugates for electrophoresis as described in Subheading 3.2.1. 5. Prepare a QD-only control sample at the same volume and concentration as the QD-[pep(dye)]N samples. 6. Prepare a pep(dye) control sample at the same volume as the QD-[pep(dye)]N samples and a concentration equivalent to that of the pep(dye) in one of the intermediate QD-[pep (dye)]N samples. 7. Run the gel as described in Subheading 3.2.1. 8. Image the gel as soon as possible after the run is complete. One or more images should be acquired under conditions that permit independent visualization of both the QDs and dye-labeled peptides. 9. Analyze the gel image(s). Below the maximum loading, the band of dye fluorescence will overlap with the band of QD

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PL. Above the maximum loading, a new band of dye fluorescence will appear (see Note 35). This new band will be aligned with the band for the pep(dye) control sample and will not be co-localized with the QD band (see Note 36). In certain cases, the trend in the electrophoretic mobility of the QD-[pep (dye)]N samples with increasing N may also be useful for estimating the maximum loading, but may also be unreliable (see Note 37). 10. If the gel was inconclusive, another gel should be run with either higher values of N or a more narrow range of N for QD-[pep(dye)]Nconjugates. Figure 5 shows an example of a gel used for estimating the maximum peptide loading of a sample of QDs. For 60 or fewer peptides per QD, a single band is observed and corresponds to QD-[pep(dye)]N. The color of PL from this band changes from orange to yellow-orange at 60 peptides. For 90 and 120 peptides, the PL from the QD-[pep(dye)]N band is more distinctly yellow and a new band with green PL appears. This new band corresponds to unbound pep(dye) and green is the color of dye PL. (The observed yellow color of the QD-[pep(dye)]N arises from the superposition of orange QD PL and green dye PL.) The estimated average maximum loading for these peptides and QDs is thus greater than 60 and less than 90 peptides per QD.

Fig. 5 Example of the electrophoretic determination of peptide loading capacity using a 1% w/v agarose gel. The green, blue, and red arrows indicate the electrophoretic mobilities of the pep(dye), QD605-[pep(A488)]N conjugates, and QD, respectively. The QDs are coated with dihydrolipoic acid (DHLA) ligands and are anionic. In the image, QD PL appears orange and Alexa Fluor 488 (A488) dye fluorescence appears green. Some streaking and a band for excess peptide is observed in the lanes with 90 and 120 equivalents of pep(dye), but not in the lane with 60 equivalents of pep(dye). The maximum peptide loading capacity is therefore at least 60 peptides/QD. (Reproduced with permission from Ref. (18). Copyright 2018 American Chemical Society)

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3.3 Capillary Polyacrylamide Gel Electrophoresis

Poly(acrylamide) gel electrophoresis (PAGE) also enables confirmation of binding of Hisn-tagged peptides to QDs on the basis of changes in electrophoretic mobility. Whereas slab agarose gel electrophoresis is widely used for this purpose, PAGE is not. A reason is that the pores of a slab poly(acrylamide) gel (PAG), though more uniform than those of agarose, tend to be smaller than ideal for QDs and yield poor quality separations. This limitation was overcome by implementing a “stubby” capillary format for PAGE (see Note 38) [96]. The format is a hybrid of conventional capillary and slab PAGE formats and offers several advantages, including minimal sample consumption, high field strength at low voltages, short runtimes, and high resolution. The QD must have a significant net charge to use this method.

3.3.1 Preparing the Polyacrylamide Gel-Filled Capillaries

This section describes a method of preparing PAG-filled capillaries starting from commercially available hollow glass capillaries. 1. Place 10 capillaries in a 50-mL centrifuge tube containing ~45 mL EtOH. Sonicate the tube for 2 h to clean the capillaries. 2. Drain the EtOH from the capillaries using a paper towel and dry the capillaries with clean compressed air. 3. In a 15-mL centrifuge tube, dilute 200 μL TMSM and 100 μL glacial acetic acid with EtOH to a final volume of 10 mL. This solution should be prepared fresh. 4. Plasma clean the capillaries. We use air plasma at 18 W RF power for 45 s. 5. Immerse the capillaries in the TMSM/acetic acid solution immediately after completion of the plasma cleaning. Let stand at room temperature for at least 4 h. 6. Drain and dry the capillaries as described in step 2. 7. Place the dried capillaries in a ~70  C oven for at least 1 h. 8. For a 3.0% w/w gel (see Note 39), dilute 113 μL of acrylamide: bis-acrylamide (37.5:1, 40% w/w in water) with 1381 μL borate buffer (100 mM, pH 9.2; Note 40) in a 1.7-mL microcentrifuge tube. Add 1.5 μL of TEMED and 4.5 μL of freshly prepared APS solution (25% w/v). Mix vigorously. 9. Create three holes in the lid of the microcentrifuge tube (see Note 41). Immediately insert the capillaries through the holes and immerse their ends in the acrylamide:bis-acrylamide solution to the center of the depth of the solution. The capillaries will fill with the solution through capillary action. 10. Leave the capillaries at room temperature for 2 h to allow the gel to polymerize.

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11. Remove the filled capillaries from the microcentrifuge tube and wipe their outer surface to clean off any adhered gel. 12. Soak the PAG-filled capillaries in borate buffer (100 mM, pH 9.2) for at least 20 min before conducting experiments (see Note 42). The capillaries can be prepared in batches and stored in buffer at room temperature for a month. 13. Truncate the PAG-filled capillary to ~5.5 cm prior to experiments (see Note 43). 3.3.2 Confirmation of Peptide Binding

This section describes a protocol for capillary PAGE with real-time detection. Our apparatus is shown in Fig. 3. Fluorescein is used as an internal standard against which the migration times of the QDs are standardized (see Notes 44 and 45). This protocol is for QDs with a net negative charge. 1. Set up the microscope for time-based measurements of fluorescence near the anode end of the capillary (see Notes 46 and 47). 2. Prepare a primary stock solution of 1–25 mg fluorescein in 10 mL borate buffer (100 mM, pH 9.2). Dilute 50 μL of this stock to a final volume of 1.5 mL to produce a 10–250 μM secondary fluorescein stock. 3. Prepare as many samples of QD-[pep(dye)]Nconjugates as of interest (see Subheading 3.1). If necessary, dilute each sample in buffer to a volume of 45 μL and QD concentration of 0.1–0.6 μM. 4. Prepare a QD-only control sample at the same concentration as the QD-[pep(dye)]Nconjugates (see Note 26). 5. Spike 45 μL of each sample with 5 μL of the secondary fluorescein stock. The final fluorescein concentration should be 1–25 μM (see Note 48) and the QD concentration should be 0.1–0.5 μM. 6. Add 250 μL of borate buffer (100 mM, pH 9.2; Note 49) to the anode reservoir and 50 μL of sample to the cathode reservoir (see Note 46). 7. Electrokinetically inject QDs into the capillary by applying a voltage of 300 V across the capillary for 5 s (see Note 50). 8. Remove the residual sample from the cathode reservoir and wipe the exterior surface of the capillary to remove any residual sample. 9. Add 350 μL of borate buffer (100 mM, pH 9.2; Note 49) to the cathode reservoir and remount the capillary between the anode and cathode reservoirs.

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10. Adjust the position of the capillary (if necessary) and set the objective lens to focus as close to the center of the interior of the capillary as possible (see Note 51). 11. Run the sample at 300 V for a field strength of ~55 V/cm. Most gels should run for a minimum of 10 min and a maximum of 30 min (see Note 52). 12. Turn off the high voltage after the allotted time. 13. Repeat the above steps for additional samples (see Note 53). 14. The desired form of the data are electropherograms that plot PL intensity versus migration time. If necessary, adjust the migration time (measured at the electropherogram peak) of each QD by subtracting any differences in the migration time of the fluorescein internal standard. As shown in Fig. 6, QDs conjugated with increasing equivalents of peptide will have increasingly longer migration times. 3.4 FRET-Based Characterization

Given its short range (see Note 54), FRET is a useful means of confirming the assembly of peptides to QDs when the peptides have been labeled with a suitable fluorescent dye (see Note 55). FRET can also be used to measure the kinetics of assembly of Hisntagged peptides and estimate the maximum peptide loading

Fig. 6 Example of capillary PAGE data for QDs and QD-peptide conjugates. PL electropherograms for QD600-[pep]Nconjugates with increasing N and a 3.0% w/w PAG capillary. The migration time increases with the number of peptides assembled. Separate peaks are observed for the QD and fluorescein internal standard. Adapted with permission from Ref. (96). Copyright 2018 Royal Society of Chemistry

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capacity of a QD. The results obtained with dye-labeled peptides are often a good proxy for the same metrics with unlabeled peptides. 3.4.1 Confirmation of Peptide Binding

1. Prepare a QD-[pep(dye)]N conjugate as described in Subheading 3.1. 2. Prepare QD-only and pep(dye)-only control samples at the same concentrations and volumes. 3. Measure the PL spectra (see Note 56) of the samples using an excitation wavelength that minimizes direct excitation of the dye (see Note 57). 4. If the QD PL intensity of the QD-[pep(dye)]N sample decreased relative to the QD-only sample and the dye PL intensity of the QD-[pep(dye)]N sample increased relative to the pep(dye)-only sample, then the result is consistent with FRET and successful assembly (see Notes 58–60). Figure 7a shows examples of PL emission spectra for the assembly of increasing equivalents of dye-labeled peptide per QD. 5. If the data was not consistent with the criteria in step 4, then an excitation scan is warranted. For each sample and control, collect excitation spectra. The emission wavelength should be one at which the dye emits but the QD does not (see Note 61). The scan should span a range of excitation wavelengths across which the QD absorbs but the dye does not.

Fig. 7 Examples of PL spectra for FRET-based confirmation of binding between N equivalents of His6-tagged, Alexa Fluor 647 (A647) dye-labeled peptides and QD635. (a) Emission spectra (λexc ¼ 405 nm) show a progressive decrease in QD PL and increase in A647 PL with increasing N for QD635-[pep(A647)]Nconjugates. Directly excited dye PL from pep(A647)-only control sample (12 equivalents) is negligible. (b) The excitation spectrum (λem ¼ 670 nm) for a pep(A647)-only control sample show that there is minimal direct excitation of dye PL at wavelengths shorter than ~525 nm. In contrast, dye PL is excited by these wavelengths with QD635[pep(A647)]N samples, indicative of FRET from excitation of the QD

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6. Compare the excitation spectra between the samples. Figure 7b shows examples of PL excitation spectra for the assembly of increasing equivalents of dye-labeled peptide per QD. The indicators of assembly are that the excitation spectrum looks like a combination of the individual excitation (or absorbance) spectra for the QD and dye, and that the QD excitation feature for the QD-[pep(dye)]N samples is more intense than for the QD-only sample and scales with N (see Note 60). In some cases, FRET can be used to directly confirm the assembly of two or more dye-labeled peptide sequences to a QD (see Notes 62 and 63). 3.4.2 Assessment of Assembly Kinetics

In our experience, the kinetics of Hisn tag assembly on QDs is a function of the QD surface chemistry. An assembly time of 30 min is ample for some surface chemistries, whereas up to 120 min may be prudent for other surface chemistries. The following steps may be used to determine the time required for the assembly of dye-labeled peptides to a QD. The assay monitors the QD and dye PL intensity as a function of time after addition of the dye-labeled peptide to the QD sample. 1. Prepare two samples of QDs at twice the desired final QD-[pep (dye)]N concentration in the desired buffer. We typically use 2–5 pmol QD per sample with a concentration of 50–100 nM (see Notes 20 and 21). The QD sample will be diluted to the desired concentration upon addition of an equal volume of either buffer or dye-labeled peptide solution. 2. Prepare two pep(dye) samples at twice the desired final dye-labeled peptide concentration. These samples should be equal in volume and N-fold more concentrated relative to the QD samples from step 1. 3. Set up a plate reader for fluorescence measurements at regular time intervals, usually between 30 s and 2 min over a duration of 1 h (see Note 64). Select the excitation wavelength to minimize direct excitation of the dye-labeled peptide. The PL intensity at λQD and λDye should be measured at each time point. 4. Dilute one of the two QD samples with an equal volume of the desired buffer. Transfer to a sample well in a microtiter plate. 5. Dilute one of the two pep(dye) samples with an equal volume of the desired buffer. Transfer to a sample well in a microtiter plate. 6. To the remaining undiluted QD sample, add the remaining undiluted pep(dye) sample. Transfer to a sample well in a microtiter plate and immediately begin the kinetic assay.

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7. For the QD-[pep(dye)]N sample data, calculate the ratio of the dye and QD PL intensities as a function of time, as described in Eq. 1, where IQD-dye(λ,t) is the PL intensity of the QD-peptide conjugate at wavelength λ and time t. If the direct excitation and spectral crosstalk corrections described in Notes 58 and 59 were required, these corrections should be applied before calculating the PL ratio.
I QDdye λDye , t
Sample PL ratioðt Þ ¼ ð1Þ I QDdye λQD , t 8. For the control samples data, calculate the ratio of the separate dye and QD PL intensities as a function of time, as per Eq. 2:
I Dye λDye , t
ð2Þ Control PL ratioðt Þ ¼ I QD λQD , t where IDye(λ,t) and IQD(λ,t) are the PL intensity of the dye-labeled peptide-only and QD-only control samples at wavelength λ and time t. 9. Plot both the sample and the control PL ratios as a function of time. As the peptides assemble to the QD surface, FRET causes the dye emission to increase and the QD emission to decrease, yielding a net increase in the sample PL ratio. The time at which the sample ratio plateaus marks the completion of the assembly process (see Note 65). Example data are shown in Fig. 8 for equilibrium binding within ~10 min.

Fig. 8 Example of binding kinetics for the assembly of 8 equivalents of A647labeled peptide on glutathione-coated QD605. The increase and subsequent plateau in the dye/QD PL ratio for the QD-[pep(A647)]8 sample indicate that binding was complete after ~10 min

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The next two sections provide protocols to determine the maximum peptide loading capacity of a QD. Both protocols assume that FRET will approach 100% efficiency prior to the QD reaching its maximum capacity for assembled peptides, which is usually the case (see Note 66). As such, a combination of dye-labeled and unlabeled peptides is used (see Notes 67 and 68), assembled to the QD either simultaneously (see Subheading 3.4.3) or sequentially (see Subheading 3.4.4). The use of unlabeled peptides enables probing of large values of assembled peptides without FRET inducing nearcomplete quenching of the QD PL. We denote a conjugate with M equivalents of unlabeled peptide and N equivalents of labeled peptide as QD-[pep]M[pep(dye)]N. The simultaneous addition protocol addresses the need to avoid 100% FRET efficiency by diluting dye-labeled peptide with unlabeled peptide. That is, we leverage QD-[pep]M[pep(dye)]Nconjugates where only one of every five or ten of the peptides is labeled (see Note 69). The maximum loading can thereby be reached and identified prior to saturation of FRET. The assumption in this method is that the unlabeled peptides have minimal impact on FRET between the QDs and dye-labeled peptides (see Notes 70 and 71). When the maximum peptide loading is reached, no further dye-labeled peptides assemble to the QDs and the indicators of FRET (e.g., QD PL intensity, dye PL intensity, dye/QD PL ratio) should approach plateaus or show other deviations in their overall trend with increasing M + N (see Note 72). 1. Prepare a stock solution of pep(dye). 2. Prepare an additional stock solution containing a mixture of pep(dye) and unlabeled peptide at a ratio of 1:5 (i.e., 20% labeled peptide). 3. Prepare an additional stock solution containing a mixture of pep(dye) and unlabeled peptide at a ratio of 1:10 (i.e., 10% labeled peptide). 4. Using the stock solution of pep(dye), prepare a series of QD-[pep(dye)]Nconjugates with increasing values of N as described in Subheading 3.1. The values of N should span the range over which there is still significant QD PL in the spectrum of the assembled conjugate. 5. Using the stock solution of 20% pep(dye) and unlabeled peptide, prepare a series of QD-[pep]M[pep(dye)]Nconjugates with increasing values of total assembled peptide (i.e., M + N). The series should include multiple conjugates in which the value of M + N exceeds the anticipated maximum loading of the QD. The values of N for the QD-[pep]M[pep (dye)]Nconjugates should span the same values of N as the conjugates prepared in step 4.

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6. Repeat step 5 with the stock solution of 10% pep(dye) and unlabeled peptide. 7. Prepare a QD-only control sample at the same concentration and volume as each of the QD-peptide conjugate samples. 8. If it is not known from prior experiments that the direct excitation of dye fluorescence is negligible in measurements, then pep(dye)-only control samples should be prepared at the same concentrations as in step 4 (see Note 73). 9. Measure a PL emission spectrum (see Note 56) for each sample using an excitation wavelength that minimizes direct excitation of the dye. 10. Determine the IQD and IDye values from each sample as described in Subheading 3.4.1, including crosstalk and direct excitation corrections (see Notes 58 and 59) if necessary. 11. For each conjugate series, plot the QD PL intensity, the dye PL intensity, and the dye-to-QD PL ratio as a function of N. Figure 9a shows example data for conjugates prepared using 100%, 20%, and 10% dye-labeled peptide. 12. For each set of QD-[pep]M[pep(dye)]N samples, identify the value of N at which the QD PL intensity, dye PL intensity, or dye-to-QD PL ratio deviates from the qualitative trend observed with the QD-[pep(dye)]Nconjugates. The value of M + N at which the deviation occurs corresponds (approximately) to the maximum peptide loading capacity of the QD. For example, the plots of 10% and 20% pep(dye) in Figs. 9b–d show inflection points near N ¼ 5 and N ¼ 12, respectively, that are not observed with 100% pep(dye). The data thus suggests an average peptide loading capacity of ~50–60 peptides per QD (see Note 74). 3.4.4 Estimation of Peptide Loading Capacity via Sequential Addition

In this protocol, two-step assembly of QD-[pep]M[pep(dye)]Nconjugates is used to overcome the obstacle of FRET efficiencies that approach 100%. Increasing equivalents, M, of unlabeled peptide are added to the QD in a first step, given time to bind, and a constant N equivalents of dye-labeled peptide are added in a second step (see Notes 67 and 68). As M reaches the maximum loading for the QDs, the dye-labeled peptides will no longer be able to assemble. The QD PL should be a plateau at low intensity for values of M well below the maximum loading, then transition to a plateau at high intensity for values of M above the maximum loading. The inverse trend should be observed for the dye PL. 1. Prepare a series of at least five QD-[pep]Mconjugates of increasing M, as described in Subheading 3.1. The range of M should include multiple values that exceed the anticipated maximum peptide loading capacity of the QDs (see Note 34).

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Fig. 9 Example of FRET-based estimation of the peptide loading capacity of QDs using Method 1 (simultaneous addition). (a) PL emission spectra of QD-[pep]M[pep(A647)]Nconjugates with (i) 100%, (ii) 20%, and (iii) 10% labeled peptide. The spectra in (ii) and (iii) are denoted by “N / (M+N )” to indicate both the equivalents of labeled peptides and the total equivalents of peptide. The PL spectra for the peptide-only controls (mixtures of labeled and unlabeled peptide without QD, and denoted only by the equivalents of dye-labeled peptide) are indistinguishable from the baseline. Corresponding plots of (b) normalized QD PL intensity, (c) A647 PL intensity, and (d) A647/QD PL ratio as functions of N. Fit curves are intended to guide the eye and are not based on FRET theory. The inflection points at N  5 for the data with 10% labeled peptide and at N  12 for the data with 20% labeled peptide indicate that the surface is saturated with approximately 50–60 peptides

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2. Prepare two QD-only samples at the same concentration and volume as step 1. 3. Prepare a pep(dye)-only sample at the same volume and pep (dye) concentration as the QD-[pep]M[pep(dye)]N samples. 4. Let the QD-[pep]M samples stand for 1–2 h so that all of the unlabeled peptides have had ample time to assemble (see Note 22). 5. Add N equivalents of dye-labeled peptide to all samples except one of the two QD-only samples. 6. Let the samples stand for 1–2 h so that all of the pep(dye) have had ample time to assemble. 7. Measure the PL emission spectrum (see Note 56) of each sample using an excitation wavelength that minimizes direct excitation of the dye. 8. Determine the IQD and IDye values from each sample as described in Subheading 3.4.1, including crosstalk and direct excitation corrections as described in Notes 58 and 59 if necessary. 9. Plot the IQD and IDye values as functions of M. Figure 10 is an example of such data for the same QDs and peptides as those used in Subheading 3.4.3 and Fig. 9. 10. Identify the approximate value of M at which the QD and dye PL intensities transition between high and low plateaus. The maximum peptide loading capacity is estimated to be the sum of M and N at this point (see Note 75). In Fig. 10, the onset of the increase in QD PL and decrease in dye PL suggests that the total number of peptides per QD that can assemble without significant excess remaining in bulk solution is ~40–50 (M ¼ 30–40, N ¼ 10). However, the average maximum loading for the population of QDs, as indicated by the halfway point in the transition, is closer to ~60–70 peptides per QD (M ¼ 50–60, N ¼ 10). 11. If there is no clear transition point for the QD or dye PL intensities, the experiments should be repeated with larger values of M because maximum loading was likely not attained. At maximum loading, the QD PL intensity of the sample with the largest value of M should be roughly equal (usually within 10%) to that of the QD-only control sample.

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Fig. 10 Example of FRET-based estimation of the peptide loading capacity of QDs using Method 2 (sequential addition). (a) PL emission spectra showing the effect of assembling increasing equivalents of unlabeled peptide on the QD prior to assembling 10 equivalents of labeled peptide. Labels in the legend are of the form (M, 10); the QD-only sample is denoted (0,0). (b) Corresponding plots of QD and dye PL intensity as a function of M. Fit curves are intended to guide the eye and are not based on FRET theory. The horizontal dashed line corresponds to the QD PL intensity for the QD-only sample. The value of M + N at which the QD and dye PL intensities deviate from their initial plateaus and reach inflection points represent threshold (M ¼ 30–40, N ¼ 10) and average loading capacities (M ¼ 50–60, N ¼ 10), respectively. (c) Electrophoretic confirmation of the successful assembly of both labeled and unlabeled peptides, indicated by the decrease in mobility between (0,0) and (0,10) and the progressive decrease in mobility of the (10  M  30,10) samples, respectively

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Notes 1. Photoluminescence (PL) is defined as the emission of a photon from a photoexcited species. The term is typically used to describe the emission from a photoexcited QD rather than fluorescence (which, like phosphorescence, is a type of photoluminescence). For the purpose of this chapter, readers should feel free to conceptually interchange the terms PL and fluorescence. 2. The cytotoxicity of CdX QDs and their core/shell analogs depends on several factors, including the QD surface chemistry, the presence or absence of a shell, the QD diameter, and the route and duration of exposure [23, 24, 97]. In practice, QDs with high-quality shells and surface chemistry are no more toxic than molecular fluorophores [24]. Cd-free QDs (e.g., InP, CuInS2, Ag2S) and their core/shell analogs are also under development; however, these materials do not yet have PL properties on par with those of CdX and PbX QDs [98, 99]. 3. FRET is non-radiative, through-space energy transfer from an electronically excited donor to a ground-state acceptor. The acceptor effectively takes the excitation energy from the donor, resulting in quenching of donor fluorescence and, if fluorescent, emission from the acceptor. In order for FRET to occur, the emission spectrum of the donor and the absorption spectrum of the acceptor must overlap, and the donor and acceptor must typically be