Nanopore Technology: Methods and Protocols [1st ed.] 9781071608050, 9781071608067

This detailed collection explores techniques involved in the main strategies of nanopore sensing, such as translocation,

521 26 7MB

English Pages X, 231 [231] Year 2021

Report DMCA / Copyright


Polecaj historie

Nanopore Technology: Methods and Protocols [1st ed.]
 9781071608050, 9781071608067

Table of contents :
Front Matter ....Pages i-x
Front Matter ....Pages 1-1
Preparation of Fragaceatoxin C (FraC) Nanopores (Natalie Lisa Mutter, Gang Huang, Nieck Jordy van der Heide, Florian Leonardus Rudolfus Lucas, Nicole Stéphanie Galenkamp, Giovanni Maglia et al.)....Pages 3-10
Preparation of Cytolysin A (ClyA) Nanopores (Nicole Stéphanie Galenkamp, Veerle Van Meervelt, Natalie Lisa Mutter, Nieck Jordy van der Heide, Carsten Wloka, Giovanni Maglia)....Pages 11-18
Building Synthetic Transmembrane Peptide Pores (Kozhinjampara R. Mahendran)....Pages 19-32
Design and Assembly of Membrane-Spanning DNA Nanopores (Kerstin Göpfrich, Alexander Ohmann, Ulrich F. Keyser)....Pages 33-48
Front Matter ....Pages 49-49
Determining the Orientation of Porins in Planar Lipid Bilayers (Sandra A. Ionescu, Sejeong Lee, Hagan Bayley)....Pages 51-62
Revelation of Function and Inhibition of Wza Through Single-Channel Studies (Lingbing Kong)....Pages 63-76
Protein Analyte Sensing with an Outer Membrane Protein G (OmpG) Nanopore (Monifa A. V. Fahie, Bib Yang, Christina M. Chisholm, Min Chen)....Pages 77-94
Nanopore Enzymology to Study Protein Kinases and Their Inhibition by Small Molecules (Leon Harrington, Leila T. Alexander, Stefan Knapp, Hagan Bayley)....Pages 95-114
A Selective Activity-Based Approach for Analysis of Enzymes with an OmpG Nanopore (Monifa A. V. Fahie, Bach Pham, Fanjun Li, Min Chen)....Pages 115-133
Oligonucleotide-Directed Protein Threading Through a Rigid Nanopore (Garbiñe Celaya, David Rodriguez-Larrea)....Pages 135-144
Unfolding and Translocation of Proteins Through an Alpha-Hemolysin Nanopore by ClpXP (Jeff Nivala, Logan Mulroney, Qing Luan, Robin Abu-Shumays, Mark Akeson)....Pages 145-155
Front Matter ....Pages 157-157
Simulation of pH-Dependent, Loop-Based Membrane Protein Gating Using Pretzel (Alan Perez-Rathke, Monifa A. V. Fahie, Christina M. Chisholm, Min Chen, Jie Liang)....Pages 159-169
Free Energy Minimization for Vesicle Translocation Through a Narrow Pore (Hamid R. Shojaei, Ahad Khaleghi Ardabili, Murrugappan Muthukumar)....Pages 171-183
Front Matter ....Pages 185-185
Single Ion-Channel Analysis in Droplet Interface Bilayer (Arash Manafirad)....Pages 187-195
Continuous and Rapid Solution Exchange in a Lipid Bilayer Perfusion System Based on Droplet-Interface Bilayer (En-Hsin Lee)....Pages 197-211
Protein Transport Studied by a Model Asymmetric Membrane Army Arranged in a Dimple Chip (Xin Li, Min Chen)....Pages 213-225
Back Matter ....Pages 227-231

Citation preview

Methods in Molecular Biology 2186

Monifa A.V. Fahie Editor

Nanopore Technology Methods and Protocols




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

For further volumes:

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.

Nanopore Technology Methods and Protocols

Edited by

Monifa A. V. Fahie Molecular and Cellular Biology Program, University of Massachusetts Amherst, Amherst, MA, USA

Editor Monifa A. V. Fahie Molecular and Cellular Biology Program University of Massachusetts Amherst Amherst, MA, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-0805-0 ISBN 978-1-0716-0806-7 (eBook) © Springer Science+Business Media, LLC, part of Springer Nature 2021 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 The field of nanopore technology for single molecule sensing has garnered much attention with its robust application in DNA sequencing. Much effort over the last two decades in nanopore design and computational analysis has made nanopore-based DNA sequencing a real-world (and outer space) technique, not only used in the confines of a research laboratory. Nanopore sensing, however, is also used for the analysis of other biological molecules such as RNA and proteins as well as chemicals such as metabolites, toxins, or drugs. In recent years, interest in protein analysis and peptide sequencing has gained traction in and among the members of the nanopore technology field. However, unlike DNA and RNA sequencing, precise and accurate protein analysis has several obstacles. Therefore, this volume primarily focuses on a few of the single molecule methods and nanopores developed for the specific and selective characterization of protein analytes. Nanopores are nanometer-sized holes. In nature, small holes exist as membrane channels that act as gatekeepers, allowing or disallowing molecules to enter or exit the cell. Membrane channels exist in all forms of life. It is here where the field of nanopore technology gained inspiration from as far back as in the 1980s, where the intrinsic activity of acetylcholine receptors were being studied or the nonspecific activity of voltagedependent anion channels (VDAC) against synthetic molecules were studied in model lipid bilayers [1, 2]. Not only are nanopores based on protein channels but also solid-state materials and synthetic DNA origami. Solid-state nanopores, although not represented in this book, are a class of nanopores that have shown great promise in protein analysis [3–6]. The advantage of synthetic nanopores is the option of size tunability while the advantage of purified protein nanopores is consistent and precise pore characteristics such as diameter and asymmetric charge distribution. Hybrid nanopores combine both biological and synthetic nanopores and are a developing technique that can push the boundaries of protein analysis by single nanopore sensing technology. Nanopore sensing is performed in either one of three main strategies. Firstly, early nanopore research focused heavily on the passage of analytes through the nanopore’s lumen, a process called translocation. Molecules upon entering the nanopore lumen would displace water and, therefore, block ion movement through the pore, resulting in a measurable decrease in ionic current. The size of the current blockages is congruent with the molecular weight or size of the translocating molecule. Currently, this detection method is primarily used by solid-state nanopores but also used by protein nanopores, as exampled in Chapters 3–6, 10, 11, and 13–15. In recent years, other methods of analyte detection by protein nanopores have become popular. For example, analyte trapping, which can be considered as incomplete translocation, has only been successfully performed with protein nanopores with asymmetric lumen characteristics and constriction sites significantly smaller than the rest of the lumen, i.e., goblet-shaped nanopores. One example, the cytolysin A (ClyA) nanopore has been used to trap protein analytes up to ~40 kDa in size [7, 8]. The protein analytes’ behavior inside the ClyA nanopore can give information about its compactness or rigidity and can also report on conformational changes due to ligand–protein interactions. Finally, protein nanopores can also detect analytes that do not enter its lumen but those that interact with an external binding site that is either intrinsic to the nanopore or that




which has been engineered. Chapters 7–9 give detailed methodologies for this type of nanopore detection system. This book, with its focus on nanopore technology and biomolecule characterization, will hold the interest of the biophysicists, biochemists, bioengineers, and molecular biologists among us. This book’s contributions come from a collective of 39 scientists from all over the world, working diligently in this growing field of nanopore sensing application. This book includes 16 chapters which are grouped into four parts: Part I consists of four chapters which lay the framework for the foundation of nanopore technology: nanopore design and nanopore production. Part II consists of seven chapters discussing various biological nanopores, nanopore engineering, and their uses in single molecule sensing. In particular, the sensing of proteins and one example of sugar sensing are outlined. Part III consists of two chapters outlining computational methods to study intrinsic nanopore behavior as well as the formulations for characterizing the specific translocation activity of a vesicle particle through a nanopore. Part IV consists of three chapters specifically detailing the use of the technique droplet interface bilayer (DIB) in nanopore and membrane biophysical studies. The editor wishes to thank all the contributors for their dedication and patience during the creation of this book. Amherst, MA, USA

Monifa A. V. Fahie

References 1. Suarez-Isla BA, Wan K, Lindstrom J, Montal M (1983) Single-channel recordings from purified acetylcholine receptors reconstituted in bilayers formed at the tip of patch pipets. Biochemistry 22:2319–2323 2. Zimmerberg J, Parsegian VA (1986) Polymer inaccessible volume changes during opening and closing of a voltage-dependent ionic channel. Nature 323:36–39 3. Han A, Schu¨rmann G, Mondin G, Bitterli RA, Hegelbach NG, De Rooij NF, Staufer U (2006) Sensing protein molecules using nanofabricated pores. Appl Phys Lett 88:1–4 4. Fologea D, Ledden B, McNabb DS, Li J (2007) Electrical characterization of protein molecules by a solid-state nanopore. Appl Phys Lett 91:53901-1–53901-3 5. Talaga DS, Li J (2009) Single-molecule protein unfolding in solid state nanopores. J Am Chem Soc 131:9287–9297 6. Nir I, Huttner D, Meller A (2015) Direct sensing and discrimination among ubiquitin and ubiquitin chains using solid-state nanopores. Biophys J 108:2340–2349 7. Meervelt V Van, Soskine M, Maglia G (2014) Detection of two isomeric binding configurations in a protein aptamer complex with a biological nanopore. ACS Nano 8:12826–12835 8. Willems K, Ruic´ D, Biesemans A, Galenkamp NS, Van Dorpe P, Maglia G (2019) Engineering and modeling the electrophoretic trapping of a single protein inside a nanopore. ACS Nano 13:9980–9992

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



1 Preparation of Fragaceatoxin C (FraC) Nanopores . . . . . . . . . . . . . . . . . . . . . . . . . . Natalie Lisa Mutter, Gang Huang, Nieck Jordy van der Heide, Florian Leonardus Rudolfus Lucas, Nicole Ste´phanie Galenkamp, Giovanni Maglia, and Carsten Wloka 2 Preparation of Cytolysin A (ClyA) Nanopores . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nicole Ste´phanie Galenkamp, Veerle Van Meervelt, Natalie Lisa Mutter, Nieck Jordy van der Heide, Carsten Wloka, and Giovanni Maglia 3 Building Synthetic Transmembrane Peptide Pores . . . . . . . . . . . . . . . . . . . . . . . . . . Kozhinjampara R. Mahendran 4 Design and Assembly of Membrane-Spanning DNA Nanopores . . . . . . . . . . . . . . Kerstin Go¨pfrich, Alexander Ohmann, and Ulrich F. Keyser


v ix



19 33


5 Determining the Orientation of Porins in Planar Lipid Bilayers . . . . . . . . . . . . . . . 51 Sandra A. Ionescu, Sejeong Lee, and Hagan Bayley 6 Revelation of Function and Inhibition of Wza Through Single-Channel Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63 Lingbing Kong 7 Protein Analyte Sensing with an Outer Membrane Protein G (OmpG) Nanopore . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77 Monifa A. V. Fahie, Bib Yang, Christina M. Chisholm, and Min Chen 8 Nanopore Enzymology to Study Protein Kinases and Their Inhibition by Small Molecules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Leon Harrington, Leila T. Alexander, Stefan Knapp, and Hagan Bayley 9 A Selective Activity-Based Approach for Analysis of Enzymes with an OmpG Nanopore . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115 Monifa A. V. Fahie, Bach Pham, Fanjun Li, and Min Chen 10 Oligonucleotide-Directed Protein Threading Through a Rigid Nanopore . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 ˜ e Celaya and David Rodriguez-Larrea Garbin 11 Unfolding and Translocation of Proteins Through an Alpha-Hemolysin Nanopore by ClpXP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 145 Jeff Nivala, Logan Mulroney, Qing Luan, Robin Abu-Shumays, and Mark Akeson






Simulation of pH-Dependent, Loop-Based Membrane Protein Gating Using Pretzel . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 159 Alan Perez-Rathke, Monifa A. V. Fahie, Christina M. Chisholm, Min Chen, and Jie Liang Free Energy Minimization for Vesicle Translocation Through a Narrow Pore. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 171 Hamid R. Shojaei, Ahad Khaleghi Ardabili, and Murrugappan Muthukumar

PART IV 14 15




Single Ion-Channel Analysis in Droplet Interface Bilayer . . . . . . . . . . . . . . . . . . . . 187 Arash Manafirad Continuous and Rapid Solution Exchange in a Lipid Bilayer Perfusion System Based on Droplet-Interface Bilayer. . . . . . . . . . . . . . . . . . . . . . . . 197 En-Hsin Lee Protein Transport Studied by a Model Asymmetric Membrane Army Arranged in a Dimple Chip . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 213 Xin Li and Min Chen

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


Contributors ROBIN ABU-SHUMAYS • UC Santa Cruz Genomics Institute, University of California, Santa Cruz, Santa Cruz, CA, USA MARK AKESON • UC Santa Cruz Genomics Institute, University of California, Santa Cruz, Santa Cruz, CA, USA LEILA T. ALEXANDER • Nuffield Department of Clinical Medicine, Structural Genomics Consortium and Target Discovery Institute, University of Oxford, Oxford, UK; Personalized Health Informatics, SIB Swiss Institute of Bioinformatics, Basel, Switzerland HAGAN BAYLEY • Department of Chemistry, University of Oxford, Oxford, UK GARBIN˜E CELAYA • Department of Biochemistry and Molecular Biology (UPV/EHU), Biofisika Institute (CSIC, UPV/EHU), Leioa, Spain MIN CHEN • Department of Chemistry, University of Massachusetts Amherst, Amherst, MA, USA CHRISTINA M. CHISHOLM • Molecular and Cellular Biology Program, University of Massachusetts Amherst, Amherst, MA, USA MONIFA A. V. FAHIE • Molecular and Cellular Biology Program, University of Massachusetts Amherst, Amherst, MA, USA NICOLE STE´PHANIE GALENKAMP • Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, The Netherlands KERSTIN GO¨PFRICH • Cavendish Laboratory, University of Cambridge, Cambridge, UK LEON HARRINGTON • Department of Chemistry, University of Oxford, Oxford, UK; Max Planck Institute of Biochemistry, Martinsried, Germany GANG HUANG • Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, The Netherlands SANDRA A. IONESCU • Department of Chemistry, University of Oxford, Oxford, UK ULRICH F. KEYSER • Cavendish Laboratory, University of Cambridge, Cambridge, UK AHAD KHALEGHI ARDABILI • School of Engineering and Natural Sciences, Altınbas¸ University, Istanbul, Turkey STEFAN KNAPP • Nuffield Department of Clinical Medicine, Structural Genomics Consortium and Target Discovery Institute, University of Oxford, Oxford, UK; Institute for Pharmaceutical Chemistry, Structural Genomics Consortium, and Buchmann Institute for Molecular Life Sciences, Johann Wolfgang Goethe-University, Frankfurt am Main, Germany LINGBING KONG • International Institute of Rare Sugar Research and Education, Department of Applied Biological Science, Faculty of Agriculture, Kagawa University, Miki, Kagawa, Japan EN-HSIN LEE • Department of Chemistry, University of Massachusetts Amherst, Amherst, MA, USA SEJEONG LEE • Department of Chemistry, University of Oxford, Oxford, UK JIE LIANG • Department of Bioengineering, University of Illinois at Chicago, Chicago, IL, USA FANJUN LI • Department of Chemistry, University of Massachusetts Amherst, Amherst, MA, USA XIN LI • Department of Chemistry, University of Massachusetts Amherst, Amherst, MA, USA




QING LUAN • Department of Chemistry and Biochemistry, University of Notre Dame, Notre Dame, IN, USA FLORIAN LEONARDUS RUDOLFUS LUCAS • Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, The Netherlands GIOVANNI MAGLIA • Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, The Netherlands KOZHINJAMPARA R. MAHENDRAN • Membrane Biology Laboratory, Interdisciplinary Research Program, Rajiv Gandhi Centre for Biotechnology, Thiruvananthapuram, India ARASH MANAFIRAD • Department of Physics, University of Massachusetts Amherst, Amherst, MA, USA; Department of Chemistry, University of Massachusetts Amherst, Amherst, MA, USA LOGAN MULRONEY • UC Santa Cruz Genomics Institute, University of California, Santa Cruz, Santa Cruz, CA, USA MURRUGAPPAN MUTHUKUMAR • Department of Polymer Science and Engineering, University of Massachusetts Amherst, Amherst, MA, USA NATALIE LISA MUTTER • Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, The Netherlands JEFF NIVALA • Paul G. Allen School of Computer Science and Engineering, University of Washington, Seattle, WA, USA ALEXANDER OHMANN • Cavendish Laboratory, University of Cambridge, Cambridge, UK ALAN PEREZ-RATHKE • Department of Bioengineering, University of Illinois at Chicago, Chicago, IL, USA BACH PHAM • Department of Chemistry, University of Massachusetts Amherst, Amherst, MA, USA DAVID RODRIGUEZ-LARREA • Department of Biochemistry and Molecular Biology (UPV/ EHU), Biofisika Institute (CSIC, UPV/EHU), Leioa, Spain HAMID R. SHOJAEI • Department of Polymer Science and Engineering, University of Massachusetts Amherst, Amherst, MA, USA NIECK JORDY VAN DER HEIDE • Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, The Netherlands VEERLE VAN MEERVELT • Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, The Netherlands CARSTEN WLOKA • Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, The Netherlands BIB YANG • Department of Chemistry, University of Massachusetts Amherst, Amherst, MA, USA

Part I Design and Preparation of Biological Based Nanopores

Chapter 1 Preparation of Fragaceatoxin C (FraC) Nanopores Natalie Lisa Mutter, Gang Huang, Nieck Jordy van der Heide, Florian Leonardus Rudolfus Lucas, Nicole Ste´phanie Galenkamp, Giovanni Maglia, and Carsten Wloka Abstract Biological nanopores are an emerging class of biosensors with high-end precision owing to their reproducible fabrication at the nanometer scale. Most notably, nanopore-based DNA sequencing applications are currently being commercialized, while nanopore-based proteomics may become a reality in the near future. Although membrane proteins often prove to be difficult to purify, we describe a straightforward protocol for the preparation of Fragaceatoxin C (FraC) nanopores, which may have applications for DNA analysis and nanopore-based proteomics. Recombinantly expressed FraC nanopores are purified via two rounds of Ni-NTA affinity chromatography before and after oligomerization on sphingomyelin-containing liposomes. Starting from a plasmid vector containing the FraC gene, our method allows the production of purified nanopores within a week. Afterward, the FraC nanopores can be stored at +4  C for several months, or frozen. Key words Nanotechnology, Nanopore, Porin, Actinoporin, Fragaceatoxin C, FraC, Protein purification, Protein oligomerization, Electrophysiology, Artificial bilayers


Introduction The understanding of single ion channels was revolutionized over four decades ago when it was shown that single channel currents can be recorded [1]. The small ion current flowing through a single channel in a planar lipid bilayer [2] has been subsequently used for single-molecule studies. Single channels (or nanopores) with a β-barrel comprise toxins such as α-hemolysin (αHL) [3, 4], aerolysin [5, 6], Mycobacterium smegmatis porin A (MspA) [7], ferric hydroxamate uptake component A (FhuA) [8], and outer membrane proteins (OmpG [9–11], OmpF [12], OmpC [13]), and have been used for the detection of small molecules as well as for DNA, protein, and peptide analysis. Our laboratory introduced the α-helical Cytolysin A (ClyA) nanopore from Salmonella typhi for studies of proteins [14] and DNA [15]. More recently the

Monifa A. V. Fahie (ed.), Nanopore Technology: Methods and Protocols, Methods in Molecular Biology, vol. 2186,, © Springer Science+Business Media, LLC, part of Springer Nature 2021



Natalie Lisa Mutter et al.

Fragaceatoxin C (FraC) nanopore [16] has been introduced in our lab for peptide detection. The octameric crystal structure of FraC has been disclosed [17] after being discovered to be an ingredient of the venom of the strawberry anemone (Actinia fragacea) [18]. This α-helical pore forms a relatively narrow (V-shaped) trans entry, which proved to be useful for the analysis of DNA [19] and physiologically important proteins and peptides [16, 20]. Based on previous work [17, 21], we describe here an easy-tofollow protocol to obtain FraC nanopores. In this protocol, histidine-tagged FraC monomers are recombinantly expressed in Escherichia coli and purified using Ni-NTA affinity chromatography. Monomers are then assembled into oligomers on sphingomyelin-containing liposomes. In the final step, liposomes are solubilized and oligomeric FraC nanopores are purified using a second round of Ni-NTA affinity chromatography. FraC nanopores can be obtained within a week and stored at 80  C as well as at 4  C for several months.


Materials All solutions are prepared using deionized water (Millipore). Special attention should be paid to the final pH of all solutions, in particular to the high amount of imidazole-containing buffers (e.g., elution buffers). All reagents can be prepared and stored at room temperature (~25  C), unless otherwise noted.

2.1 Transformation and Expression of FraC in Escherichia coli BL21 (DE3) Strain

1. Sterile Lysogeny broth (LB). 2. Sterile 2 YT medium. 3. 1 M Isopropyl-β-D-thiogalactopyranoside (IPTG). Store at 20  C. 4. 100 mg/mL Ampicillin. Store at

20  C.

5. LB-Agar plates containing 1% (w/v) glucose (optional) and 100 μg/mL ampicillin. Store at 4  C. 6. 50 μL aliquot of electrocompetent E. cloni® EXPRESS BL21 (DE3) (Lucigen, stored at 80  C). 7. Fragaceatoxin C pT7-SC1 [22] plasmid DNA containing a C-terminal His6-tag (75–150 ng/μL DNA), stored at 20  C. 8. LB medium containing 1% (w/v) glucose (500 μL). 9. 100 mL Erlenmeyer flask (starter culture). 10. 1 L Erlenmeyer flask (expression culture). 11. Centrifuge capable of maintaining 4  C and 5000  g.

Preparation of FraC Nanopores


12. Spectrophotometer. 13. Shaking incubator capable of maintaining 37 and 20  C as well as 200 rpm. 14. Electroporation device and electroporation cuvettes. 15. Crushed ice. 16. T-shaped spreader, sterile. 2.2 Purification of Fragaceatoxin C Monomers

1. Lysis buffer: 0.015 M Tris–HCl, pH 7.5, 0.150 M NaCl, 0.02 M imidazole, 1 mM MgCl2, and 4 M urea (see Note 1). Store at 4  C for up to 1 month. Freshly prepared buffers are preferred, however. 2. Wash buffer (WB): 0.015 M Tris–HCl, pH 7.5, 0.150 M NaCl, 0.02 M imidazole. Store at 4  C. 3. Elution buffer (EB): 0.015 M Tris–HCl, pH 7.5, 0.150 M NaCl, 0.300 M imidazole. Store at 4  C (see Note 2). 4. DNase I. 5. Chicken Lysozyme powder. 6. Sonication device. 7. Falcon™ tubes and compatible shaking/turning device. 8. Gravity flow columns with a volume of 1–2 mL. 9. Ni-NTA beads.

2.3 Preparation of Sphingomyelin: DPhPC Liposomes

1. Dilution buffer (DB): 0.015 M of Tris–HCl, pH 7.5, 0.150 M of NaCl. Store at 4  C. 2. Sphingomyelin (Brain, Porcine). 3. 1,2-Diphytanoyl-sn-glycero-3-phosphocholine (DPhPC). 4. Pentane (99%). 5. Ethanol, absolute (99.8%). 6. Round-bottom flask (25 mL) and rotary evaporator (or heat gun). 7. Sonication water bath.

2.4 Oligomerization of Fragaceatoxin C Monomers

1. 5% (w/v) N,N-Dimethyldodecylamine-N-oxide (LDAO). Store at 20  C (room temperature is also acceptable). 2. 10% (w/v) Dodecyl-β-D-maltoside (DDM). Store, for example, 1 mL aliquots at 20  C. 3. Elution buffer for oligomers (EBO). 0.200 M Na2EDTA, pH 8.0, 0.02% (w/v) DDM (see Note 3). Store at 4  C.



Natalie Lisa Mutter et al.


3.1 Transformation and Expression of Fragaceatoxin C in an Escherichia coli BL21 (DE3) Strain

1. Transform the FraC plasmid DNA into the electrocompetent E. cloni® EXPRESS BL21 (DE3) strain for protein expression. Place the electrocompetent cells on ice and add 1 μL plasmid DNA (75–150 ng/μL DNA) to 50 μL cells. Transfer the cells to an electroporation cuvette (cooled down on ice) and electroporate (settings: bacteria, 5 ms pulse). 2. Add 500 μL prewarmed LB supplemented with 1% (w/v) glucose quickly to the cells and incubate for 1 h at 37  C. 3. Plate the transformed cells out on LB plates containing 100 μg/mL ampicillin and 1% (w/v) glucose. Add 300 μL cells to one plate and 100 μL cells to another plate, to ensure well-spaced colonies. Incubate the plates at 37  C overnight. 4. The following day (day 2), inoculate 10 mL LB medium supplemented with 100 μg/mL ampicillin with a single colony and incubate at 37  C and 200 rpm overnight in a 100 mL Erlenmeyer flask. 5. On day 3, inoculate 200 mL 2 YT medium containing 100 μg/mL ampicillin (expression culture) with 2 mL of the overnight starter culture. This dilutes the culture 1:100. Incubate the expression culture for 2–3 h at 37  C and 200 rpm until the optical density at 600 nm wavelength (OD600) is between 0.6 and 0.8. 6. Transfer the expression culture to 20  C and 200 rpm, add 100 μL IPTG for a final concentration of 0.5 mM IPTG, to induce protein expression, and continue growth overnight.

3.2 Purification of Fragaceatoxin C Monomers

1. Harvest cells in 50 mL Falcon™ tubes by centrifugation at 5000  g (30 min at 4  C) and discard the supernatant (see Note 4). 2. Freeze the bacterial cell pellets for at least 1 h at Note 5).

80  C (see

3. Solubilize two bacterial cell pellets (100 mL of the expression culture total) in 20 mL lysis buffer (see Note 6). 4. Add 0.2 units/mL of DNase I and 20 μg/mL of lysozyme to each resuspended pellet (see Note 7). Incubate the pellets for 30 min at room temperature (~25  C) in a shaking device. 5. Sonicate by putting the tip of the sonication device at least 3 cm beneath the surface of the liquid (see Note 8). Sonicate on ice at 30% output power for 3  30 s to break the cells open (see Note 9). 6. Pellet the cell debris by centrifugation for 30 min at 5000  g and 4  C. Discard the pellet. The protein is in the supernatant.

Preparation of FraC Nanopores


7. Wash 100 μL of Ni-NTA beads (~200 μL of Ni-NTA slurry) with wash buffer (WB), add 500 μL WB to the Ni-NTA beads and centrifuge for 30 s at 10,000  g, discard the supernatant and repeat three times (see Note 10). 8. Incubate the supernatant from step 6 with the washed Ni-NTA beads for 1 h at room temperature (~25  C) on a shaking device or tube rotator to prevent sinking of Ni-NTA beads. 9. Prewash a gravity flow column with about five column volumes (~ 1 mL, depending on the column used) WB to check the flow and equilibrate the column. 10. Add supernatant from step 8 containing the incubated Ni-NTA beads into the column (see Note 10). 11. Wash the column with 10 mL of WB. After washing, remove the WB by putting the column in a clean microtube and centrifuge at 1500  g for 30 s (see Note 11). 12. Elute the monomers from the Ni-NTA beads with five elution steps of 150 μL EB. Incubate for 5–10 min with EB and centrifuge (1500  g, 30 s) in order to collect elution fractions in clean microtubes. 13. Check the protein concentration of the monomers with a spectrophotometer at wavelength 280 nm (see Note 12). The experimentally determined extinction coefficient is 1 1 38,688 M cm . 14. Store the monomers at 4  C overnight or for up to a week, or continue with the next steps (see Note 13). 3.3 Preparation of Sphingomyelin: DPhPC Liposomes

1. Solubilize 25 mg sphingomyelin and 25 mg DPhPC (1:1) in approximately 2 mL pentane and about 200 μL of ethanol in a round-bottom flask (see Note 14). 2. Make a film of sphingomyelin:DPhPC on the surface of the round-bottom flask by rotary evaporation. Keep the flask open for 30 min to let all solvent evaporate (see Note 15). 3. Solubilize the lipid film in 5 mL dilution buffer (DB) to a final concentration of 10 mg/mL. 4. Briefly sonicate the lipid film solution in a sonication bath and freeze-thaw at least one time, freezing at 20  C (see Note 16).

3.4 Oligomerization of Fragaceatoxin C Monomers

1. Add sphingomyelin:DPhPC liposomes to FraC monomers in a mass ratio of 10:1 (lipid:protein), mix thoroughly/vortex, and incubate for 30 min at 37  C (see Notes 17 and 18). 2. Solubilize proteoliposomes by adding a final concentration of 0.6% (w/v) LDAO. The solution should turn clear immediately.


Natalie Lisa Mutter et al.

3. Dilute the solubilized mixture 20-fold with dilution buffer supplemented with 0.02% (w/v) DDM. 4. Wash 50 μL of Ni-NTA beads (100 μL of Ni-NTA suspension) with WB supplemented with 0.02% (w/v) DDM. Add 500 μL WB containing 0.02% (w/v) DDM to the Ni-NTA beads, centrifuge for 30 s at 10,000  g, discard the supernatant, and repeat this wash three times. 5. Incubate the oligomer mixture with the washed Ni-NTA beads for 1 h at room temperature (~25  C) while shaking (using an orbital shaker/tube rotator). 6. Prewash a gravity flow column with 5 column volumes (~ 1 mL) WB supplemented with 0.02% (w/v) DDM. 7. Add the incubated Ni-NTA beads to the column (see Note 10). 8. Wash the FraC oligomers incubated with the Ni-NTA beads with 10 mL of WB supplemented with 0.02% (w/v) DDM. Remove excess WB containing 0.02% (w/v) DDM by centrifuging at 1500  g for 30 s (see Note 11). 9. Elute the oligomers from the Ni-NTA beads in one elution step with 100 μL EBO. Incubate the Ni-NTA beads for 5–10 min with EBO and centrifuge (1500  g, 30 s) to collect the elution in a clean microtube (see Note 19). 10. Oligomers can be stored for several months at 4  C. Store oligomers at 80  C for longer periods (see Note 20). 11. Oligomers can be used in electrophysiology as described before [23].


Notes 1. Concentrated HCl can be used first to get close to the required pH. Then, use less concentrated HCl to avoid the pH getting below the desired value. 2. When stored at 4  C, the buffer can be used for a longer period of time. 3. A pH of 8.0 is required to dissolve the Na2EDTA. The NaOH powder should be used to bring the pH close to the required pH. Use a lower-concentrated NaOH solution (1 M) to avoid the pH getting above the required value. 4. Before centrifugation, balance the tubes on a weighing scale to a difference of less than 0.1 g. Use buffer or transfer cell culture from one tube to another in order to balance. 5. Pellets frozen at 80  C can be kept for long-term storage, allowing purification at a later time.

Preparation of FraC Nanopores


6. Pellets can be combined in this step in a ratio of 1 mL lysis buffer per 5 mL bacterial cells. 7. The DNase I is added to break down DNA and the lysozyme is added to break down the bacterial cell wall. 8. Consult manufacturer instructions on how to operate the sonication device. 9. It is also possible to sonicate the cells for 2 min continuously on ice. Do not overheat the sample. 10. Ni-NTA beads settle to the bottom; therefore make sure you resuspend the Ni-NTA beads before pipetting. 11. After spinning the Ni-NTA beads to dryness, add EB directly. 12. Expect a yield of about 0.5 mg/mL or less from 100 mL expression culture. 13. Freezing of monomers results in precipitation of the proteins. 14. The solvents should evaporate relatively fast, so make sure the pentane and ethanol you use contain little water. 15. A rotary evaporator can be used. However, manual turning and heat application to evaporate solvents, using a heat gun for example, will achieve the same result. It is crucial that all solvent is evaporated. 16. Freeze-thawing may be repeated many times. It creates fewer multilamellar liposomes; thus, they will be more uniform. 17. To yield smaller FraC pores [20], it is also possible to use sphingomyelin-containing liposomes in a mass ratio of 1:1 instead. 18. Incubation for up to 1 h does not influence the oligomerization. 19. It is also possible to use a different elution buffer containing imidazole instead of EDTA. Add 100 μL DB supplemented with 0.02% (w/v) DDM and 1 M imidazole. Adjust the pH of the solution to 8.0. Incubate for 5–20 min, and centrifuge afterward to collect the elution. 20. FraC nanopores can be stored at 4  C for several months. It is possible to store oligomers at 80  C. After thawing, store at 4  C. References 1. Neher E, Sakmann B (1976) Single-channel currents recorded from membrane of denervated frog muscle fibres. Nature 260:619–621 2. Moczydlowski E, Latorre R (1983) Gating kinetics of Ca2+-activated K+ channels from rat muscle incorporated into planar lipid bilayers. Evidence for two voltage-dependent

Ca2+ binding reactions. J Gen Physiol 82:511–542 3. Division B, Biology C, Cruz S (1996) Characterization of individual polynucleotide molecules using a membrane channel. Proc Natl Acad Sci 93:13770–13773


Natalie Lisa Mutter et al.

4. Gu L-Q, Braha O, Conlan S, Cheley S, Bayley H (1999) Stochastic sensing of organic analytes by a pore-forming protein containing amolecular adapter. Nature 398:686–690 5. Cao C, Ying YL, Hu ZL, Liao DF, Tian H, Long YT (2016) Discrimination of oligonucleotides of different lengths with a wild-type aerolysin nanopore. Nat Nanotechnol 11:713–718 6. Piguet F, Ouldali H, Pastoriza-Gallego M, Manivet P, Pelta J, Oukhaled A (2018) Identification of single amino acid differences in uniformly charged homopolymeric peptides with aerolysin nanopore. Nat Commun 9:966 7. Manrao EA, Derrington IM, Laszlo AH, Langford KW, Hopper MK, Gillgren N, Pavlenok M, Niederweis M, Gundlach JH (2012) Reading DNA at single-nucleotide resolution with a mutant MspA nanopore and phi29 DNA polymerase. Nat Biotechnol 30:349–353 8. Mohammad MM, Iyer R, Howard KR, McPike MP, Borer PN, Movileanu L (2012) Engineering a rigid protein tunnel for biomolecular detection. J Am Chem Soc 134:9521–9531 9. Fahie MA, Yang B, Mullis M, Holden MA, Chen M (2015) Selective detection of protein homologues in serum using an OmpG nanopore. Anal Chem 87:11143–11149 10. Fahie MA, Yang B, Pham B, Chen M (2016) Tuning the selectivity and sensitivity of an OmpG nanopore sensor by adjusting ligand tether length. ACS Sensors 1:614–622 11. Kahlstatt J, Reiß P, Halbritter T, Essen LO, Koert U, Heckel A (2018) A light-triggered transmembrane porin. Chem Commun 54:9623–9626 12. Ghai I, Bajaj H, Bafna JA, El Damrany Hussein HA, Winterhalter M, Wagner R (2018) Ampicillin permeation across OmpF, the major outer-membrane channel in Escherichia coli. J Biol Chem 293:7030–7037 13. Prajapati JD, Solano CJF, Winterhalter M, Kleinekatho¨fer U (2018) Enrofloxacin permeation pathways across the porin OmpC. J Phys Chem B 122:1417–1426

14. Wloka C, Van Meervelt V, Van Gelder D, Danda N, Jager N, Williams CP, Maglia G (2017) Label-free and real-time detection of protein ubiquitination with a biological nanopore. ACS Nano 11:4387–4394 15. Franceschini L, Brouns T, Willems K, Carlon E, Maglia G (2016) DNA translocation through nanopores at physiological ionic strengths requires precise nanoscale engineering. ACS Nano 10:8394–8402 16. Huang G, Willems K, Soskine M, Wloka C, Maglia G (2017) Electro-osmotic capture and ionic discrimination of peptide and protein biomarkers with FraC nanopores. Nat Commun 8:935 17. Tanaka K, Caaveiro JMM, Morante K, Gonza´˜ as JM, Tsumoto K (2015) Structural lez-Man basis for self-assembly of a cytolytic pore lined by protein and lipid. Nat Commun 6:6337 18. Bellomio A, Morante K, Barlicˇ A, Gutie´rrez˜ as JM Aguirre I, Viguera AR, Gonza´lez-Man (2009) Purification, cloning and characterization of fragaceatoxin C, a novel actinoporin from the sea anemone Actinia fragacea. Toxicon 54:869–880 19. Wloka C, Mutter NL, Soskine M, Maglia G (2016) Alpha-helical fragaceatoxin C nanopore engineered for double-stranded and singlestranded nucleic acid analysis. Angew Chem Int Ed Engl 55:12494–12498 20. Huang G, Voet A, Maglia G (2019) FraC nanopores with adjustable diameter identify the mass of opposite-charge peptides with 44 dalton resolution. Nat Commun 10:835 21. Tanaka K, Caaveiro JMM, Tsumoto K (2015) Bidirectional transformation of a metamorphic protein between the water-soluble and transmembrane native states. Biochemistry 54:6863–6866 22. Miles G, Cheley S, Braha O, Bayley H (2001) The staphylococcal leukocidin bicomponent toxin forms large ionic channels. Biochemistry 40:8514–8522 23. Maglia G, Heron AJ, Stoddart D, Japrung D, Bayley H (2010) Analysis of single nucleic acid molecules with protein nanopores. Methods Enzymol 475:591–623

Chapter 2 Preparation of Cytolysin A (ClyA) Nanopores Nicole Ste´phanie Galenkamp, Veerle Van Meervelt, Natalie Lisa Mutter, Nieck Jordy van der Heide, Carsten Wloka, and Giovanni Maglia Abstract The ionic currents passing through nanopores can be used to sequence DNA and identify molecules at the single-molecule level. Recently, researchers have started using nanopores for the detection and analysis of proteins, providing a new platform for single-molecule enzymology studies and more efficient biomolecular sensing applications. For this approach, the homo-oligomeric Cytolysin A (ClyA) nanopore has been demonstrated as a powerful tool. Here, we describe a simple protocol allowing the production of ClyA nanopores. Monomers of ClyA are expressed in Escherichia coli and oligomerized in the presence of detergent. Subsequently, different oligomer variants are electrophoretically resolved and stored in a gel matrix for long-term use. Key words Nanopores, Cytolysin A, ClyA, Protein purification, Membrane protein, Protein assembly, Pore-forming toxin, Single-molecule, Electrophysiology, Protein trapping


Introduction In a typical nanopore experiment, a potential is applied across a nanopore reconstituted into a lipid bilayer. The resulting ionic current flowing through nanopores provides the signal by which molecules are recognized, studied, or sequenced. Over the past 20 years, nanopores have been used to sequence DNA, recognize small molecules, and sample chemical reactions at the single-molecule level [1–9]. Advantages include high resolution on a micro- to millisecond timescale [10], and the ability to observe native single molecules with no intrinsic limitation of molecular size or observation time. Because of their single-molecule nature, nanopore experiments can unravel processes and transient intermediates normally hidden in ensemble measurements [11, 12]. Single-molecule analysis with Cytolysin A nanopores has been introduced 7 years ago by our laboratory [13]. The crystal structure of the nanopore revealed a 13 nm long nanopore comprising 12 identical protomers [14], with a 5.5 nm cis entry and 3.3 nm

Monifa A. V. Fahie (ed.), Nanopore Technology: Methods and Protocols, Methods in Molecular Biology, vol. 2186,, © Springer Science+Business Media, LLC, part of Springer Nature 2021



Nicole Ste´phanie Galenkamp et al.

trans entry, which allows the entry of proteins or dsDNA [15]. Under a negative applied potential, folded proteins between ~20 and ~40 kDa added to the cis side of a dodecameric nanopore can be accommodated inside its lumen exerted by the electroosmotic flow [9, 16–18]. Importantly, the narrower trans entry of the nanopore (3.3 nm) allows trapping of proteins without the need of labeling, immobilization, or chemical modification [9, 17– 19]. Furthermore, the ionic current is large enough to permit working at physiological salt conditions (150 mM), which is important when sampling protein–protein, protein–DNA interactions [20] or dsDNA [15]. Inside the nanopore, proteins can be identified by their specific ionic current blockades. Notably, the asymmetric environment of the nanopore lumen also allowed resolving isomeric protein–DNA interactions [20]. Furthermore, the ionic current across the nanopore was exquisitely sensitive to small changes in the structure of the trapped protein, which in turn allowed sampling protein conformational changes, due to binding of substrates [9, 17]. Hence, ClyA nanopores can be used in single-molecule enzymology studies [21], while the ability of identifying ligand binding to internalized protein has been exploited to quantify metabolites in bodily fluids [18]. In our method, we describe the recombinant expression of histidine-tagged ClyA monomers in Escherichia coli, purification of the monomers using Ni-NTA affinity chromatography, and assembly into oligomers by addition of detergent. Because ClyA monomers are water-soluble and assemble into nanopores, only upon addition of a detergent (or liposomes) large amounts of nanopores can easily be obtained. Finally, we describe how to resolve the ClyA oligomers by separation on blue-native polyacrylamide gel electrophoresis. Within 1 week ClyA nanopores can be obtained and the nanopores can be stored long-term at 4 or 20  C when embedded within the gel matrix.


Materials Deionized water is used for all solutions and care is given to the final pH of the solution. Unless otherwise noted, reagents are prepared and stored at 25  C.

2.1 Transformation and Expression of ClyA-AS in Escherichia coli

1. 1 L Lysogeny broth (LB). Weigh 10 g of bacto tryptone, 5 g of yeast extract, and 10 g of NaCl and transfer to a 1.5 or 2 L bottle. Add water (about 800–900 mL), dissolve all components by mixing, and fill up to 1 L with water. Sterilize by autoclaving. Store at room temperature. 2. 1 L 2 YT-medium. Weigh 16 g of bacto tryptone, 10 g of bacto yeast extract, and 5 g of NaCl and transfer to a 1.5 or 2 L

Preparation of ClyA Nanopores


bottle. Mix components in 900 mL water, then fill up to 1 L with water. Sterilize by autoclaving. Store at room temperature. 3. 10 mL of 1 M Isopropyl-β-D-thiogalactopyranoside (IPTG). Add 2.38 g IPTG to 10 mL water and mix well. Aliquots can be stored at 20  C. 4. 10 mL of 100 mg/mL (1000) ampicillin. Add 1.0 g ampicillin to 10 mL water and mix well. Aliquots can be stored at 20  C. 5. LB-agar plates: LB-agar plates containing 1% (w/v) glucose and 100 μg/mL ampicillin which should be stored at 4  C. For 1 L of LB agar: add 15 g of bacto agar to 1.0 L of LB medium and autoclave. Use a temperature-controlled water bath to cool the LB medium to 60  C and add 1 mL of 100 mg/mL ampicillin and 10 g of glucose. In a sterile environment, pour plates (about 30 mL per dish). Dry the plates for 30 min. Store at 4  C. 6. 50 μL Electrocompetent E. cloni® EXPRESS BL21 (DE3) (Lucigen, stored at 80  C). 7. pT7-SC1 [22] plasmid containing the ClyA-AS gene with C-terminal His6-tag (75–150 ng/μL DNA), stored at 20  C. ClyA-AS contains eight mutations relative to the Salmonella typhi ClyA-WT: C87A, L99Q, E103G, F166Y, I203V, C285S, K294R, and H307Y (the H307Y mutation is in the C-terminal His6-tag added for purification). 8. LB medium with 1% (w/v) glucose (400 μL). 9. 100 mL Erlenmeyer flask for the starter culture. 10. 1 L Erlenmeyer flask for the expression culture. 11. Cooled centrifuge (4  C). 12. Spectrophotometer, such as a Nanodrop. 13. Shaking incubator (37 and 20  C). 14. Electroporation device and electroporation cuvettes. 15. Sterile T-shaped spreader. 16. Ice. 2.2 For Purification of ClyA-AS

1. 1 L Lysis buffer: 15 mM Tris–HCl, pH 7.5, 150 mM NaCl, 10 mM imidazole, pH 8.0, 1 mM MgCl2. Weigh 1.82 g Tris– HCl, 8.77 g NaCl, 0.68 g imidazole, and 95.2 mg MgCl2, and transfer to a 1 L glass bottle. Add about 900 mL water to dissolve all components. Adjust the pH to 7.5 with HCl. Fill up with water to 1 L and store at room temperature for up to 2 months. Fresh buffers are preferred, however. If the buffer is stored for a period of time, the pH should be checked regularly.


Nicole Ste´phanie Galenkamp et al.

2. 1 L Wash buffer (WB): 10 mM imidazole, 150 mM NaCl, 15 mM Tris–HCl, pH 7.5. Weigh 1.82 g Tris–HCl, 8.77 g NaCl, and 0.68 g imidazole, and transfer to a 1 L glass bottle. Add about 900 mL water and mix until all components are dissolved. Adjust the pH to 7.5 with HCl. Fill up to 1 L with water and store at 4  C. 3. 100 mL Elution buffer (EB): 150 mM NaCl, 15 mM Tris– HCl, pH 7.5 supplemented with 300 mM imidazole, pH 8. Weigh 182 mg Tris–HCl, 877 mg NaCl, and 2.04 g imidazole. Transfer to a 100 mL glass bottle. Add about 80 mL water and mix until all components are dissolved. Adjust the pH to 7.5 with HCl. Fill to 100 mL water and store at 4  C. 4. 1 mL Extraction buffer: 150 mM NaCl, 15 mM Tris–HCl, pH 7.5 supplemented with 0.2% (w/v) β-dodecylmaltoside (DDM) and 5 mM EDTA. Weigh 1.82 mg Tris–HCl and 8.77 mg NaCl and transfer to a microtube. Add water to a volume of 800 μL and mix until all components are dissolved. Adjust the pH to 7.5 with HCl. Add 20 μL of 10% (w/v) DDM stock and 1.46 mg of EDTA. Fill to 1 mL and store at 20  C. 5. DNase I (1 U/μL). 6. Lysozyme (Chicken). 7. Vortex. 8. Centrifuge tubes and compatible shaking/rotating device. 9. Sonication device. 10. Ni-NTA resin (e.g., Qiagen). 11. Cooled centrifuge (4  C). 12. Gravity flow columns. 13. Nanodrop or Bradford assay to measure protein concentration. 14. 10% (w/v) DDM stock prepared in deionized water. Weigh 1 g DDM and transfer to a 10 mL tube. Fill up to 10 mL with water and mix until all DDM is dissolved. Store aliquots at 20  C. 15. Criterion TGX 4–20% polyacrylamide pre-cast gels (Bio-rad). 16. 20 native running buffer (Invitrogen). 17. 20 cathode buffer additive (Invitrogen). 18. 5 blue native sample buffer (BN SB): mix 2.5 mL of 20 native running buffer with 4 g of glycerol (99%), 2.5 mL of 20 cathode buffer additive and 1 mL of deionized water and store aliquots at 20  C. 19. Electrophoresis cell and power supply fitting the respective gel system. 20. Surgical blade.

Preparation of ClyA Nanopores




3.1 Transformation and Recombinant Expression of ClyA-AS in Escherichia coli (Day 1–3)

1. Transform electrocompetent E. cloni® EXPRESS BL21 (DE3) with pT7-SC1 plasmid DNA containing the ClyA-AS gene. On ice, add 1 μL DNA (75–150 ng/μL plasmid) to 50 μL cells and mix gently. Add the mixture to a precooled electroporation cuvette and electroporate (2  5 ms pulse, setting: bacteria). 2. Immediately add 400 μL pre-warmed (37  C) LB medium containing 1% (w/v) glucose to the cells and incubate for 1 h at 37  C. 3. Transfer the cell suspension to a LB agar plate, supplemented with 1% (w/v) glucose and 100 μg/mL ampicillin. Spread out the suspension using a T-shaped spreader until the plate is dry. Incubate the plates at 37  C overnight. 4. On day 2, inoculate a single colony into 10 mL LB-medium containing 100 μg/mL ampicillin (starter culture) and incubate in a 50 mL Erlenmeyer flask at 37  C and 200 rpm overnight. 5. On day 3, transfer 1–5 mL of the starter culture to an Erlenmeyer flask containing 200 mL 2 YT medium supplemented with 100 μg/mL ampicillin. Incubate the expression culture for ~2–3 h at 37  C and 200 rpm until OD600 (optical density) is 0.6–0.8. 6. Add 0.5 mM IPTG to induce protein expression and incubate the culture overnight in a 25  C shaking incubator at 200 rpm.

3.2 Purification of ClyA-AS (Day 4)

1. Harvest the bacterial cells by centrifugation in 50 mL centrifuge tubes at 4000  g for 30 min at 4  C. Discard the supernatant. Store the pellet at 80  C. 2. Freeze the cell pellet for at least 1 h at 80  C. To allow a more thorough cell disruption, the pellet should be frozen at 80  C and thawed two more times. It takes approximately 20 min to be frozen again. 3. Resuspend the pellets in 20 mL of Lysis buffer supplemented with 0.2 units/mL DNase I. Use a vortex to homogenize the solution well. 4. Add 10 μg/mL of lysozyme. Mix well by vortexing and incubate the suspension for 20–30 min at room temperature while shaking. 5. Place the tube containing the cell suspension on ice and disrupt the cells by sonication: 3  30 s at 30% output (see Note 1). 6. Spin down debris by centrifugation for 30 min at 5000  g while cooled at 4  C. The protein is in the supernatant. Transfer the cell lysate to a fresh 50 mL centrifuge tube. 7. During the centrifugation of the previous step, start preparing the Ni-NTA resin. Take 100 μL of Ni-NTA resin (200 μL of


Nicole Ste´phanie Galenkamp et al.

Ni-NTA suspension) into a microtube and wash the resin by adding 500 μL of WB. Mix by inverting the tube and centrifuge the suspension for 30 s at 11,000  g. Carefully remove the supernatant and add 500 μL of fresh WB. Repeat three times. 8. Add the washed Ni-NTA resin to the cell lysate in the centrifuge tube. Incubate for 10–30 min while turning at room temperature (see Note 2). 9. Prewash the empty gravity flow column (1.2 mL bed volume) with approximately 5 bed volumes of WB (~5 mL). 10. Transfer the lysate with the incubated Ni-NTA resin stepwise into the column (see Note 3). 11. Wash the column with at least 50 resin volumes (~5 mL) of WB and remove excess WB by centrifugation of the column in a clean microtube for 30 s (max. 2000  g). 12. Place the column into a clean microtube and elute the ClyA monomers by adding 100 μL of EB. For larger columns, use a volume of EB that covers the resin volume. Incubate for 5–10 min and elute by a quick (~30–60 s) centrifugation spin (max. 2000  g). Repeat the elution step ~3–4 times. Store the monomer solution at 4  C (see Note 4). 3.3 Oligomerization and Gel Purification of ClyA-AS Nanopores (Day 4)

1. Measure the protein concentration using a spectrophotometer or a Bradford assay and dilute the purified protein to 1 mg/mL in EB. 2. Add 0.2% (w/v) DDM and mix well. Incubate for 30 min at 37  C without shaking. 3. Add 5 blue native sample buffer to the purified protein so that the final concentration in the sample is 1 BN SM (dilute down five times). 4. Prepare the blue native PAGE. Assemble it into the electrophoresis cell and fill the container and the top compartment of the gel with 1 native running buffer. Using a pipette, wash the wells of the gel with 1 native running buffer into the wells. Make sure air bubbles are removed. 5. Load 45 μL of sample into each well (depending on the well size). Then, add 2 mL of 20 cathode buffer additive to the top compartment and mix gently (final concentration in top compartment should be 1 cathode buffer additive). Then allow the gel to run for 10 min at 120 V (see Note 5). 6. After 10 min, take out the gel and discard the cathode buffer additive solution (blue solution) from the upper compartment. Wash thoroughly (six times or more) with water in order to remove the blue solution from the wells. 7. Fill the upper compartment with fresh 1 native running buffer and insert the gel back in the container.

Preparation of ClyA Nanopores


Fig. 1 Isolation of different ClyA-AS types separated on 4–20% acrylamide BN-PAGE

8. Allow the gel to run for approximately 1 h at 120 V until the protein bands corresponding to the oligomeric ClyA types are clearly separated (Fig. 1). 9. Using a surgical blade, cut out the bands corresponding to the desired oligomeric state (see Note 6). Store the bands at 20  C. 10. Extract the protein from a smaller piece (1/10th) of the gel by adding 20 μL Extraction buffer. Incubate at room temperature for 1–2 h. Store the gel-extracted ClyA oligomers for 1–2 months at 4  C.


Notes 1. Make sure not to overheat by keeping samples on ice. 2. Incubate for no longer than 45 min. Longer incubation times will cause the ClyA to aggregate. 3. If the column does not flow through well, a tabletop centrifuge can be used to facilitate the flow through. Do not use speeds higher than 2000  g as they might damage the column and centrifuge. 4. ClyA monomers will aggregate when frozen; therefore do not freeze the ClyA monomers. 5. The 20 cathode buffer additive should have entered the gel. 6. Oligomeric ClyA proteins show multiple bands in blue-native polyacrylamide gels (Fig. 1). To extract the 12-mer form of ClyA (Type I), extract the lowest band from the polyacrylamide gel. The band above is corresponding to the 13-mer variant of ClyA (Type II).

References 1. Bezrukov SM, Vodyanoy I, Parsegian VA (1994) Counting polymers moving through a single ion channel. Nature 370:279–281 2. Kasianowicz JJ, Brandin E, Branton D, Deamer DW (1996) Characterization of individual polynucleotide molecules using a

membrane channel. Proc Natl Acad Sci U S A 93:13770–13773 3. Wanunu M, Morrison W, Rabin Y, Grosberg AY, Meller A (2009) Electrostatic focusing of unlabelled DNA into nanoscale pores using a salt gradient. Nat Nanotechnol 5:160–165


Nicole Ste´phanie Galenkamp et al.

4. Meller A, Nivon L, Brandin E, Golovchenko J, Branton D (2000) Rapid nanopore discrimination between single polynucleotide molecules. Proc Natl Acad Sci U S A 97:1079–1084 5. Akeson M, Branton D, Kasianowicz JJ, Brandin E, Deamer DW (1999) Microsecond time-scale discrimination among polycytidylic acid, polyadenylic acid, and polyuridylic acid as homopolymers or as segments within single RNA molecules. Biophys J 77:3227–3233 6. Clarke J, Wu HC, Jayasinghe L, Patel A, Reid S, Bayley H (2009) Continuous base identification for single-molecule nanopore DNA sequencing. Nat Nanotechnol 4:265–270 7. Luchian T, Shin SH, Bayley H (2003) Singlemolecule covalent chemistry with spatially separated reactants. Angew Chem Int Ed Engl 42:3766–3771 8. Movileanu L, Howorka S, Braha O, Bayley H (2000) Detecting protein analytes that modulate transmembrane movement of a polymer chain within a single protein pore. Nat Biotechnol 18:1091–1095 9. Soskine M, Biesemans A, Maglia G (2015) Single-molecule analyte recognition with ClyA nanopores equipped with internal protein adaptors. J Am Chem Soc 137:5793–5797 10. Maglia G, Heron AJ, Stoddart D, Japrung D, Bayley H (2010) Analysis of single nucleic acid molecules with protein nanopores. Methods Enzymol 475:591–623 11. Lin Y, Ying YL, Gao R, Long YT (2018) Single-molecule sensing with nanopore confinement: from chemical reactions to biological interactions. Chemistry 24:13064–13071 12. Ramsay WJ, Bell NAW, Qing Y, Bayley H (2018) Single-molecule observation of the intermediates in a catalytic cycle. J Am Chem Soc 140:17538–17546 13. Soskine M, Biesemans A, Moeyaert B, Cheley S, Bayley H, Maglia G (2012) An engineered ClyA nanopore detects folded target

proteins by selective external association and pore entry. Nano Lett 12:4895–4900 14. Mueller M, Grauschopf U, Maier T, Glockshuber R, Ban N (2009) The structure of a cytolytic alpha-helical toxin pore reveals its assembly mechanism. Nature 459:726–730 15. Franceschini L, Brouns T, Willems K, Carlon E, Maglia G (2016) DNA translocation through nanopores at physiological ionic strengths requires precise nanoscale engineering. ACS Nano 10:8394–8402 16. Soskine M, Biesemans A, De Maeyer M, Maglia G (2013) Tuning the size and properties of ClyA nanopores assisted by directed evolution. J Am Chem Soc 135:13456–13463 17. Van Meervelt V, Soskine M, Singh S, Schuurman-Wolters GK, Wijma HJ, Poolman B, Maglia G (2017) Real-time conformational changes and controlled orientation of native proteins inside a protein nanoreactor. J Am Chem Soc 139:18640–18646 18. Galenkamp NS, Soskine M, Hermans J, Wloka C, Maglia G (2018) Direct electrical quantification of glucose and asparagine from bodily fluids using nanopores. Nat Commun 9:4085 19. Wloka C, Van Meervelt V, van Gelder D, Danda N, Jager N, Williams CP, Maglia G (2017) Label-free and real-time detection of protein ubiquitination with a biological nanopore. ACS Nano 11:4387–4394 20. Van Meervelt V, Soskine M, Maglia G (2014) Detection of two isomeric binding configurations in a protein-aptamer complex with a biological nanopore. ACS Nano 8:12826–12835 21. Willems K, Van Meervelt V, Wloka C, Maglia G (2017) Single-molecule nanopore enzymology. Philos Trans R Soc Lond B Biol Sci 372:1726 22. Miles G, Cheley S, Braha O, Bayley H (2001) The staphylococcal leukocidin bicomponent toxin forms large ionic channels. Biochemistry 40:8514–8522

Chapter 3 Building Synthetic Transmembrane Peptide Pores Kozhinjampara R. Mahendran Abstract Membrane protein pores have demonstrated applications in nanopore technology. Previous studies have mostly focused on β-barrel protein pores, whereas α-helix-based transmembrane protein pores are rarely explored in nanopore applications. Here, we developed a synthetic transmembrane peptide pore built entirely from short synthetic α-helical peptides. We examined the formation of a stable uniform ion-selective pore in single-channel electrical recordings. Furthermore, we show that cyclodextrins (CDs) block the peptide pores and determine the kinetics of CD binding and translocation. We suggest that such designed synthetic transmembrane pores will be useful for several applications in biotechnology, including stochastic sensing. Key words Peptide, Pores, Bilayer, Conductance, Single-channel, Substrates


Introduction Membrane-spanning proteins are found in all major groups of organisms and involved in a wide variety of biological functions, including transport of small molecules across the membrane [1, 2]. They are mostly hydrophobic, usually requiring appropriate detergents during their extraction and purification from the membrane [1, 3]. Often, they fold into their native structures and can exert their functions in specific membrane-mimetic conditions [4– 6]. Notably, membrane proteins are engineered extensively for applications in single-molecule sensing of a wide variety of biomolecules [5, 6]. Previously engineering and redesign of membrane proteins have been predominantly based on β-barrels [3, 5]. For example, heptameric α-hemolysin pore (αHL) derived from Staphylococcus aureus has been engineered extensively for the detection of a wide variety of analytes through stochastic sensing [5, 7]. Most of the natural ion channels consist of α-helical bundles and charge selectivity, which has not been produced with engineered αHL pores [5, 8]. Importantly, alpha-helical pore-forming proteins include a broad range of antimicrobial peptides (AMPs) and

Monifa A. V. Fahie (ed.), Nanopore Technology: Methods and Protocols, Methods in Molecular Biology, vol. 2186,, © Springer Science+Business Media, LLC, part of Springer Nature 2021



Kozhinjampara R. Mahendran

alpha-pore-forming toxins (α-PFTs) with diverse structures and membrane assembly mechanisms [2, 9–11]. However, engineering α-helical bundles from existing scaffolds can cause misfolding of proteins, and therefore α-helical bundles are engineered mainly by de novo design [12–16]. Also, the interaction of alpha-helical peptides with lipid membrane can produce complex structures hindering a stable pore formation [12, 13, 16]. Recently, there is a significant interest in developing and engineering synthetic transmembrane pores that can be used for stochastic sensing of small biomolecules [5, 17]. Specifically, engineering α-helical peptides that can selectively conduct specific ions can play a major role in nanobiotechnology. Notably, a short synthetic peptide based on the membrane-spanning domain of polysaccharide transporter Wza has been previously characterized [13]. This synthetic pore inserts into the membrane to form an active stable pore, but the assembly pathway of the pore consists of several intermediate conductance states [13]. Currently, various approaches are used for the de novo design of an entire protein based on the physical principles of protein folding [18]. In particular, computational methods are used extensively to design possible structures with molecular-level accuracy for building suitable structural models for engineering [15, 19, 20]. We propose that a stable, functional alpha-helical synthetic pore with larger inner diameter can be used for single-molecule sensing of complex biomolecules. Here, we focus on an alpha-helical peptide pore, pPorA, based on short synthetic peptides consisting of 40 amino acids derived from the porin PorACj produced by a Gram-positive bacteria Corynebacterium jeikeium [16, 21].



2.1 Peptide Structure Analysis

1. Pore-forming 40 amino acid pPorA peptides (Peptide Protein Research Ltd. at >95% purity as lyophilized powder). (MIDQITEIFGQLGTFLGGFGNIFKGLKDVIETIVKWTAAK). Solubilize peptides in 10 mM potassium phosphate buffer, pH 7.4 and store at 4  C for 3 months. 2. MOS-500 spectropolarimeters fitted with Peltier temperature controllers (Bio Logic Science Instruments) for Circular Dichroism Analysis. 3. Quartz cuvettes measurements.






4. PBS buffer: 8.2 mM sodium phosphate, 1.8 mM potassium phosphate, 137 mM sodium chloride, 2.7 mM potassium chloride at pH 7.4.

Synthetic Peptide Pores


5. CD buffer: 8.2 mM sodium phosphate, 1.8 mM potassium phosphate, 137 mM sodium chloride, 2.7 mM potassium chloride at pH 7.4 with 1% n-dodecyl β-D-maltoside (DDM). 2.2

Gel Extraction

1. SDS PAGE reagents: 2 Laemmli sample buffer (Bio-Rad), Any kD™ Mini-PROTEAN® TGX™ precast gel (Bio-Rad), Precision Plus Protein™ Dual Color Standards (Bio-Rad). 2. Microfilterfuge tubes.

2.3 Single-Channel Recording

1. Pre-painting solution: Use either 10 mg/mL hexadecane in npentane or 10 mg/mL hexadecane in hexane. This stock solution can be applied directly to Teflon aperture and can be stored at 20  C for 3 months. 2. Bilayer forming lipid solution: 1,2-diphytanoyl-sn-glycero-3phosphocholine (Avanti Polar Lipids) dissolved in pentane (5 mg/mL). This lipid stock solution can be applied directly to bilayer chambers and can be stored at 20  C for at least 6 months. 3. Electrolyte solution: Typically contains 1 M KCl, 10 mM HEPES, pH 7.4. Dithiothreitol (DTT) can be used with electrolyte buffer to facilitate cysteine peptide insertion into the bilayer. 4. Detergent buffer: 0.1% n-dodecyl β-D-maltoside (DDM) solubilized in 10 mM potassium phosphate buffer pH 7.4 is used for directly diluting peptides. This solution can be stored at 20  C for 6 months. 5. Analyte pore blocker: octakis-(6-amino-6-deoxy)-γ-cyclodextrin octahydrochloride (am8γCD, AraChem Cyclodextrin-Shop) solubilized in 1 M KCl electrolyte buffer is used for blocking pores. This solution should be prepared fresh before adding into the bilayer chamber. 6. Bilayer chambers: We use custom-made vertical bilayer chambers made from Teflon or Delrin with a wide variety of shapes and solution volumes ranging from 500 μL to 2 mL. 7. Polytetrafluoroethylene (Teflon) film (Goodfellow, Cambridge) for the bilayer formation. 8. Teflon film of several diameter apertures (30–100 μm) are obtained by punching holes into 25-μm thickness polytetrafluoroethylene (PTFE) Teflon foils by a high-voltage discharge BD10A (Electro-Technic Products, US). 9. Ag/AgCl electrodes in 3 M KCl, 3% agarose bridge. 10. Bilayer chamber cleaning solution: KOH, isopropanol, methanol, and ethanol. 11. Equipment: Axopatch 200B patch-clamp amplifier (Molecular Devices, CA); Digidata 1550B 1440A digitizer with data


Kozhinjampara R. Mahendran

acquisition and analysis program (pClamp 10.6); Faraday cage setup and antivibration table (Holmarc Opto-Mechatronics Pvt. Ltd.) (see Note 1).



3.1 Peptide Synthesis

1. Synthesize peptides using solid-phase peptide synthesis and purify to >95% by reversed-phase high-performance liquid chromatography (HPLC). 2. Confirm their molecular mass by mass spectrometry. Store peptides as lyophilized powders at 20  C. Here, we used 40 amino acid residue peptides, pPorA: MIDQITEIFGQLGTFLGGFG NIFKGLKDVIETIVKWTAAK derived from the membrane porin PorACj of Corynebacterium jeikeium. 3. Design, synthesize, and purify a mutant peptide MIDQ ITEIFGQLGTFLGGFGNIFCGLKDVIETIVKWTAAK.

3.2 Circular Dichroism Spectroscopy

1. Determine the secondary structures of the peptides in different buffer conditions by circular dichroism (CD) spectroscopy using MOS-500 spectropolarimeters fitted with Peltier temperature controllers. 2. Prepare peptide samples as 5 and 50 μM solutions in CD buffer. Measure CD spectra of peptides only in PBS buffer as a control to confirm the role of detergent in the proper folding of alphahelical peptides. Record spectra using a 1 mm path length quartz cuvette at 20  C. 3. For each dataset (in deg.), subtract baselines from the same buffer and cuvette, and then normalize data points for amide bonds, concentration of peptide and path length of the cuvette to obtain mean residue ellipticity (MRE; deg. cm2/dmol/res) (see Note 2). CD spectra should confirm the secondary structures of wild-type and mutant pPorA, which exist in α-helical conformation in 1% n-dodecyl β-D-maltoside (DDM) micelles (Fig. 1a, b).

3.3 SDS PAGE Gel Extraction of pPorA Oligomers

SDS-polyacrylamide gel electrophoresis (SDS-PAGE) is used to assess the oligomeric state of DDM solubilized mutant pPorA peptides. 1. Solubilize the purified peptides in detergent buffer and prepare in Laemmli SDS-sample buffer. Load onto an Any kD MiniPROTEAN® TGX™ precast gel with a Protein Standard Marker and perform electrophoresis until dye front is at the bottom of gel (see Notes 3 and 4).

Synthetic Peptide Pores


Fig. 1 CD spectra of pPorA peptide pores. (a) CD spectra at 20  C for wild-type pPorA in PBS with 1% DDM at 5 μM (blue) and 50 μM (red) peptide concentration. (b) CD spectra at 20  C for cysteine mutant pPorA in PBS with 1% DDM at 5 μM (blue) and 50 μM (red) peptide concentration

2. Visualize the proteins by staining the gel with standard Coomassie technique. Determine the electrophoretic mobility and molecular mass estimate of peptides in SDS-PAGE based on the protein standards. The pPorA mutant peptides produce three bands on SDS-PAGE gel, a lower ~4.5 kDa band corresponding to the monomeric unit, ~9 kDa band corresponding to the dimeric unit, and a ~35 kDa band corresponding to a possible octameric pore structure (Fig. 2a). 3. Cut the bands containing corresponding peptide oligomers from the gel, crush the gel slices, and incubate each separately in 500 μL phosphate buffer for 45 min at room temperature or longer. 4. Obtain the cleared protein filtrate by using a microfilterfuge tube centrifuged at 15,000  g for 5 min. pPorA oligomer is ready to use in single-channel electrical recording. 3.4 Single-Channel Electrical Recordings

1. Set up the single-channel electrical recording apparatus with the following parameters: Apply a low-pass filter frequency of 10 kHz and a sampling frequency of 50 kHz. Optional-Filter the electrical signal at 2 kHz using an 8-pole Bessel digital filter based on the analysis requirement. 2. Perform single-channel recordings using planar lipid bilayers of 1,2-diphytanoyl-sn-glycero-3-phosphocholine (see Note 5) by the Montal and Muller technique across a polytetrafluoroethylene (Teflon) film aperture (25-μm thick, ~80 μm in diameter) [22] (see Note 6). 3. Coat each side of the Teflon film containing the aperture with 1–2 μL pre-painting solution using a simple stereo microscope (see Note 7).


Kozhinjampara R. Mahendran

Fig. 2 Electrical properties of gel extracted cysteine mutant pPorA peptide pores. (a) The mutant pPorA peptides run on the SDS-PAGE. The red circle indicates the self-assembled mutant pPorA peptide oligomers. (b) Electrical recording showing single gel extracted mutant pPorA pores at +50 mV and (c) 50 mV. The corresponding current-amplitude histogram is shown. (d) I–V curve obtained from a single gel extracted mutant pPorA pores. (e) Gating of gel extracted single mutant pPorA pores at +200 mV. (f) Interaction of gel extracted single mutant pPorA pores with am8γCD (1 μM, trans) at +50 mV. The current signals were filtered at 10 kHz and sampled at 50 kHz. Electrolyte: 1 M KCl, 10 mM HEPES, pH 7.4

4. Fill cis and trans compartments of the bilayer chamber with 500 μL of electrolyte buffer. 5. Add 5 μL bilayer forming lipid solution into both chambers. 6. Prepare the monolayer in each chamber by lowering the electrolyte level below the aperture and raising it again. Connect the cis compartment to the grounded electrode while connect the trans compartment to the live electrode using Ag/AgCl wires set in 3% agarose containing 3.0 M KCl (see Note 8). 7. Apply a potential difference across the bilayer and test the stability of the bilayer at different voltages ranging from 50 to 200 mV (see Notes 9 and 10). 8. Mix in purified peptides solubilized in 0.1% DDM to the cis side of the chamber (see Notes 11 and 12). Apply a voltage of +200 mV without breaking the bilayer. 9. Monitor pore formation as a stepwise increase in the conductance of the electrical reading. Analyze the electrical recording data with the latest version of pClamp software. Reconstitution

Synthetic Peptide Pores


of pore-forming peptides into planar lipid bilayer under an applied potential allows for measuring an ion conductance that provides an approximate pore size, charge selectivity, or functional properties such as channel opening and closing or substrate blocking. 10. A description of the electrical characteristics of the pPorA pore is as follows: 11. The oligomer ~35 kDa band from the gel rapidly inserts into the bilayer forming a stable pore of single-channel conductance of 4.0  0.2 nS at 50 mV (Fig. 2b, c). 12. The current–voltage (I–V) curve shows higher conductance at negative potential compared to positive potential (Fig. 2d). 13. The gating of the peptide pore is only observed at voltages higher than 60 mV (Fig. 2e). 14. The oligomeric pore can be blocked with am8γCD (Fig. 2f). The addition of 1 μM am8γCD to the trans side results in the ion current blockages only at positive potential establishing a strong electrostatic interaction between the am8γCD and the peptide pore. 3.5 Determination of Single-Channel Conductance of pPorA Peptide Pores

1. Obtain multiple insertions of at least 100 pPorA or pPorA mutant oligomers in single-channel recording by adding high concentrations of peptide pores to the cis chamber provided this do not disrupt the bilayer stability (Fig. 3a). 2. Apply a fixed voltage as multiple pores insert into the bilayer. Re-form the bilayer if it breaks to allow more pores to insert. 3. Determine the current for each pore by the stepwise increase in current. Generate a histogram and fit with Gaussian to determine the average unitary conductance that reveals the uniformity of the pores (Fig. 3b–d). A typical pore has a singlechannel unitary conductance (G) of 4.0  0.2 nS at +25 and +50 mV (n ¼ 100). The pore shows slight asymmetry in the single-channel conductance, i.e., rectification at the positive applied voltage or higher conductance obtained at negative voltages than positive voltages (Fig. 3c, d). 4. Acquire the current–voltage (I–V) curve for one single peptide pore at a time by applying different voltages ranging from 100 to +100 mV (Fig. 4a).

3.6 Charge Selectivity

1. Acquire ion selectivity measurements using asymmetric salt conditions in planar lipid bilayers. Add 1.0 M KCl in the cis side and 0.15 M KCl in the trans side of the bilayer compartment.


Kozhinjampara R. Mahendran

Fig. 3 Electrical properties of cysteine mutant pPorA peptide pores. (a) Electrical recording of multiple insertions of mutant pPorA pores into a planar bilayer at +25 mV. (b) Single mutant pPorA pores insertion at +50 mV. (c) Electrical recording of single mutant pPorA pores at +50 mV. (d) Electrical recording of single mutant pPorA pores at 50 mV. The corresponding current amplitude histogram fitted with Gaussian is shown as an inset. The current signals were filtered at 2 kHz and sampled at 10 kHz. Electrolyte: 1 M KCl, 10 mM HEPES, pH 7.4

2. Add peptides to the cis side of the bilayer chamber. Here, the peptide pore formation produces a particular ion current in the absence of an applied transmembrane voltage. 3. Manually set the ionic current to zero by fine-tuning the applied voltage. The voltage needed to produce zero current is termed as the “reverse potential” (Vm), which is used to estimate the permeability ratio of K+ and Cl ions across the pore by Goldman-Hodgkin-Katz equation [23]. ! cis P K þ ½K þ  þ P Cl ½Cl trans RT ln Vm ¼ trans F P K þ ½K þ  þ P Cl ½Cl cis 4. In this equation, R is the universal gas constant (8.314 J/K/ mol), T is the temperature in Kelvin (K ¼  C + 273.15), F is the Faraday’s constant (96,485 C/mol), P K þ is the relative

Synthetic Peptide Pores


Fig. 4 Single-channel electrical properties of cysteine mutant pPorA peptide pores. (a) Current–voltage (I–V) curve obtained from single mutant pPorA pore. (b) Reverse potential derived from the I–V curve of single mutant pPorA pore. Electrolyte: 1 M KCl, 10 mM MES, pH 6.0, cis/0.15 M KCl, 10 mM MES, pH 6.0, trans. (c) Electrical recording of single mutant pPorA showing gating at +100 mV. (d) Electrical recording of single mutant pPorA showing rapid gating at +150 mV. The current signals were filtered at 2 kHz and sampled at 10 kHz. Electrolyte: 1 M KCl, 10 mM HEPES, pH 7.4

membrane permeability for K+, P Cl is the relative membrane permeability for Cl, [K+]cis is the concentration of K+ ion in the cis side, [K+]trans is the concentration of K+ in the trans side, [Cl]cis is the concentration of Cl in the cis side, and [Cl]trans is the concentration of Cl in the trans side. Ion-selectivity studies reveal that the mutant pPorA pore is cation-selective, with a permeability ratio of 10:1 for K+ and Cl ions (Fig. 4b).


Kozhinjampara R. Mahendran

3.7 VoltageDependent Gating Analysis of pPorA Pores and SubstrateBlocking Events 3.7.1 Voltage-Dependent Gating

3.7.2 Substrate Blocking

1. Record the ionic behavior of single pPorA pores at various voltages for extended periods of time, e.g., 10 min for each voltages from 10 to 200 mV (see Note 13). 2. Analyze the ionic current (open and closed states) in pClamp analysis software. The single-channel recordings should reveal that the peptide pore is in a non-gating state at voltages lower than 50 mV. At higher voltages above 60 mV, the pore fluctuated between open and closed conductance state (gating) with more frequent gating events at the higher voltages (Fig. 4c, d). 1. Acquire the ionic behavior of a single pPorA peptide at different substrate concentrations and applied voltages, recording for 10 min each substrate concentration/voltage condition (see Notes 14 and 15). 2. Quantify substrate blocking using a statistical analysis of a single pore in its blocked and unblocked states. Count the number of blockage events and the blockage time with a single-channel search in pClamp analysis software. Obtain the average blockage time or dwell times by plotting a histogram that is fit to a standard exponential distribution. 3. Calculate the average residence time of substrate closure (τc) and the reciprocal of (τc) gives the dissociation rate of the substrate. The time between successive substrate blockages will give (τo), and the 1/[(τo)]  C, (where C is the concentration of the substrate) provides the association rate of the substrate.

3.8 Cleanup After Each Single-Channel Recording Experiment

1. An efficient washing procedure is essential for cleaning the bilayer chambers after each use. 2. Wash the bilayer chamber after experiments with large amounts of distilled water, followed by isopropanol and methanol. 3. Dry the chambers thoroughly under a stream of nitrogen to evaporate organic solvents. 4. In the case of gel extracted peptides, use 3 M KOH and ethanol washes to remove any residual peptides from the chip and Teflon.


Notes 1. An acoustic screening (Faraday cage) of the setup, in combination with an antivibration table, will allow us to measure ion conductance with better time resolution. 2. The CD spectra of peptides only in PBS buffers show that peptides exist as insoluble aggregates in the buffer.

Synthetic Peptide Pores


Importantly, the peptides are folded in an alpha-helical conformation only in the presence of the detergent. 3. Remove the seal from the precast gel before performing gel electrophoresis. If the seal is not removed, proteins cannot run through the gel efficiently. 4. Load the peptides on Any kD Mini-PROTEAN® TGX™ precast gel (Bio-Rad) with Protein Standard Marker (Bio-Rad) for resolving the monomer bands precisely. We used Plus Protein Dual Color Standards for molecular weight estimation due to increased brightness makes it easier to identify peptides on the gel in real time. This allows the rapid extraction of the peptides from the gel without staining the gel. 5. The most widely used lipid for planar lipid membranes is DPhPC as it has ideal stable bilayer-forming properties. Other lipids such as α-lecithin result in the formation of unstable leaky bilayers not suitable for peptide pore reconstitution. 6. The lipid bilayer is a perfect insulator compared to a bilayer in the presence of a pore that allows the detection of pore conductance and selectivity. The most stable lipid bilayers are achieved by using a Teflon aperture of small diameter in the range of ~30 μm, which allows longer recordings for 6–8 h but requires more patience during protein reconstitution. Also, better time resolutions can be achieved using smaller membranes. 7. The Teflon apertures should be viewed using a light microscope for accurate pre-painting of the hole, which is critical for the formation of the stable bilayers. A small amount of hexadecane is used because the Teflon film aperture should be free of excess hexadecane so that an electrical current can be recorded between the bilayer compartments before the formation of the bilayer. 8. The use of a 3 M KCl/agarose bridge between an electrode is critical for the activity of mutant cysteine peptides and ion-selectivity experiments. The 3 M KCl agarose bridge reduces the access resistance and prevents Ag+ ions from the electrode leaching into the electrolyte solution. Also, double distilled water should be used to prepare all aqueous buffers including KCl (electrolyte) which should be filtered through a 0.4-μm membrane. 9. By measuring the capacitance of the membrane we can conclude on the stability of the bilayer. 10. In the complete absence of protein pores, the planar lipid membrane itself can display pore-like events. Therefore, it is crucial to form DPhPC planar lipid bilayers and record the


Kozhinjampara R. Mahendran

current through lipid bilayers at high voltages up to 200 mV for 5 min to confirm the stability of lipid bilayers before peptide reconstitution. 11. A low concentration of gel extracted peptides should be added for the insertion into the lipid bilayer. The addition of a high concentration of gel extracted peptides can cause unstable lipid bilayers, specifically at higher voltages. Depending on the concentration of the peptide stock, the first insertion will occur after a few minutes. 12. In the case of cysteine peptides, add ~1 μM DTT to facilitate rapid pore insertion into the lipid bilayer. 13. Perfuse the bilayer compartment with electrolyte buffer to avoid numerous insertions of peptides and maintain a single channel for single-molecule sensing. 14. Usually, cationic substrates, anionic substrates, and neutral substrates are used to block the pores formed by the peptides. In the case of charged substrates, blocking with the pore depends on the external parameters such as the side of the addition, salt concentration, polarity, and magnitude of the applied voltage. For example, addition to cationic substrates to the cis side results in ion current blockages only at negative voltages as substrates are electrophoretically pulled into the pore to interact with negatively charged residues in the pore lumen. 15. Notably, we examined the interaction of cation-selective mutant pPorA pores with cationic CDs. For example, cationic cyclic octasaccharide (am8γCD) interacts with the pore in a voltage-dependent manner. Addition of 1 μM am8γCD to the cis side of the bilayer compartment resulted in time-resolved ion current blockages only at negative voltages indicting electrophoretic pulling of CDs into the pore (Fig. 5a, c). When the polarity of the voltage reversed to positive potential, no ion current blockages were observed confirming strong electrostatic interaction between am8γCD and the pore (Fig. 5b, d).

Synthetic Peptide Pores


Fig. 5 Interaction of cysteine mutant pPorA peptide pores with cationic cyclodextrins. (a) Typical ion current recordings showing the interaction of single mutant pPorA with am8γCD (1 μM, cis) at 50 mV. (b) No ion current blockages were observed at +50 mV. (c) Typical ion current recordings showing the interaction of single mutant pPorA with am8γCD (1 μM, trans) at +50 mV. (d) No ion current blockages were observed at 50 mV. The current signals were filtered at 10 kHz and sampled at 50 kHz. Electrolyte: 1 M KCl, 10 mM HEPES, pH 7.4

References 1. von Heijne G (2006) Membrane-protein topology. Nat Rev Mol Cell Biol 7:909–918 2. Dal Peraro M, Van der Goot FG (2016) Poreforming toxins: ancient, but never really out of fashion. Nat Rev Microbiol 14:77–92 3. Bayley H, Jayasinghe L (2004) Functional engineered channels and pores (review). Mol Membr Biol 21:209–220 4. Bayley H (2009) Membrane-protein structure: piercing insights. Nature 459:651–652 5. Ayub M, Bayley H (2016) Engineered transmembrane pores. Curr Opin Chem Biol 34:117–126

6. Kasianowicz JJ, Balijepalli AK, Ettedgui J, Forstater JH, Wang H, Zhang H, Robertson JW (2016) Analytical applications for poreforming proteins. Biochim Biophys Acta 1858:593–606 7. Bayley H (2015) Nanopore sequencing: from imagination to reality. Clin Chem 61:25–31 8. Tajkhorshid E, Nollert P, Jensen MO, Miercke LJ, O’Connell J, Stroud RM, Schulten K (2002) Control of the selectivity of the aquaporin water channel family by global orientational tuning. Science 296:525–530


Kozhinjampara R. Mahendran

9. Lear JD, Wasserman ZR, DeGrado WF (1988) Synthetic amphiphilic peptide models for protein ion channels. Science 240:1177–1181 10. Brogden KA (2005) Antimicrobial peptides: pore formers or metabolic inhibitors in bacteria? Nat Rev Microbiol 3:238–250 11. Puthumadathil N, Jayasree P, Santhosh Kumar K, Nampoothiri KM, Bajaj H, Mahendran KR (2019) Detecting the structural assembly pathway of human antimicrobial peptide pores at single-channel level. Biomater Sci 7:3226–3237 12. Woolfson DN (2017) Coiled-coil design: updated and upgraded. Subcell Biochem 82:35–61 13. Mahendran KR, Niitsu A, Kong L, Thomson AR, Sessions RB, Woolfson DN, Bayley H (2017) A monodisperse transmembrane alpha-helical peptide barrel. Nat Chem 9:411–419 14. Woolfson DN, Bartlett GJ, Burton AJ, Heal JW, Niitsu A, Thomson AR, Wood CW (2015) De novo protein design: how do we expand into the universe of possible protein structures? Curr Opin Struct Biol 33:16–26 15. Joh NH, Wang T, Bhate MP, Acharya R, Wu Y, Grabe M, Hong M, Grigoryan G, DeGrado WF (2014) De novo design of a transmembrane Zn2+-transporting four-helix bundle. Science 346:1520–1524 16. Krishnan RS, Satheesan R, Puthumadathil N, Kumar KS, Jayasree P, Mahendran KR (2019) Autonomously assembled synthetic transmembrane peptide pore. J Am Chem Soc 141:2949–2959

17. Wang S, Zhao Z, Haque F, Guo P (2018) Engineering of protein nanopores for sequencing, chemical or protein sensing and disease diagnosis. Curr Opin Biotechnol 51:80–89 18. Huang PS, Feldmeier K, Parmeggiani F, Velasco DAF, Hocker B, Baker D (2016) De novo design of a four-fold symmetric TIM-barrel protein with atomic-level accuracy. Nat Chem Biol 12:29–34 19. Thomson AR, Wood CW, Burton AJ, Bartlett GJ, Sessions RB, Brady RL, Woolfson DN (2014) Computational design of water-soluble alpha-helical barrels. Science 346:485–488 20. Lu P, Min D, DiMaio F, Wei KY, Vahey MD, Boyken SE, Chen Z, Fallas JA, Ueda G, Sheffler W, Mulligan VK, Xu W, Bowie JU, Baker D (2018) Accurate computational design of multipass transmembrane proteins. Science 359:1042–1046 21. Abdali N, Barth E, Norouzy A, Schulz R, Nau WM, Kleinekathofer U, Tauch A, Benz R (2013) Corynebacterium jeikeium jk0268 constitutes for the 40 amino acid long PorACj, which forms a homooligomeric and anionselective cell wall channel. PLoS One 8:e75651 22. Gutsmann T, Heimburg T, Keyser U, Mahendran KR, Winterhalter M (2015) Protein reconstitution into freestanding planar lipid membranes for electrophysiological characterization. Nat Protoc 10:188–198 23. Benz R, Schmid A, Hancock RE (1985) Ion selectivity of gram-negative bacterial porins. J Bacteriol 162:722–727

Chapter 4 Design and Assembly of Membrane-Spanning DNA Nanopores Kerstin Go¨pfrich, Alexander Ohmann, and Ulrich F. Keyser Abstract Versatile lipid membrane-inserting nanopores have been made by functionalizing DNA nanostructures with hydrophobic tags. Here, we outline design and considerations to obtain DNA nanopores with the desired dimensions and conductance properties. We further provide guidance on their reconstitution into lipid membranes. Key words DNA nanotechnology, DNA origami, Nanopores, Ionic conductance, Lipid membrane, Synthetic ion-channels


Introduction Protein pores traverse the membranes of virtually all living cells, mediating vital processes like signal transduction or substrate exchange. Yet these pores featuring nanoscale dimensions have also found technological applications—from antimicrobial agents [1] to synthetic cell assembly [2] and, most prominently, label-free single molecule detection with notable success in DNA sequencing [3]. Beyond the protein pores found in nature, synthetic membrane-spanning nanopores have been tailored for specific applications [4]. While peptide pores are a prominent example, recent advances in structural DNA nanotechnology led to the creation of diverse DNA-based membrane-spanning pores (Fig. 1). Typically, hydrophobic tags are carefully positioned and covalently liked to the DNA construct. They help overcome the energy barrier for the insertion of the hydrophilic DNA into the hydrophobic lipid bilayer. Remarkably, DNA nanopore designs span three orders of magnitude in conductance and molecular weight. Voltage [5] and ligand [6] gating has been demonstrated and DNA pores have been employed for label-free detection of translocating DNA [7]. With the large toolbox for chemical

Monifa A. V. Fahie (ed.), Nanopore Technology: Methods and Protocols, Methods in Molecular Biology, vol. 2186,, © Springer Science+Business Media, LLC, part of Springer Nature 2021



Kerstin Go¨pfrich et al.

Fig. 1 Membrane-spanning DNA nanopores. (a) Schematic illustration of diverse membrane-spanning DNA nanopores (literature references from left to right: [11, 12, 14, 15]). Adapted from [8]. (b) TEM micrograph of a lipid vesicle with multiple DNA origami nanopores as pioneered by Langecker et al. Adapted from [13]. Reprinted with permission

functionalization of DNA, membrane-spanning DNA nanopores hold great potential for the implementation of versatile functions [8]. Here, we give step-by-step instruction on the design and synthesis of a simple DNA nanopore. We further give guidance and discussion of a more complex DNA structure design.



2.1 DNA Nanopore Assembly

The following materials are required to assemble a simple DNA nanopore as presented and characterized in [9, 10]. It consists of four interconnected DNA duplexes and is anchored in the lipid membrane via four cholesterol tags as shown in Fig. 2. Prepare all solutions using ultrapure water at room temperature. 1. Custom-made unmodified DNA oligonucleotides (standard desalting purification). 2. Custom-made 30 fluorophore-modified DNA oligonucleotides (HPLC purification). 3. Custom-made 30 cholesterol-tagged oligonucleotides. An example list of DNA sequences previously used for the assembly of a four-helix bundle [9] is given in Table 1. 4. TE buffer: 10 mM Tris–HCl, pH 8, 1 mM EDTA. 5. Folding buffer: 10 mM Tris–HCl, 1 mM EDTA, 20 mM MgCl2, pH 8.0. 6. Microtubes (0.2, 0.6, 1.5 mL). 7. PCR thermocycler.

DNA Nanopore Design


Fig. 2 A simple DNA nanopore; (a) side view and (b) top view of a simple DNA nanopore consisting of four interconnected DNA duplexes. The four cholesterol modifications for membrane anchoring are shielded by six nucleotide long single-stranded DNA overhangs to avoid aggregation of the structure. Adapted from [9]

Table 1 DNA sequences employed for Cy3-labeled DNA nanostructures assembled with four cholesterol modifications shielded by six nucleotide long overhangs (orange) Adapted from [9] DNA strand

Sequence (5' to 3')
















2.2 Attachment to Giant Unilamellar Vesicles (GUVs)


1. Unlabeled lipids: 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) and 1-palmitoyl-2-oleoyl-glycero-3-phosphocholine (POPC), delivered in chloroform at a stock concentration of 25 mg/mL (Avanti Polar Lipids). 2. GUVs prepared by electroformation with a lipid composition of 50 mol% DOPC, 50 mol% POPC. 3. 300 mM sucrose solution to prepare GUVs. 4. Fluorescent lipid: Atto488-PE (from Atto-TEC), delivered in chloroform at a stock concentration of 1 mg/mL. This lipid is supplemented at 0.5 mol% in GUVs for visualization purposes. 5. Chloroform. 6. Two indium tin oxide (ITO)-coated glass slides (Nanion Technologies GmbH).


Kerstin Go¨pfrich et al.

7. Rubber ring as a spacer. 8. Cleaning reagents: acetone, isopropanol, 70% ethanol. 9. DNA nanostructures. 10. GUV buffer: 160 mM glucose, 10 mM Tris–HCl, 1 mM EDTA, 20 mM MgCl2, pH 8.0. 11. Bovine Serum Albumin (BSA)-coated cover slides for the observation chamber (to prevent fusion of the GUVs with the glass surface). 12. Observation chamber (from ibidi) or assembled from a microscopy glass slide and a cover slide spaced with, e.g., doublesided sticky tape. 13. Confocal laser scanning or epifluorescence microscope for visualization. 14. Electroformation unit Technologies GmbH).





15. Dessicator.



3.1 DNA Nanopore Assembly 3.1.1 Oligonucleotide Preparation

1. Dissolve lyophilized unmodified DNA oligonucleotides (sc1, sc3) and fluorophore-tagged oligonucleotides (st2, st4) to 100 μM in TE buffer according to manufacturer’s instructions. Aliquot into several tubes. This solution can then be stored at 20  C for long-term storage. 2. Dissolve lyophilized cholesterol-tagged DNA oligonucleotides (sc2, sc4, st1, st3) to 100 μM in ddH2O according to manufacturer’s instructions. Aliquot (e.g., 10 μL per aliquot) and store at 20  C.

3.1.2 DNA Nanopore Assembly

3. Thaw one aliquot of each of the eight of the unmodified, fluorophore-modified and cholesterol-modified oligonucleotides from Table 1. 4. In particular, heat the four aliquots containing cholesteroltagged oligonucleotides to 60  C for 5 min on a hotplate or in the PCR thermocycler while the other four oligonucleotides can be thawed at room temperature. Vortex the cholesterolmodified oligonucleotides directly after heating and pipette quickly to avoid aggregation. 5. Mix the eight DNA oligonucleotides, no preferential order, from Table 1 to a final concentration of 1 μM in folding buffer in a microtube (see Note 1). For example, mix together 1 μL of each of the eight oligonucleotides (stock at 100 μM) and then add 92 μL of folding buffer.

DNA Nanopore Design


6. Place the microtube containing the DNA mixture into a PCR thermocycler. Heat to 85  C for 5 min, subsequently gradually cool to 25  C over 18 h. Store assembled DNA nanopores at 4  C or use immediately in a downstream assay. 3.2 Preparation of Giant Unilamellar Vesicles (GUVs) by Electroformation

1. Mix DOPC and POPC lipids (dissolved in chloroform) in a 1:1 mole ratio and add Atto488 PE. Add chloroform to obtain a 10 mM lipid mix. Seal all lipid vials with Teflon film to prevent evaporation and oxidation of the lipids. This lipid mix can be stored at 20  C and reused for future electroformation preparations. 2. Clean ITO slides by sonicating for 5–10 min in acetone, then in isopropanol and lastly in ethanol. 3. Pipet 20 μL of the lipid mixture on the conducting side of each ITO slide (as determined with a voltmeter). 4. Use a clean glass coverslip to spread the lipid mix. Spreading should look even and reflect light with green colors. 5. Place in desiccator for at least 10 min (better 1–2 h) to evaporate the chloroform and dry the lipid mix. 6. Press a rubber ring onto one of the slides, enclosing the area where the lipid was spread. 7. Place this cover slide in the electroformation unit and fill entirely with 300 mM sucrose. 8. Put the second slide on top of the rubber ring so that the two lipid-coated sides are facing one another with the rubber ring in between. 9. Apply an AC-current of 3VP-P and a frequency of 5 Hz for 2 h. This may vary if you are using a different lipid mixture. 10. Remove the top glass slide and pipet up the vesicles (dispersed in the sucrose solution) with a large-diameter pipette tip immediately after the 2 h incubation is complete and store at 4  C (see Note 2).

3.3 Attachment of DNA Nanopore to Giant Unilamellar Vesicles (GUVs)

1. Dilute the DNA nanopores to a concentration of 200 nM (1:5 dilution of the initial 1 μM stock). Use the GUV buffer for the dilution in order to match the osmolarity of the GUVs. Example: GUVs were 300 mM sucrose (hence 300 mOsm). The folding buffer (10 mM Tris–HCl, 1 mM EDTA, 20 mM MgCl2) has an osmolarity of approx. 70 mOsm. Therefore the 1:5 dilution should be prepared by mixing the DNA nanopores with 358 mM sucrose to match the osmolarity of the GUVs. This gives an osmolarity of 300 mOsm for the diluted DNA nanostructures (see Note 3).


Kerstin Go¨pfrich et al.

Fig. 3 Visualization of DNA nanopore membrane insertion. GUV imaged in bright field (left) and fluorescence mode, excited at 514 nm (middle and right). At 0 s the DNA nanostructures are added to GUV suspension and are not yet bound. At 60 s after addition of the DNA nanostructures, a bright ring forms around the vesicle, indicating rapid folding and membrane attachment of the DNA nanostructures. Adapted from [8]. Reprinted with permission

2. Add the osmolarity-matched DNA nanopore solution to the GUV suspension at a ratio of 1:1 (v/v). Membrane attachment happens within seconds (see Note 4). 3. Transfer the DNA nanopore-GUV mixture to a BSA-coated observation chamber. 4. Use a confocal laser scanning microscope or an epifluorescence microscope at an excitation of 514 nm to visualize the attachment of the Cy3-labeled DNA nanopores to the lipid membrane of the GUVs. The fluorescence from the nanopores should be visible as a fluorescent ring in the confocal plane (Fig. 3, right panel). The nanopore fluorescence (Cy3 fluorescence ¼ Ex max: 550 nm and Em max: 570 nm) should overlay with the fluorescence of the lipid membrane (LissRhod PE ¼ Ex max: 545 and Em max: 567) (see Note 5). 5. As a control experiment, prepare DNA nanopores without the cholesterol modifications. This construct should remain in the bulk solution around the GUVs but not embedded in the lipid membrane. 3.4 Guidelines for DNA Nanopore Design

For more advanced applications, design and customize your own DNA nanopore with the desired features according to the protocol outlined below. 1. Choose the required pore size. The pore size will depend on the envisioned application of the DNA nanopore: For nanopore sensing or to create a transmembrane passage for specific molecules, keep in mind that the pore diameter must exceed the size of the analyte. Note that even if this criterion is met, large highly charged analytes may still be excluded from the pore due to the negative charge of the DNA backbone (see Note 6). Small pores, on the other hand, are preferable to engineer

DNA Nanopore Design


selectivity and to mimic protein ion channels. The nominal inner pore diameters of existing DNA nanopores ranges from sub-nanometer [11] to 6 nm [12], spanning three orders of magnitude in ionic conductance (from tens of pS to tens of nS). 2. Derive membrane anchoring strategy. Hydrophobic modifications, covalently linked to the DNA, facilitate the insertion of DNA nanopores into lipid membranes (see Note 7). Cholesterol [9, 11–13], tocopherol [7], and ethane [14] are all DNA modifications commercially available. Alternatives such as inhouse-synthesized porphyrin [15, 16] have been successfully used as hydrophobic tags for lipid membrane anchoring. Alternatively, biotin-functionalized DNA nanopores have been designed to bind to biotinylated lipids via biotin-streptavidin linkage [7] (also commercially available). For the design, it is important to note that the anchoring strategy can also influence the ion conductance of the pore. Experiments and molecular dynamics simulations have shown that the lipid head groups tilt toward the membrane-inserted DNA nanostructure (see Fig. 4a). In this configuration, the hydrophobic lipid tails are shielded from the hydrophilic DNA. This leads to a toroidal pore at the DNA-lipid interface. Ions can flow though this toroidal lipid pore as well as through the central cavity of the DNA nanopore, increasing the overall conductance (Fig. 4a, red arrows). The emergence of a continuous lipid bilayer leaflet allows for lipid molecules to diffuse from one bilayer sheet to the other (Fig. 4a in blue). For this reason, membraneinserted DNA nanostructures have also been shown to act as scramblases [17]. To confine the ion pathway to the central

Fig. 4 Computer-aided modeling of DNA nanopores. (a) Schematic illustration of a DNA nanopore with hydrophilic exterior. Ions can pass through the central cavity as well as though the toroidal lipid pore forming at the DNA-lipid interface [15]. Lipids can be transferred from one bilayer leaflet to the other, leading to scramblase activity of DNA nanopores [17]. (b) Coarse grained simulation of the free energy of DNA nanopore insertion as a function of the pore radius and the number of hydrophobic tags (here: cholesterol). Adapted from [12]. Reprinted with permission. Copyright 2016, American Chemical Society. This direct link is supposed to be included:


Kerstin Go¨pfrich et al.

cavity and to inhibit scrambling, remove the charges of the DNA backbone in the membrane-spanning section of the pore by using ethane modifications [14], C12 spacers or uncharged nucleic acids (e.g., PNA) in the stem section of the designed DNA pore. 3. Determine the required number of membrane anchors. To trigger self-insertion of the DNA nanopores, the energy cost for pore formation in the lipid membrane has to be compensated by the energy gain for the insertion of the hydrophobic tags or the formation of other molecular bonds. Therefore, it is crucial to choose the right number of hydrophobic tags. Referencing the coarse-grained simulations in Fig. 4b, one can extract the required number of cholesterol tags for a certain pore radius (the blue region of the plot represents a net energy gain for pore insertion) (see Note 8). 4. Choose between scaffolded DNA origami or scaffold-free DNA tiles. This will depend on the application of the DNA nanopore. Assembly of nanopores from short synthetic single strands of DNA is useful when the target structure is small. Assembly from single strands, however, becomes less efficient for large finite-size assemblies (12–17% yield [18]), which is why scaffolded DNA origami is the method of choice for larger nanopores. When using a kilobase long DNA scaffold (e.g., the commonly used DNA from the virophage M13mp18), the size of the DNA nanostructure falls into the megadalton regime. However, the achievable concentrations are typically restricted to nanomolar amounts. Table 2 compares advantages

Table 2 Comparison of scaffolded DNA origami nanopores and scaffold-free DNA nanopores Scaffolded DNA origami nanopores

Scaffold-free DNA nanopores

Molecular weight

MDa range

kDa range

Number of DNA strands

1 scaffold, typically 200–350 synthetic DNA oligonucleotides

Typically 2–20 synthetic DNA oligonucleotides

Achievable concentrations

Typically 1–100 nM

Typically 1–10 μM


Large number of possible attachment sites on staples, typically no modifications on scaffold

Can be placed on all strands, however limited number


Thermal annealing for hours to days

Thermal annealing or RT assembly within minutes


Required (e.g., spin filtration)

Mostly not required


[7, 12, 13]

[5, 6, 9, 10, 11, 14–16]

DNA Nanopore Design


and disadvantages of both DNA assembly methods with corresponding literature references. 5. Conceive target shape. The target shape of the nanopore will depend on the chosen pore diameter. The pore cavity is typically lined by a single layer of DNA, although multiple layers may improve the mechanical stability. The transmembrane section of the pore should be at least 5 nm long to fully span the lipid bilayer. For a scaffold-free design, the transmembrane section is the only required part. For scaffolded DNA origami, the remaining scaffold has to be used up. This can be achieved by extending the pore length far beyond the thickness of the bilayer up to 40 nm [12, 12], and/or by implementing a base plate, which lies flat on the membrane and provides space for the attachment of hydrophobic groups [7]. Such an asymmetric mushroom-shaped pore with a large hydrophilic head will always insert in the same orientation. To implement the envisioned design, it is useful to employ computer-aided design tools like caDNAno (see Note 9). While caDNAno has been developed for scaffolded DNA origami, the scaffold path can be broken up into short strands if the target structure is scaffold-free. The same design principles apply as for DNA nanostructures in general. A useful reference with guidance is given elsewhere [19]. Once the target shape is drawn, the design-file can be submitted to CanDo (see Note 10) for structural and fluctuation analysis. Fluctuations can indicate areas of weak structural stability, which can be stabilized by additional crossovers. Following basic DNA origami design principles [19] and combining caDNAno and CanDo, one can be relatively certain that a new DNA origami design will assemble as designed; however, the yield is to be experimentally determined. Other design tools and software analysis packages are outlined in Note 11. 6. Optimize the positioning of membrane anchors. For scaffolded DNA origami, hydrophobic moieties are often placed on a base plate adjacent to the stem of the pore [12, 13]. By extending single-stranded DNA overhangs from the DNA origami structure, it is possible to attach many hydrophobic tags with just one sequence (to reduce the cost of the DNA synthesis). These overhangs are typically 15–20 bases long to ensure stable binding at room temperature. They can be placed in a standard [12] or a zipper-like arrangement [13]. For scaffold-free designs, the hydrophobic moieties are often placed on the transmembrane section of the pore itself [5, 6, 11, 13, 15]. To avoid twist and strain, it may be useful to choose positions where the helices point outward [9]. For both, scaffolded and scaffoldfree designs, it is crucial that the hydrophobic tags cannot all be inserted without insertion of the DNA nanopore—otherwise


Kerstin Go¨pfrich et al.

the DNA nanopores will lie flat on the membrane without additional energy gain for flipping into the ion conducting orientation. Careful geometrical considerations are required to design a functional pore. Here, it is important to consider the linker length and the length of the hydrophobic tag itself. Porphyrin, for instance, is more compact than cholesterol, making it better suited for very narrow architectures [15]. Note that the positioning of the anchors will also affect diffusion and membrane attachment [20]. To avoid aggregation of the structures, it may be helpful to shield the hydrophobicity of the tags. This can be realized with a dynamic hinge, hiding the hydrophobic tags prior to membrane insertion [21]. Alternatively, the extension of single-stranded overhangs which can wrap around the hydrophobic tags has been shown to prevent aggregation effectively [9]. 7. Design the DNA sequences. Custom DNA sequences can be purchased commercially. When placing the order, in addition to the DNA sequence, it is necessary to provide the type of purification. Standard desalting is sufficient for unmodified strands, however, HPLC purification is recommended for cholesterol- or fluorophore-tagged DNA sequences. Designing the correct DNA sequence is crucial. For scaffolded DNA origami nanopores, the DNA sequences of the staples are predetermined by the scaffold sequence. However, custom sequences can be assigned to single-stranded DNA overhangs for the attachment of hydrophobic groups. Designing the correct DNA sequence is crucial for both scaffolded DNA and scaffold-free DNA nanopores. NUPACK’s thermodynamic analysis can help select sequences without stable secondary structures and unwanted complementarities with other strands (see Note 12). A favorable set of DNA sequences will reduce the assembly times and produce a higher yield of the target structure. 8. Optional: Implement stimuli-response and/or selectivity. DNA nanopores provide the exciting possibility to implement stimuli-response and/or selectivity. The ideal pore for nanopore sensing should exhibit a stable low-noise conductance state. Yet many other applications, including controlled drug release systems, may require gated pores which are responsive to external stimuli and/or selective for specific analytes. Due to the negative charge of the DNA backbone, DNA nanopores are, to some degree, cation selective. They transport proportionally more cations compared to anions, and charge-based exclusion of larger analytes is possible [15, 22]. G-quadruplexes or other ion-sensitive DNA motifs could be attached to the mouth of the pore in order to engineer more specific selectivity. In terms of stimuli-response, note that voltage gating has been observed for all DNA nanopores. A high transmembrane

DNA Nanopore Design


voltage (typically above 50 mV) favors a lower conductance state [5, 23]. Ligand gating can be implemented by incorporating specific binding sites at the pore entry, for instance binding sites for complementary DNA [6]. In future designs, versatile stimuli-response could be achieved by exploiting light-sensitive modifications, pH-responsive DNA motifs, aptamers or other sequence motifs and functional groups (see Note 13). 9. DNA nanopore assembly. Following the design and the synthesis of the required DNA sequences, assembly of the DNA nanopores is carried out according to protocols used for assembly of standard DNA nanostructures [19]. Sequences carrying hydrophobic tags are often added in excess—either after assembly for DNA origami pores or in the folding mix of DNA tilebased architectures. 3.5 Guidelines for Lipid Membrane Reconstitution of DNA Nanopores

1. Choose lipid composition. The insertion efficiency does not depend on the design of the DNA nanopore alone. The lipid membrane itself has a strong influence on the insertion characteristics. Hydrophobic tags show lipid membrane preferences. For example, cholesterol and tocopherol insert almost exclusively into liquid disordered domains composed of unsaturated lipids like DOPC at room temperature. Palmitate, on the other hand, has been shown to insert preferentially into the liquid ordered phase containing saturated lipids [24]. Charged lipid membranes may lead to nonspecific orientation absorption of the DNA nanopores. At the same time, it is important to select phospholipids which form stable bilayers with minimal leakage (like DPhPC, DOPC, POPC, EggPC) to avoid measurement artifacts related to ion conduction through transient lipid pores (see Note 14). 2. Choose experimental system to test for DNA nanopore insertion. The functionality of DNA nanopores has been tested by electrical ionic conductance measurements [11, 13] as well as optical observation of transmembrane transport [7]. However, only electrical measurements can give insights into the ionic conductance on the single-channel level. For these measurements, a voltage is applied across a lipid membrane while recording the ionic current. DNA nanopore insertion increases the permeability of the membrane and hence increases the ionic current in a stepwise manner. Different setups have been developed for single-channel ionic current measurements of protein pores [25]. For DNA nanopore insertion, high membrane fluidity and high membrane curvature are beneficial. Solventcontaining membranes have been shown to be suitable for DNA nanopore insertion [25]. Alternatively, the DNA nanopores can be added to GUVs which are then used for patch


Kerstin Go¨pfrich et al.

clamping. Instead of a stepwise conductance increase, the overall conductance of the lipid bilayer patch is measured. Patches can readily be formed and broken, giving a first indication for the insertion rate in a high throughput manner [26]. Generally, it is advisable to use two different setups to confirm the singlechannel conductance of a DNA nanopore, since different properties such as the lateral membrane tension can influence their conductance behavior [5, 22].


Notes 1. DNA nanopores typically require divalent ions (10–20 mM Mg2+) for long-term stability and membrane attachment. Buffers often contain EDTA, a chelator for divalent ions. Without EDTA, the concentration of divalent ions can be reduced. Alternatively, increased concentrations of monovalent ions can be used [27]. For some applications, UV crosslinking may be the method of choice to prevent degradation [28]. 2. If there is a low yield of GUVs, try the following: (a) Prepare fresh lipid stocks, sucrose solution, use new ITO slides and then make a fresh batch of GUVs. (b) Remove GUVs immediately after the electroformation is finished, or else they will fuse with the glass slides. (c) Try different lipid spreading techniques and spread smaller/larger volumes. Temperature and humidity can affect the yield of GUVs, so alter the procedure to suit your lab environment. 3. This ensures that the osmotic pressures inside and outside the GUV are equilibrated and sufficient MgCl2 is present for attachment and for the structures to remain intact. 4. Not all DNA nanopores will insert in the desired orientation perpendicular to the lipid membrane. Generally, only a small fraction of the pores that are absorbed on the membrane will create a transmembrane passage for ions. 5. If no insertions can be observed, first ensure that the DNA nanopores are assembled correctly in the final measurement buffer (e.g., with a combination of AFM, TEM, gel electrophoresis, DLS). Then check if the pores are attached to the membrane and not aggregated (e.g., by functionalizing them with a fluorescent dye). Confirm the unilamellarity of the membrane with a standard widely used protein pore, e.g., α-hemolysin. Increase the DNA nanopore concentration and the observation time. Finally, reconsider your design, in particular, the number and positioning of the hydrophobic tags. Also consider using a different measurement setup.

DNA Nanopore Design


6. Apart from DNA, other types of natural and unnatural nucleic acids (e.g., RNA, PNA, XNA [29]) may be suitable and interesting for the construction of nanopores as described here. Both DNA-hybrids and nanopores made entirely of other forms of nucleic acids are conceivable. Alternatively, DNA nanotechnology can be used to scaffold the arrangement of peptides and proteins leading to protein-DNA hybrid nanopores [30]. 7. Addition of surfactants (e.g., Triton, OPOE) can increase the insertion efficiency of DNA pores, but it also alters the membrane properties and can lead to artifacts related to the formation of transient lipid pores. Therefore, appropriate control experiments are particularly crucial if surfactants are employed. 8. Note that aggregation of the DNA nanopores can occur if too many hydrophobic tags are used [9, 13] or if their positioning (see Subheading 3.4, step 6) or their sequence design (see Subheading 3.4, step 7) is not optimal. 9. caDNAno is an open-source DNA origami design software developed by Douglas et al., available at [31]. Multiple online tutorials offer guidance for inexperienced users, and template designs are available for download. caDNAno can be installed as a stand-alone program as well as a plug-in for Autodesk Maya providing an interface for threedimensional visualization. Within caDNAno, possible DNA geometries are currently limited to hexagonal and square lattices. 10. CanDo is a free online tool to predict the three-dimensional shape and flexibility of scaffolded and scaffold-free DNA nanostructures in solution using finite-element analysis [32]. It is available at It can thus help to make design choices before the more cost- and time-intensive DNA synthesis. The structural predictions are regularly improved and updated [33]. 11. Other computational tools, including SARSE [34], Tiamat [35], oxDNA [36], tacoxDNA [37], vHelix [38], DAEDALUS [39], and ENRG MD [40] MrDNA [41], are useful for design and structure prediction of DNA nanostructures. 12. NUPACK is a free online tool for the analysis and the design of secondary structures of one or more interacting DNA sequences available at [42]. It is especially useful to optimize the sequence design for small scaffold-free DNA nanostructures and sequences that carry hydrophobic groups. 13. DNA nanopores have also been inserted into solid state nanopores [43], where they can serve as adapters to tune the pore size or to implement molecular recognition [44, 45]. While the


Kerstin Go¨pfrich et al.

functionalization with hydrophobic tags is normally not necessary in this case, the protocol provided here can also give guidance on the design of DNA nanopores for solid state supports. 14. Note that DNA nanopores will influence the lipid bilayer properties. The absorption of highly charged polymers creates an asymmetry and influences the spontaneous membrane curvature [46]. Lipid tubulation of GUVs has been observed as a consequence of the attachment of micromolar concentrations of DNA nanopores [8].

Acknowledgements K.G. received funding from the European Union’s Horizon 2020 research and innovation program under the Marie SkłodowskaCurie grant agreement No. 792270. K.G. further acknowledges support from the Winton Programme for the Physics of Sustainability, Gates Cambridge and the Oppenheimer PhD studentship. A.O. was supported by funding from the Engineering and Physical Sciences Research Council (EPSRC) and through the Vice Chancellor’s Award from the Cambridge Trust. U.F.K. received funding from an ERC Consolidator Grant (Designerpores 647144) and Oxford Nanopore Technologies. References 1. Fernandez-Lopez S, Kim HS, Choi EC, Delgado M, Granja JR, Khasanov A, Kraehenbuehl K, Long G, Weinberger DA, Wilcoxen KM, Ghadiri MR (2001) Antibacterial agents based on the cyclic D,L-α-peptide architecture. Nature 412:452–456 2. Go¨pfrich K, Platzman I, Spatz JP (2018) Mastering complexity: towards bottom-up construction of multifunctional eukaryotic synthetic cells. Trends Biotechnol 36:938–951 3. Deamer D, Akeson M, Branton D (2016) Three decades of nanopore sequencing. Nat Biotechnol 34:518–524 4. Sakai N, Matile S (2013) Synthetic ion channels. Langmuir 29:9031–9040 5. Seifert A, Go¨pfrich K, Burns JR, Fertig N, Keyser UF, Howorka S (2015) Bilayer-spanning DNA nanopores with voltage-switching between open and closed state. ACS Nano 9:1117–1126 6. Burns JR, Seifert A, Fertig N, Howorka S (2016) A biomimetic DNA-based channel for the ligand-controlled transport of charged

molecular cargo across a biological membrane. Nat Nanotechnol 11:152–156 7. Krishnan S, Ziegler D, Arnaut V, Martin TG, Kapsner K, Henneberg K, Bausch AR, Dietz H, Simmel FC (2016) Molecular transport through large-diameter DNA nanopores. Nat Commun 7:12787 8. Go¨pfrich K (2017) Rational design of DNA-based lipid membrane pores. Dissertation, University of Cambridge 9. Ohmann A, Go¨pfrich K, Joshi H, Thompson RF, Sobota D, Ranson NA, Aksimentiev A, Keyser UF (2019) Controlling aggregation of cholesterol-modified DNA nanostructures. Nucleic Acids Res 47:11441–11451 10. Ohmann A (2020) A synthetic lipid scramblase built from DNA. Dissertation, University of Cambridge, uk/handle/1810/303262 11. Go¨pfrich K, Zettl T, Meijering AE, Herna´ndez-Ainsa S, Kocabey S, Liedl T, Keyser UF (2015) DNA-tile structures induce ionic currents through lipid membranes. Nano Lett 15:3134–3138

DNA Nanopore Design 12. Go¨pfrich K, Li CY, Ricci M, Bhamidimarri SP, Yoo J, Gyenes B, Ohmann A, Winterhalter M, Aksimentiev A, Keyser UF (2016) Large-conductance transmembrane porin made from DNA origami. ACS Nano 10:8207–8214 13. Langecker M, Arnaut V, Martin TG, List J, Renner S, Mayer M, Dietz H, Simmel FC (2012) Synthetic lipid membrane channels formed by designed DNA nanostructures. Science 338:932–936 14. Burns JR, Stulz E, Howorka S (2013) Selfassembled DNA nanopores that span lipid bilayers. Nano Lett 13:2351–2356 15. Go¨pfrich K, Li CY, Mames I, Bhamidimarri SP, Ricci M, Yoo J, Mames A, Ohmann A, Winterhalter M, Stulz E, Aksimentiev A, Keyser UF (2016) Ion channels made from a single membrane-spanning DNA duplex. Nano Lett 16:4665–4669 16. Burns JR, Go¨pfrich K, Wood JW, Thacker VV, Stulz E, Keyser UF, Howorka S (2013) Lipidbilayer-spanning DNA nanopores with a bifunctional porphyrin anchor. Angew Chem Int Ed 52:12069–12072 17. Ohmann A, Li CY, Maffeo C, Al Nahas K, Baumann KN, Go¨pfrich K, Yoo J, Keyser UF, Aksimentiev A (2018) A synthetic enzyme built from DNA flips 107 lipids per second in biological membranes. Nat Commun 9:2426 18. Wei B, Dai M, Yin P (2012) Complex shapes self-assembled from single-stranded DNA tiles. Nature 485:623–626 19. Wagenbauer KF, Engelhardt FAS, Stahl E, Hechtl VK, Sto¨mmer P, Seebacher F, Meregalli L, Ketterer P, Gerling T, Dietz H (2017) How we make DNA origami. Chembiochem 18:1873–1885 20. Khmelinskaia A, Mu¨cksch J, Petrov EP, Franquelim HG, Schwille P (2018) Control of membrane binding and diffusion of cholesteryl-modified DNA origami nanostructures by DNA spacers. Langmuir 34:14921–14931 21. List J, Weber M, Simmel FC (2014) Hydrophobic actuation of a DNA origami bilayer structure. Angew Chem Int Ed 53:4236–4239 22. Maingi V, Burns JR, Uusitalo JJ, Howorka S, Marrink SJ, Sansom MS (2017) Stability and dynamics of membrane-spanning DNA nanopores. Nat Commun 8:1–12 23. Maingi V, Lelimousi M, Howorka S, Sansom MSP (2015) Gating-like motions and wall porosity in a DNA nanopore scaffold revealed by molecular simulations. ACS Nano 9:11209–11217


24. Loew M, Springer R, Scolari S, Altenbrunn F, Seitz O, Liebscher J, Huster D, Herrmann A, Arbuzova A (2010) Lipid domain specific recruitment of lipophilic nucleic acids: a key for switchable functionalization of membranes. J Am Chem Soc 132:16066–16072 25. Gutsmann T, Heimburg T, Keyser U, Mahendran KR, Winterhalter M (2015) Protein reconstitution into freestanding planar lipid membranes for electrophysiological characterization. Nat Protoc 10:188–198 26. Go¨pfrich K, Kulkarni CV, Pambos OJ, Keyser UF (2013) Lipid nanobilayers to host biological nanopores for DNA translocations. Langmuir 29:355–364 27. Martin TG, Dietz H (2012) Magnesium-free self-assembly of multi-layer DNA objects. Nat Commun 3:1103 28. Gerling T, Kube M, Kick B, Dietz H (2018) Sequence-programmable covalent bonding of designed DNA assemblies. Sci Adv 4:eaau1157 29. Pinheiro VB, Holliger P (2014) Towards XNA nanotechnology: new materials from synthetic genetic polymers. Trends Biotechnol 32:321–328 30. Spruijt E, Tusk SE, Bayley H (2018) DNA scaffolds support stable and uniform peptide nanopores. Nat Nanotechnol 13:739–745 31. Douglas SM, Marblestone AH, Teerapittayanon S, Vazquez A, Church GM, Shih WM (2009) Rapid prototyping of 3D DNA-origami shapes with caDNAno. Nucleic Acids Res 37:5001–5006 32. Castro CE, Kilchherr F, Kim DN, Shiao EL, Wauer T, Wortmann P, Bathe M, Dietz H (2011) A primer to scaffolded DNA origami. Nat Methods 8:221–229 33. Pan K, Kim DN, Zhang F, Adendorff MR, Yan H, Bathe M (2014) Lattice-free prediction of three-dimensional structure of programmed DNA assemblies. Nat Commun 5:5578 34. Andersen ES, Dong M, Nielsen MM, Jahn K, Lind-Thomsen A, Mamdouh W, Gothelf KV, Besenbacher F, Kjems J (2008) DNA origami design of dolphin-shaped structures with flexible tails. ACS Nano 2:1213–1218 35. Williams S, Lund K, Lin C, Wonka P, Lindsay S, Yan H (2009) Tiamat: a threedimensional editing tool for complex DNA structures. International workshop on DNAbased computers. Springer, Berlin, Heidelberg, 2008 36. Sˇulc P, Romano F, Ouldridge TE, Rovigatti L, Doye JPK, Louis AA (2012) Sequencedependent thermodynamics of a coarsegrained DNA model. J Chem Phys 137:135101


Kerstin Go¨pfrich et al.

37. Suma A, Poppleton E, Matthies M, Sˇulc P, Romano F, Louis AA, Doye JPK, Micheletti C, Rovigatti L (2019) TacoxDNA: a user-friendly web server for simulations of complex DNA structures, from single strands to origami. J Comput Chem 40:2586–2595 38. Benson E, Mohammed A, Gardell J, Masich S, Czeizler E, Orponen P, Ho¨gberg B (2015) DNA rendering of polyhedral meshes at the nanoscale. Nature 523:441–444 39. Veneziano R, Ratanalert S, Zhang K, Zhang F, Yan H, Chiu W, Bathe M (2016) Designer nanoscale DNA assemblies programmed from the top down. Science 352:1534 40. Maffeo C, Aksimentiev A (2020) MrDNA: a multi-resolution model for predicting the structure and dynamics of DNA systems. Nucleic Acids Res 48(9):5135–5146 41. Maffeo C, Yoo J, Aksimentiev A (2016) De novo reconstruction of DNA origami structures through atomistic molecular dynamics simulation. Nucleic Acids Res 44:3013–3019

42. Zadeh JN, Steenberg CD, Bois JS, Wolfe BR, Pierce MB, Khan AR, Dirks RM and Pierce NA (2011) NUPACK: analysis and design of nucleic acid systems. J. Comput. Chem 32:170–173. 43. Bell NAW, Engst CR, Ablay M, Divitini G, Ducati C, Liedl T, Keyser UF (2012) DNA origami nanopores. Nano Lett 12:512–517 44. Herna´ndez-Ainsa S, Bell NAW, Thacker VV, Go¨pfrich K, Misiunas K, Fuentes-Perez ME, Moreno-Herrero F, Keyser UF (2013) DNA origami nanopores for controlling DNA translocation. ACS Nano 7:6024–6030 45. Wei R, Martin TG, Rant U, Dietz H (2012) DNA origami gatekeepers for solid-state nanopores. Angew Chem Int Ed 51:4864–4867 46. Nikolov V, Lipowsky R, Dimova R (2007) Behavior of giant vesicles with anchored DNA molecules. Biophys J 92:4356–4368

Part II Single Molecule Detection and Analysis of Protein Analytes

Chapter 5 Determining the Orientation of Porins in Planar Lipid Bilayers Sandra A. Ionescu, Sejeong Lee, and Hagan Bayley Abstract Single-channel planar lipid bilayer (PLB) recording of bacterial porins has revealed molecular details of transport across the outer membrane of Gram-negative bacteria, including antibiotic permeation and protein translocation. To explore directional transport processes across cellular membranes, the orientation of porins or other pore-forming proteins must be established in a lipid bilayer prior to experimentation. Here, we describe a direct method for determining the orientation of porins in a PLB—with a focus on E. coli OmpF—by using targeted covalent modification of cysteine mutants. Each of the two possible orientations can be correlated with the porin conductance asymmetry, such that thereafter an I–V curve taken at the start of an experiment will suffice to establish orientation. Key words Porin, Outer membrane protein, OmpF, Planar lipid bilayer, Orientation, Electrophysiology, Cysteine labeling, Thiol reagents, Membranes


Introduction Bacterial porins are the most abundant proteins in the outer membrane of Gram-negative bacteria where they mediate the passive transport of nutrients and toxins, such as antibiotics and bacteriocins (strain-specific antibacterial proteins) [1]. Detergentsolubilized porins can be reconstituted into artificial bilayers, where they retain properties consistent with their in vivo function. Single-channel planar lipid bilayer (PLB) recording has become a powerful tool for measuring the currents that flow through individual protein pores. The technique can detect interactions between pores and other molecules with sub-millisecond resolution [2], and has been used to investigate porin physiology, including sugar transport through maltoporin [3], phage binding to FhuA [4], and antibacterial peptide translocation through OmpF [5]. It has also been used to develop stochastic sensors [6] based on the α-hemolysin pore for the detection of kinases and sugars [7, 8]. Bacterial porins comprise an attractive class of transmembrane proteins

Monifa A. V. Fahie (ed.), Nanopore Technology: Methods and Protocols, Methods in Molecular Biology, vol. 2186,, © Springer Science+Business Media, LLC, part of Springer Nature 2021



Sandra A. Ionescu et al.

for the development of additional sensor elements. Compared to multi-subunit pore-forming proteins—such as α-hemolysin [9], which is made up of seven polypeptide chains—each porin barrel is composed of a single chain. Modifications of the pore at desired locations can therefore be easily achieved by mutagenesis because (1) assembly of heteromers is not required and (2) mutations can be readily introduced over several β-strands. Because porins comprise largely symmetrical β-barrel structures, it is feasible for a porin to insert into a PLB with either the extracellular or periplasmic loops first. For example, the most abundant porin outer membrane protein F (OmpF), which adopts only one orientation in vivo [10], has been shown to insert into a PLB in both orientations [11, 12]. This is known from the two types of I–V curve asymmetry, in which the porin conductance is higher at either positive or negative applied voltages. Asymmetric ion conduction has also been observed with other porins, including maltoporin [13] and OmpG [14]. In order to employ the highly informative PLB technique to investigate directional transport through porins or to develop new sensors, the orientation of the porin in question must first be established. Previously, the orientation of the bacterial porins OmpF [11], maltoporin [13], OmpG [14], and mitochondrial VDAC [15] were indirectly inferred from interactions with antibiotics, reducing reagents and sugars applied from one side or the other of a bilayer, and by conductance changes in response to applied potentials, respectively. These approaches are not generally applicable. In this chapter a general and direct method for establishing porin orientation is presented based on the authors’ work with the trimeric E. coli OmpF [12]. The method uses targeted covalent modification of peripheral cysteine residues—which are not present in wild-type porins—introduced on the periplasmic and extracellular loops. Once a single-cysteine mutant has inserted into a PLB, an I–V curve is measured to establish porin I–V asymmetry. A thioldirected PEG reagent (mPEG-maleimide or -orthopyridyl disulfide) is then introduced on each side of the bilayer in turn; the molecular weight of the PEG is chosen such that it cannot diffuse through the porin. When the reagent is introduced on the side of the bilayer where the cysteine residue is exposed, a reaction will occur generating a PEG adduct that causes a concomitant decrease in the ionic current through the pore [12]. The site of reaction (cis or trans) can then be correlated to the I–V asymmetry of the channel to establish the absolute orientation of the porin with respect to the I–V curve. After the connection between the porin I–V curve and orientation has been established, an I–V curve measured at the start of each subsequent experiment is enough to establish the porin’s orientation. Caveats are that the wild-type porin must have an asymmetric I–V curve and the chosen cysteine mutation(s) must not significantly change this asymmetry. This

Orientation of Porins in Planar Lipid Bilayers


targeted covalent modification approach might be further applied to determine the orientation of a wide range of membranes channels and pores, including toxins, MACPF proteins, and transporters.



2.1 Solutions and Reagents

Prepare all solutions using ultrapure deionized and double-distilled water. Store buffers and reagents at room temperature unless otherwise indicated. Use Hamilton syringes for the handling of organic solvents. 1. Electrophysiology recording buffer (used for OmpF): 100 mM KCl, 20 mM potassium phosphate buffer, pH 7.0. Weigh 7.5 g KCl (see Note 1) and transfer to a graduated cylinder. Add 24.6 mL of K2HPO4 (0.5 M) and 15.4 mL of KH2PO4 (0.5 M). Add water to 900 mL. If necessary, adjust solution to pH 7.0 (see Note 2). Make up to 1 L with water. Filter buffer through a 0.22 μm polyethersulfone (PES) filter before use. 2. TCEP: Prepare a 20 mM tris(2-carboxyethyl)phosphine (TCEP) solution by dissolving 5 mg in 1 mL water. Make fresh before use. 3. Hexadecane (1% v/v in pentane): Prepare a 1 mL stock of 10% anhydrous hexadecane in anhydrous pentane by dissolving 100 μL hexadecane in 900 μL pentane. Dilute the solution tenfold in pentane and aliquot into glass vials. Use a PTFElined cap to minimize evaporation and store at 20  C. 4. DPhPC lipid (2.5 mg/mL in pentane): For bilayer formation, 1,2-diphytanoyl-sn-glycerol-3-phosphocholine (DPhPC) is used. Dissolve 25 mg of powdered lipid in 1 mL anhydrous pentane. Dilute the solution tenfold in pentane and aliquot into glass vials. Use a PTFE-lined cap to minimize evaporation and store at 20  C. 5. mPEG-MAL-5K: Prepare a 5 mM solution of methoxy polyethylene glycol-maleimide-5K (mPEG-MAL-5K) by dissolving 0.125 g in 0.5 mL electrophysiology recording buffer. Make fresh before use. 6. mPEG-OPSS-5K: Prepare a 5 mM solution of methoxy polyethylene glycol-orthopyridyl disulfide-5K (mPEG-OPSS-5K) by dissolving 0.125 g in 0.5 mL electrophysiology recording buffer (see Note 3). Make fresh before use. 7. DTT: Prepare a 200 mM dithiothreitol (DTT) solution by dissolving 31 mg in 1 mL electrophysiology recording buffer. Make fresh before use.



Sandra A. Ionescu et al.


1. Wild-type (WT) porin: Transform the plasmid containing the porin gene of interest into E. coli. Express and purify proteins according to a protocol of choice. For OmpF, express in E. coli BZB1107 (ompf knockout strain) and purify as previously described [5]. Store the purified protein in buffer containing 1% (w/v) n-octyl-β-D-glucopyranoside (β-OG) at 80  C. 2. Single-cysteine porin mutants: Introduce single-cysteine mutations on extracellular or periplasmic loops at a position where the thiol group will be exposed for reaction with thiol-directed PEG reagents (see Note 4). For OmpF, mutations E29C, N246C (extracellular), and D221C (periplasmic) have been successfully labeled after PLB reconstitution [12]. The extracellular and periplasmic designation is based on the orientation found in vivo [10]. Express according to item 1, supplementing the storage buffer with TCEP or DTT (1 to 2 mM) to avoid thiol oxidation.

2.3 Electrophysiology Setup

1. Recording compartment: two Delrin (acetal resin) compartments (0.5 mL volume). 2. PTFE film (25 μm thick). 3. RTV silicone coating. 4. PVC-coated copper wire. 5. Faraday cage. 6. Ag/AgCl electrodes: Oxidize silver wire (1.0 mm thick) overnight in 5% sodium hypochlorite. Solder tip of silver electrodes to PVC-coated copper wires. 7. Agarose salt bridge: 1.5% low-gelling agarose in 3 M KCl. Weigh out 1.5 g low-gelling agarose and 22.4 g KCl and add water to 100 mL. Microwave solution until agarose is fully dissolved. Transfer 1 mL aliquots into 1.5 mL centrifuge tubes. One tube is enough to make two electrodes. 8. Recording equipment and software for analysis. The authors use an Axopatch 200B patch-clamp amplifier connected to a CV 203BU head stage. Filter data through a 2 kHz low-pass Bessel filter and digitize with a Digidata 1322A converter at a sampling frequency of 10 kHz. Perform data analysis with pClamp 10.3 software.


Methods All experiments can be carried out at room temperature.

3.1 Planar lipid bilayer recording setup

1. Create a ~50 μm aperture in PTFE film: Suspend the film between a 2–5 mm spark-gap and discharge a spark for approximately 5 s. The aperture can be visualized and the diameter determined with a light microscope.

Orientation of Porins in Planar Lipid Bilayers


Fig. 1 Electrophysiology setup and the orientations of OmpF in planar lipid bilayers. (a) Planar lipid bilayer (PLB) recording setup. The recording compartment (black) resides in a Faraday cage and has two compartments: cis at ground and trans to which a voltage is applied. A 3 M KCl agarose salt bridge (yellow tips) forms an electrical connection between the recording buffer and each Ag/AgCl electrode, which is soldered to an insulated copper wire (orange). The PLB (usually made from DPhPC) is formed over a ~50 μm-sized aperture in a PTFE film (white box) sandwiched between the cis and trans compartments. A porin of interest, e.g., OmpF, can insert into the bilayer from the cis compartment in two possible orientations. (b) OmpF I–V curves obtained in 0.1 M KCl, 20 mM phosphate buffer, pH 7.0. The two asymmetric curves represent the two orientations that OmpF can adopt in a PLB. Positive asymmetry denotes higher conductance at positive applied voltages (blue); negative asymmetry denotes higher conductance at negative applied voltages (orange)

2. Sandwich the PTFE film between two Delrin compartments with RTV silicone coating. Allow to set overnight. 3. Make agarose bridges for the Ag/AgCl electrodes: Melt one aliquot of agarose salt bridge solution. Fill a 100 or 200 μL pipette tip with molten agarose and insert an electrode containing a rubber stopper at the top until a seal is formed (Fig. 1a). Allow agarose to solidify. 4. Connect the electrodes to the recording apparatus. The compartment containing the grounded electrode is designated “cis” and the opposite compartment to which voltage is applied “trans” (Fig. 1a). 5. Form a bilayer according to the Montal-Mueller solvent-free method [16, 17]. In brief, paint both sides of the PTFE film with 1% (v/v) hexadecane using a pipette or capillary and allow to dry for 10–20 min. Add electrophysiology recording buffer and TCEP (200 μM final) to both compartments followed by 2–4 μL of lipid solution (2.5 mg/mL) on each side (see Note 5). Wait 5–10 min for the pentane to evaporate, then slowly lower and raise the buffer level in both of the compartments to create the bilayer.


Sandra A. Ionescu et al.

3.2 Porin Reconstitution and Establishing I–V Asymmetry

For our setup, cis is at ground and voltage is applied to the trans side. Protein is introduced into the cis compartment. 1. Reconstitute a single porin (WT or cysteine mutant) into the bilayer. For OmpF, successful insertion has been realized by the addition of 50–500 ng of protein in 1% (w/v) β-OG to 500 μL of electrophysiology recording buffer [6]. Facilitate protein insertion by stirring, pipette mixing, or applying voltages in the 150–250 mV range. 2. To prevent further porin insertions, replace 20% of the buffer in the protein-containing cis compartment with fresh buffer a minimum of three times. Mix the compartment solution with a pipette between each exchange. 3. Measure the channel I–V curve by recording the current readout at voltages ranging from 100 to +100 mV in 10 mV increments (see Note 6). Positive asymmetry refers to a channel that exhibits higher conductance at positive applied potentials, whereas negative asymmetry refers to a channel that exhibits higher conductance at negative applied potentials. The two asymmetries will be assigned to the two possible orientations of the porin within the bilayer (see Note 7) (Fig. 1b).

3.3 Cysteine Mutant Labeling with mPEG-MAL-5K

1. Insert a cysteine mutant porin into a PLB and establish the I–V asymmetry according to step 3 of Subheading 3.2. 2. Record the porin baseline conductance at the applied potential that will be used for the cysteine-targeted covalent modification. For OmpF, use 70 mV (see Note 8). 3. Add mPEG-MAL-5K (1 mM final) to the cis compartment (see Note 9). A successful reaction between the porin thiol and mPEG-MAL-5K should produce a stable stepwise drop in conductance (see Note 10). The number of steps corresponds to the stoichiometry of the pore. The drop(s) in conductance should not be reversible. For trimeric OmpF, it is typical to observe three steps ranging from 1 to 5 pA, depending on the cysteine mutant [12] (Fig. 2a, TOP). Adduct formation is typically seen within 5 min of reagent addition, but this may vary among porins and with the buffer pH. 4. Add mPEG-MAL-5K to the trans compartment according to step 3. If a stepwise reaction has already been observed upon reagent addition to the cis side, no further reaction should be seen upon reagent addition to the trans side (Fig. 2b). 5. If no reaction (current step) is observed on either side of the bilayer, then the cysteine thiol may be (a) buried in the bilayer or (b) oxidized or may have reacted with reagent impurities. Cysteine reactivity can be assessed by carrying out the targeted covalent modification in bulk solution and analyzing the

Orientation of Porins in Planar Lipid Bilayers


Fig. 2 Determining cysteine mutant orientation by targeted covalent modification. (a) OmpF cysteine mutant inserted in an orientation that exposes the thiol groups to the cis side of the bilayer and exhibits a positive (Pos) I–V curve asymmetry (left panel). Top: Upon addition of mPEG-MAL-5K to the cis compartment, a PEG adduct is formed through a thio-ether linkage at each cysteine thiol, causing stepwise drops in the current. In the case of trimeric OmpF, three steps are observed (associated with levels L0–L3). The addition of DTT does not alter the conductance, as the thio-ether bond cannot be broken. Bottom: Upon addition of mPEG-OPSS-5K to the cis compartment, thiol-disulfide exchange initiated by the porin cysteine thiols generates three PEG adducts (associated with levels L0–L3). The resultant disulfide linkages are cleaved upon cis-side addition of DTT, returning the conductance to the initial open pore level (L0). (b) OmpF extracellular loop cysteine mutant inserted in an orientation that exposes the thiol groups to the trans side of the bilayer and exhibits negative (Neg) asymmetry. Upon addition of a thiol-directed PEG reagent into the cis side of the bilayer, no adduct is formed (the labeling reaction does not occur) as the bulky PEG molecule cannot pass through the porin. Therefore, as expected, there is no change in the open pore current (L0)


Sandra A. Ionescu et al.

products by SDS-PAGE. If labeling is efficient in the bulk phase but is not observed in the PLB recording, then (a) may be the case: repeat steps 1–4 with a new cysteine mutant. If no or poor labeling is observed on SDS-PAGE, then (b) may be the case: the preparation of the cysteine mutant (exposure to oxidizing conditions) and reagent purity should be considered. 6. Note the connection between the I–V curve asymmetry (positive or negative, as defined in step 3 of Subheading 3.2) and the side of cysteine labeling (cis or trans). 7. Repeat steps 1–4 several times to ensure that the side of adduct formation (cis or trans) is dependent on the orientation of the channel, as determined from the I–V curve. If a cysteine mutant porin exhibiting positive asymmetry reacts with the reagent on the cis side of the bilayer, then a porin with the same cysteine mutation inserted in the opposite orientation (negative asymmetry) should react with the reagent on the trans side of the bilayer. 3.4 Cysteine Mutant Labeling with mPEG-OPSS-5K

1. Repeat steps 1–7 in Subheading 3.3 using mPEG-OPSS-5K (1 mM final) (Fig. 2a, BOTTOM). In this case, the PEG adduct is attached to the porin via a disulfide bond (see Note 11). The OPSS reaction is slower than the MAL reaction. With OmpF, the reaction of all three cysteines with the OPSS reagent is sometimes observed only after 30 min [12]. 2. If a stepwise reaction is observed (see Note 12), add DTT (20 mM final) to the same compartment to which the mPEG-OPSS-5K successfully reacted with the porin. The DTT will cleave the disulfide bond to release the PEG adduct, leading to a stepwise increase in conductance with the same amplitudes seen during adduct formation.

3.5 Determining Porin Orientation in a PLB from the I– V Curve

1. Based on the targeted covalent modification results from Subheadings 3.3 and 3.4, establish the orientation of each porin cysteine mutant relative to the I–V curve, e.g., positive asymmetry indicates insertion into the bilayer from the cis compartment with the periplasmic loops first, leaving the extracellular loops in cis and the periplasmic loops in trans. 2. Ensure that the results are consistent across multiple porin cysteine mutants. If a particular cysteine mutant exhibits positive asymmetry upon bilayer insertion and reacts with reagent added to the cis side, then that same mutant exhibiting negative asymmetry should react with reagent on the trans side. Likewise, if an extracellular cysteine mutant exhibits positive asymmetry upon bilayer insertion and reacts with reagent added to the cis side, then a periplasmic cysteine mutant that exhibits positive asymmetry should react with reagent added to the trans side only (Fig. 3).

Orientation of Porins in Planar Lipid Bilayers


Fig. 3 Orientation of OmpF in planar lipid bilayers summary. The connection between I–V curve asymmetry and OmpF cysteine mutant bilayer orientation for our setup, in which the cis compartment is at ground and voltage is applied to the trans compartment. For porins exhibiting positive I–V asymmetry (blue trimers), the cysteine thiols will be exposed on the cis side for extracellular cysteine mutants and on the trans side for periplasmic cysteine mutants. For porins exhibiting negative I–V asymmetry (orange trimers), the opposite will be true. Therefore, for WT OmpF, positive I–V asymmetry means the extracellular loops are exposed in the cis compartment, leaving the periplasmic loops in trans. The opposite orientation applies for WT OmpF exhibiting negative I–V asymmetry

3. The absolute orientation of the cysteine mutant porin(s) can now be determined from a single-channel I–V curve. 4. Identify a porin substrate that shows side-dependent (cis vs trans addition) interaction patterns. For OmpF, enrofloxacin can be used [11, 12]. 5. Determine if the side-dependent interaction pattern relative to the channel I–V asymmetry (positive or negative) is consistent across cysteine mutants and WT to confirm that the relative orientations are the same (see Note 13). 6. The absolute orientation of the WT porin can now be determined from a single-channel I–V curve.



Sandra A. Ionescu et al.

Notes 1. The authors recommend using ultrapure KCl (99.999% trace metal basis) to minimize metal impurities that may interact with protein pores and affect the current read-out. KCl was chosen because the mobility of K+ and Cl ions are similar. Salt concentrations higher than 100 mM can be used so long as the I–V asymmetry of the pore is maintained (see Subheading 3.2). 2. Other buffers with different pH values can also be used. A pH between 7.0 and 8.0 is recommended to favor thiol deprotonation to the reactive thiolate, while avoiding side reactions between primary amines and the maleimide reagent. 3. The authors noted that successful reaction between the mPEGOPSS reagent and the OmpF cysteine mutant was highly susceptible to reagent impurities. Ask the supplier for a purity assessment or repurify the reagent in-house, for example, by using HPLC, where possible. Impurities remaining after synthesis, such as dipyridyl disulfide, may react with the free porin cysteine(s) to form a disulfide adduct, thus precluding the formation of a subsequent PEG adduct via thiol-disulfide exchange. 4. Multiple periplasmic and extracellular single-cysteine mutants should be prepared in parallel for testing. Depending on the position and orientation of a thiol group, adduct formation may not be favorable (e.g., if the cysteine is buried in or located near the lipid bilayer) or may not cause a decrease in porin conductance (e.g., if the adduct is too distant from the ion conducting pathway). PyMOL was used to choose free and exposed residues for cysteine mutagenesis based on the OmpF crystal structure (PDB: 3POX). Avoiding charged residues is recommended, as the glutamate (E29C) and aspartate (D221C) mutants showed a bigger deviation from the WT I–V curve than the asparagine mutant (N246C). 5. Addition of TCEP to the recording compartment prevents cysteine oxidation during the recording and will react less readily than DTT with the maleimide group. However, high concentrations of TCEP (>5 mM) can destabilize the lipid bilayer. 6. An increase in current asymmetry was noted at higher voltages (up to 200 mV was tested) for OmpF. This may be a general trend for the porins and an I–V curve that includes higher voltages (within the constraints of bilayer stability) can be measured in cases of weak asymmetry. 7. The two types of I–V curve (positive and negative asymmetry) should be linked by a 180 rotation, as expected for proteins in

Orientation of Porins in Planar Lipid Bilayers


opposite orientations in a symmetric bilayer. The I–V curves of WT porin and the cysteine mutants should be compared to ensure that they are similar, i.e., the pore conductance and asymmetry should not have been significantly affected by the single mutation. This allows extrapolation of the orientation obtained with cysteine mutants to the WT porin. For other methods of confirming the connection between the orientation of cysteine mutants and WT porin, see Subheading 3.5. 8. Any voltage can be used for the PEG labeling experiments so long as the bilayer remains stable for the duration of the experiment (for DPhPC, we recommend 150 mV as the maximum voltage) and the change in conductance upon PEG adduct formation can be resolved from the noise in the current trace. The noise will depend on the diameter of the bilayer and the acquisition frequency. 9. Because of the susceptibility of the OPSS reaction to impurities (see Note 3), the authors recommend investigating porin orientation using the maleimide (MAL) reagent first. Orientation and cysteine specificity can then be confirmed using the OPSS reagent. 10. Test the mPEG-MAL-5K reaction with WT porin to ensure there are no off-target interactions between the PEG and the porin, which may obfuscate results. 11. The labeling reaction is carried out with the reversible mPEGOPSS-5K reagent to affirm the results obtained with mPEGMAL-5K and to prove cysteine specificity, as primary amines can also react with maleimides at high pH (>8). 12. The amplitude of the current step(s) observed with the OPSS reagent should be similar to the current step(s) observed with the MAL reagent if the same sized PEG is used, e.g., 5 kDa was used for OmpF [12]. 13. Enrofloxacin exhibits side-dependent interactions with OmpF in the presence of Mg2+. In the case of positive I–V curve asymmetry and under negative applied potential, enrofloxacin added from the cis side causes fast blocking events, whereas trans-side addition does not significantly alter OmpF baseline conductance [11, 12]. An OmpF exhibiting negative I–V curve asymmetry should show the opposite interaction pattern, with fast blocking events observed after enrofloxacin addition from the trans side.


Sandra A. Ionescu et al.

References 1. Page`s JM, James CE, Winterhalter M (2008) The porin and the permeating antibiotic: a selective diffusion barrier in Gram-negative bacteria. Nat Rev Microbiol 6:893–903 2. Qing Y, Pulcu GS, Bell NAW, Bayley H (2018) Bioorthogonal cycloadditions with sub-millisecond intermediates. Angew Chem Int Ed 57:1218–1221 3. Kullman L, Winterhalter M, Bezrukov SM (2002) Transport of maltodextrins through maltoporin: a single-channel study. Biophys J 82:803–812 4. Udho E, Jakes KS, Buchanan SK, James KJ, Jiang X, Klebba PE, Finkelstein A (2009) Reconstitution of bacterial outer membrane TonB-dependent transporters in planar lipid bilayer membranes. Proc Natl Acad Sci U S A 22:21990–21995 5. Housden NG, Hopper JTS, Lukoyanova N, Rodriguez-Larrea D, Wojdyla JA, Klein A, Kaminska R, Bayley H, Saibil HR, Robinson CV, Kleanthous C (2013) Intrinsically disordered protein threads through the bacterial outer-membrane porin OmpF. Science 340:1570–1574 6. Bayley H, Cremer PS (2001) Stochastic sensors inspired by biology. Nature 413:226–230 7. Harrington L, Cheley S, Alexander LT, Knapp S, Bayley H (2013) Stochastic detection of Pim protein kinases reveals electrostatically enhanced association of a peptide substrate. Proc Natl Acad Sci U S A 110:E4417–E4426 8. Ramsay WJ, Bayley H (2018) Single-molecule determination of the isomers of D-glucose and D-fructose that bind to boronic acids. Angew Chem Int Ed 57:2841–2845

9. Song L, Hobaugh MR, Shustak C, Cheley S, Bayley H, Gouaux JE (1996) Structure of staphylococcal alpha-hemolysin, a heptameric transmembrane pore. Science 274:1859–1866 10. Hoenger A, Page`s JM, Fourel D, Engel A (1993) The orientation of porin OmpF in the outer membrane of Escherichia coli. J Mol Biol 233:400–413 11. Brauser A, Schroeder I, Gutsmann T, Cosentino C, Moroni A, Hansen UP, Winterhalter M (2012) Modulation of enrofloxacin binding in OmpF by Mg2+ as revealed by the analysis of fast flickering single-porin current. J Gen Physiol 140:69–82 12. Ionescu SA, Lee S, Housden NG, Kaminska R, Kleanthous C, Bayley H (2017) Orientation of the OmpF porin in planar lipid bilayers. Chembiochem 18:554–562 13. Danelon C, Brando T, Winterhalter M (2003) Probing the orientation of reconstituted maltoporin channels at the single-protein level. J Biol Chem 278:35542–35551 14. Chen M, Li QH, Bayley H (2008) Orientation of the monomeric porin OmpG in planar lipid bilayers. Chembiochem 9:3029–3036 15. Marques EJ, Carneiro CM, Silva AS, Krasilnikov OV (2004) Does VDAC insert into membranes in random orientation? Biochim Biophys Acta 1661:68–77 16. Gutsmann T, Heimburg T, Keyser U, Mahendran KR, Winterhalter M (2015) Protein reconstitution into freestanding planar lipid membranes for electrophysiological characterization. Nat Protoc 10:188–198 17. Zakharian E (2013) Recording of ion channel activity in planar lipid bilayer experiments. Methods Mol Biol 998:109–118

Chapter 6 Revelation of Function and Inhibition of Wza Through Single-Channel Studies Lingbing Kong Abstract Antibacterial resistance (AR) is causing more and more bacterial infections that cannot be cured by using the antibacterial drugs that are currently available. It is predicted that 10 million people will die every year by 2050 from infections caused by antibacterial resistant strains, surpassing the predicted numbers of deaths caused by cancer. AR is therefore a global challenge and novel antibacterial strategies are in high demand. To this end, the work on exploring the pore properties of a bacterial sugar transporter, WzaK30, has led to the discovery of the first inhibitor against bacterial capsular polysaccharides export. Recently, single-molecule recapitulation of capsular polysaccharide (CPS) export and pore formation properties of Wza barrel peptides have also revealed the possibility of a next-generation of Wza strategies. These strategies are based upon the first examination and understanding of the pore properties of wild-type (WT) and mutant WzaK30 in single-molecule electrical channel recording. The initially reported experimental procedures have been further developed to enable efficient studies of other Wza homologs that are more common in bacterial pathogens causing significant bacterial infections. Therefore, this chapter presents the most recent protocols and logistics behind the research on Wza channel activity, antibacterials, and strategies. The disciplines covered here include computation, molecular biology, biochemistry, electrophysiology, microbiology, and biophysics. Key words Single-molecule, Antibacterial, Wza, Capsular polysaccharides, Inhibition


Introduction Bacteria have various defense mechanisms that enable them to resist environmental challenges, which include immune response agents inside animal hosts. Among the defensive machineries in Gramnegative bacteria, the outermost layer called the outer membrane is often a thick sugar-rich layer which forms a barrier to regulate the interaction of the organism with environmental stimuli such as antibodies, chemokines, cytokines, or antibacterials. Capsular polysaccharides (CPS) and exopolysaccharides (EPS) are the most common sugar moieties in the outer membrane of Gram-negative bacteria. In the biosynthesis of the outer membrane, CPS and EPS moieties must be exported from the cell [1]. One lipoprotein,

Monifa A. V. Fahie (ed.), Nanopore Technology: Methods and Protocols, Methods in Molecular Biology, vol. 2186,, © Springer Science+Business Media, LLC, part of Springer Nature 2021



Lingbing Kong

Wza, is the translocon essential for the CPS export in various bacteria [2]. Research has shown that blocking channels could be a strategy to combat pathogens. For example, the anthrax PA63 channel has been studied by single-channel recording and a library of blockers have been prepared and tested for this channel [3]. Some of these compounds, especially the positively charged beta-cyclodextrins, have shown to be effective against the anthraxcausing bacteria at concentrations in the low nanomolar level. In a similar fashion, specifically blocking the Wza channel and therefore the translocation of CPS would inhibit the formation of the outermost capsule protective layer of the bacteria. In this way, targeting Wza would serve as a novel antibacterial strategy to combat the rise of antibacterial resistance [4, 5]. The pore properties of various mutant WzaK30 pores as well as proteolytically cleaved WT pores by proteinase K in situ were examined previously [6]. The WzaK30 mutants were then used in a screen of blockers and identification of binding sites in the pore. In this chapter, we outline the techniques used to characterize Wza channel activity and blockage on a single-molecule level [6, 7].



2.1 Reagents and Kits for Plasmid Construction and Extraction

1. Computational AutoDock.





2. Competent cells for bacterial transformation with WzaK30 and other Wza homologs: XL10-Gold or DH5α E. coli from Agilent or NIPPON GENE CO., LTD. 3. Polymerase chain reaction (PCR) for mutated WzaK30 and other Wza constructs: Phusion High-Fidelity PCR Kit and TA Cloning™ Kit with pCR™2.1 Vector. 4. Purification of PCR products: agarose gel electrophoresis with Gel Extraction Kit. 5. Bacterial culture and plates: The LB-Agar plates and LB medium are placed in an Incubated/Refrigerated Stackable Shakers at typically 37  C. 6. Plasmid DNA extraction kits: Miniprep and/or Maxiprep kits.

2.2 Reagents for Protein Expression

1. IVTT system: E. coli T7-S30 Extract System for Circular DNA kit (Promega). 2. SDS-PAGE: Criterion™ XT Bis-Tris precast gels (4–12% gel). 3. 20 XT MOPS running buffer (Biorad). 4. Gel drier with vacuum (e.g., the Model 583 gel dryer from Bio Rad). 5. TE buffer: 10 mM Tris–HCl, pH 8, 1 mM EDTA.

Wza Single Channel Studies


6. 4 Laemmli Sample Buffer: 200 mM Tris–HCl, pH 6.8, 400 mM DTT, 8% w/v SDS, 0.4% bromophenol blue, 40% glycerol. 7. Radiolabel: L-(35S)-Methionine. 8. Amino acid mixtures (minus methionine; minus leucine). 9. Centrifuge (25,000  g). 10. Whatman filter paper and X-ray film. 11. Microcentrifuge microfiltration device (0.45 μm pore size). 12. NanoDrop or UV-Vis spectrophotometer. 2.3 Reagents for Wza Channel Studies

1. High salt buffer for initial channel studies: 2 M KCl, 5 mM HEPES, pH 7.5. 2. Low salt buffer for evaluation of blockers under physiological conditions: 300 mM KCl, 5 mM HEPES, pH 7.5. 3. Oil: Hexadecane (10% v/v in pentane), Lipid: DphPC (10 mg/mL in pentane). 4. A planar bilayer setup: Two chamber Delrin chip. 5. Silver/silver chloride electrodes. 6. Methanethiosulfonate (MTSES). 7. Wza blocker: (am8γCD).


8. Equipment: Axopatch 200B (Axon Instruments), Digidata 1500B converter (Axon Instruments). 2.4 Reagents for In Vivo Blocker Inhibition (Polysaccharide Analysis)

1. Pro-Q Emerald 300 Glycoprotein Stain Kit (Molecular Probes). 2. M9 minimal growth media (Sigma) for growth of E69 strain with or without blocker. 3. Small molecules to be tested for Wza blocking activity. 4. 10% SDS-PAGE gel. 5. Gel imager suitable for detecting fluorescence in 500 nm range.



3.1 Computational Analysis for Open-Form Wza Mutants and Blocker Design

An outline of the methods referenced here is shown in Scheme 1. The availability of a high-resolution full structure of the WT Wza protein is crucial for this development. However, the X-ray crystal structure of WzaK30 is incomplete. It is therefore necessary to create a homology model. This homology model is used to base the design of Wza mutants and the selection of blockers for screening.


Lingbing Kong

Scheme 1 Flowchart to discover initial drug candidates against membrane protein channels. This flowchart summarizes the efficient development of the strong inhibitor of the WzaK30 channels and cellular functions in live bacteria. It is presented in a way that is applicable not only to the Wza homologs but also other membrane proteins, especially multimeric membrane proteins forming stable pore structures in cells

Wza Single Channel Studies


1. Download the protein structure from the protein data bank (PDB): ID 2j58.pdb for WzaK30 from structure/2J58. 2. Generate the homology model for the desired Wza by using Modeller (see Note 1). (a) Download the latest version of Modeller from https:// (b) Follow the instructions at: release.html. 3. Perform local relaxation or energy-minimization of the homology model by using Modeller. However, a newer common practice is to perform molecular dynamics with the protein in explicit water (see Note 2). 4. Obtain channel or pore diameters by manual measurements in PyMOL or plotted with software like HOLE. These values, especially those at the constriction sites (the narrowest sites), help in the prediction of potential channel current levels, signal-to-noise ratios, and stability. 5. Design initial channel blockers based on Wza structure using AutoDock and a template molecule such as γ-cyclodextrin [6]. The constriction sites of a channel are more stable than many soluble drug target proteins whose binding sites could be simply at the interface of a homo-dimeric protein complex. Most of the pore sizes of a multimeric channel are relatively unchanged (see Note 3). The criteria to select initial blockers is based upon size, shape, and charges or polarities. (a) Size: The relatively rigid constriction site of the Wza protein is at the alpha-helix barrel. The constriction site of WT WzaK30, for example, is 17 A˚ at the barrel. Therefore, a potential strong blocker should be in the same or similar size (see Note 4). (b) Shape: The Wza channels are multimeric and symmetric protein complexes (eightfold symmetry). The potential blockers should therefore be ideally cyclic and eightfold symmetric (see Note 5). (c) Charges or polarities: The affinities of Wza channels to potential blockers follow the general binding energies of bimolecular interactions, i.e., electrostatic interactions > hydrogen bonds > hydrophobic interactions. In the case of WzaK30, the residues at the constriction site of the alpha-helix barrel are composed of amino acids E369 and Y373, whose side chains are negatively charged in the case of E369 or act as hydrogen donor or acceptor in the case of the sidechain oxygen on Y373 (see Note 6).


Lingbing Kong

3.2 Mutagenesis of Wza Channels Using Molecular Biology Techniques

1. Using the Wza homology model, identify residues in the Wza channel to mutate in order to increase the size of the channel constriction site. There are two constriction sites at WT WzaK30, one at the barrel site at Domain 4 constructed by the E369 or Y373 residues while the other at a loop site at Domain 1 constructed by the Y110 residues (Fig. 1) (see Notes 7 and 8). 2. Analyze the restriction map of the Wza containing plasmid especially at the sites of interest to make sure the cloning and restriction digestions are as predicted. 3. Perform polymerase chain reaction (PCR) to acquire the desired constructs. The vector used for cloning should ideally contain an Ampicillin resistance gene. The general procedure for PCR is a two-tube approach (see Note 9). (a) In the first PCR reaction, mix the Forward primer that contains a mutation with the Reverse primer that anneals to the Ampicillin resistance gene.

Fig. 1 Domains of WzaK30 proteins (PDB: 2j58; modified with Modeller9.14). The domain in red (Domain 4) is the transmembrane domain facing the extracellular environment in live bacteria. The domain in blue (Domain 1) is the periplasmic domain located at the periplasm between the outer membrane and inner membrane

Wza Single Channel Studies


(b) In the second PCR reaction, mix the Reverse primer that contains the mutation with the Forward primer that anneals to the same site in the Ampicillin resistance gene. 4. If desired PCR products obtained, mix the two reaction mixtures in a 1:1 mole ratio and then digest with DpnI for 1–3 h. Transform the PCR products using suitable competent cells and subsequently miniprep to acquire the desired plasmid construct. 3.3 Protein Expression and Purification

1. Perform IVTT reactions by using the E. coli T7-S30 Extract System for Circular DNA kit (Promega). Place the reagents on ice before use. 2. Adjust the concentration of circular DNA to 100 ng/μL. Mix together 800 ng DNA with 2.5 μL amino acid mixture minus Methionine, 2.5 μL amino acid mixture minus Leucine, 2 μL L(35S)-Methionine, 20 μL S30 premix and 15 μL T7 S30 extract (see Note 10). 3. Place the 50 μL reaction mixture at 37  C and incubate for 3 h. 4. Centrifuge the reaction mixture at 25,000  g for 20 min. Resuspend the pellet, which contains Wza monomer and octamers, in 4 Laemmli Sample Buffer (add 12 μL buffer per 50 μL reaction). 5. Perform SDS-PAGE at 35–50 V at 4  C in 1 MOPS running buffer, which requires about 10 h or overnight to complete (see Note 11). 6. Place the gel on Whatman filter paper and dry the gel for about 7 h at room temperature (see Note 12). 7. Identify and extract octameric Wza proteins. Develop an autoradiograph of the gel in a darkroom and use the autoradiograph to locate the Wza octamer on the gel. The Wza octamer runs larger than 100 kDa [6]. 8. Excise the gel and hydrate the dried gel piece attached to filter paper in 100 μL TE buffer. Remove the filter paper and in an Eppendorf tube crush the gel into very small pieces with a homogenization pestle. Incubate the sample overnight at room temperature (see Note 13). 9. Remove the gel pieces by centrifuging in a microcentrifuge microfiltration device at around 5000  g for 5 min. 10. Aliquot the filtrate and store the samples at 80  C. The ideal concentration of the Wza octamer is approximately 100–400 μg/mL measured by using a Nanodrop.

3.4 Single-Molecule Electrical Channel Recording Protocols

1. Set up the following parameters on the single-channel recording instrumentation (i.e., the Axopatch 200B patch-clamp amplifier, pClamp software)


Lingbing Kong

(a) Axopatch 200B patch-clamp amplifier: CONFIG: WHOLE CELL β ¼ 1. Output gain: 5. Lowpass Bessel filter: 1 kHz. Scaling factor (V/pA): 0.001. (b) pClamp parameters: Acquisition mode: Gap-free. Sampling rate: 5 kHz. 2. Preparation of the bilayer: (a) Coat the aperture with a thin layer of hexadecane by adding 2 μL of (1:10) hexadecane pentane solution. (b) Add 1 mL of high salt buffer or low salt buffer to either chamber of the planar bilayer setup. Add DTT (200 μM) to the buffers when a Wza mutant with a cysteine mutation is used (see Note 14). (c) Add two drops (~10 μL) of DPhPC to each chamber. Allow 5 min for the pentane to evaporate. Pipet in both chambers to generate a bilayer. 3. Single-channel recording of Wza channels: (a) Add Wza octamer stock (1 μL) to the cis chamber (the control or ground chamber) and mix. Apply a positive potential to assist insertion of a Wza channel into the bilayer. (b) Upon insertion of a single Wza channel, apply positive and negative electrical potentials (50 mV) from the trans chamber to obtain the electrical responses of the channel, the channel properties of the Wza channels. (c) Confirm the current characteristics of the K375C or K375C/Y110G mutants: Add 1.5 mM of methanethiosulfonate (MTSES) reagent to trans chamber when one or several Wza proteins insert into the bilayer. The MTSES reagent typically blocks the pore completely when reacted with the eight K375C residues of the Wza octamer. To reverse the MTSES reaction with the Wza cysteines and reopen the pores, mix in 10 mM DTT to trans chamber (see Note 15). (d) Test the blocker efficiency against various octameric Wza mutants, e.g., WT Wza, Y110G, K375C/Y110G. An ideal blocker causes the current of the channel to drop significantly and reversibly (see Notes 16 and 17). (e) Upon identification of a suitable blocker, test a series of concentrations of the blocker with the same single channel to obtain the kinetic details of the binding events.

Wza Single Channel Studies


(f) Perform “Single-channel search” with the single-channel traces of the Wza channel interacting with the blocker in pCLAMP 10.0 software. Extract the dwell time of the “off” events and plot with a suitable bin and fit with an exponential equation to acquire the τoff of each set of binding events. koff is then obtained by 1/τoff. Extract the dwell time of the “on” events in a similar fashion. Plot the reciprocal of the dwell time of the “on” events, i.e., 1/τon against different concentrations of the blocker. The slope of this plot is the kon. Calculate the Kd by dividing the koff by the kon. At least three repeats (nr) are required to acquire an error (E) so the dissociation constant is expressed as Kd (averaged)  E (n ¼ nr) (see Note 18). 3.5 Live Bacteria Inhibitory Assay Protocols

The experimental protocols are optimized for the E. coli K30 E69 strain (see Note 19). 1. Bacteria growth and polysaccharide extraction (a) Grow the target pathogenic bacteria in M9 minimal medium in appropriate volume at 37  C to reach an OD600 between 0.5 and 1.0 (see Note 20). (b) Dilute the culture to give an approximate OD600 ¼ 0.005. Transfer 495 μL of this bacterial culture to an Eppendorf tube. (c) Add a solution of the blocker (5 μL) in a series of final concentrations from 10 μM up to 1 mM. (d) Place the samples in an incubator at 37  C with shaking at 300 rpm. (e) After overnight growth, adjust the bacterial cultures to OD600 ¼ 0.5. (f) Transfer 500 μL of the culture from each sample to a new Eppendorf tube. Centrifuge at 5000  g for 20 min. (g) Resuspend the pellet in 500 μL PBS. Mix with 500 μL of PBS-equilibrated phenol. (h) Heat the suspension to 65  C for 15 min (see Note 21). Mix the samples every 5 min until a homogeneous solution is achieved. (i) Centrifuge the solution at 4  C for 15 min. Transfer the supernatant (upper layer) to new Eppendorf tubes. (j) Wash the supernatants with dichloromethane twice to remove any residual phenol (see Note 22). Store the supernatant at 80  C.


Lingbing Kong

2. SDS-PAGE analysis of CPS content: (a) Dilute 5 μL polysaccharide extract in 5 μL PBS and then add 4 Laemmli sample loading buffer (5 μL). Load the samples onto a 10% SDS-PAGE gel. (b) Perform SDS-PAGE at constant current 20 mA at room temperature for about 80 min. 3. Staining and visualization: (a) Rinse the gel in distilled water and then stain it with the pro-Q Emerald 300 glycoprotein stain kit following manufacturer’s instructions. For example, oxidize for 30 min, wash three times, stain for 60 min, wash three times, and then immediately image. (b) Image the fluorescence in the 500 nm range with any UV lamp with excitation around 300 nm. (c) Analyze the fluorescence intensities of the CPS bands by using a modern image processing software such as Photoshop. (d) Plot the fluorescence intensities against the concentrations of the blocker to obtain the half inhibitory concentration (IC50).


Notes 1. The amino acid sequence in the WT WzaK30 used for Modeller is based on the complete gene sequence of the protein in E. coli E69. 2. The flexibility of microstructures of a membrane protein inserted in a relevant lipid bilayer and in an explicit solvent guides the proper understanding of protein functions in its natural cellular environment and thus the design and screening for a potential blocker. 3. It is common to encounter a protein structure from a protein PDB file or a snapshot out of a molecular-dynamics simulation with diminished symmetry. It is a routine practice to use perfectly circular protein structures and potential blockers for possible binding evaluations, such as Docking studies. However, it is important to also evaluate how difficult or easy these conformations are achieved through conformational changes from those with minimal energies. 4. Protons need to be added back to the structures so that potential hydrogen bonds could be evaluated accordingly. 5. Dendrimers with plenty of surface primary amines, such as DAB-Am-16 and DAB-Am-32, do block irreversibly to the

Wza Single Channel Studies


Wza channels with unaltered barrel structures on an irregular basis. These molecules do not have much value for use to inhibit the Wza functions in live bacteria and are therefore not pursued. 6. The pH of the biological environment of Wza is essential for the determination of the protonation or the deprotonation state of the relevant amino acids and the potential blockers. For example, the E369 residues at WzaK30 should therefore adopt the deprotonated state (–COO) at a slightly basic solution or environment (pH 7.5, for example), while the primary amines at other residues and the C-6 position of blocker Am8γCD should all be protonated (–NH3+). 7. WT WzaK30 pores do not generate stable ionic currents which is needed for accurate analysis of a small molecule library screen [6]. It is necessary therefore to acquire open-form WzaK30 pores through mutagenesis. For general open-form Wza mutant channels, the guideline for mutagenesis is to acquire stable channels with minimum changes to the structure of the pore. This is because the aim of protein engineering is to find blockers that are more likely to work on the WT target in live cells. The mutants with minimum changes to the WT Wza protein will increase the chance of successful blocker screening. 8. The X-ray crystal structure of wild-type WzaK30 (WT-WzaK30) (PDB ID: 2j58) shows two sites relatively narrower than the rest, one being the loop region of Domain 4 at the periplasmic side while the other being around the middle of the alpha-helix barrel. Previous reports have shown that mutation of residue Y110 has little effect on Wza protein structure. This point mutation is selected at the constriction site at the D4 domain, which has flexible loops. The ring of Y110 residues in the WzaK30 channel maintains a diameter of 3.7 A˚. Using Modeller to determine the diameter of the Y110G mutant, previous work reveals that the Y110G mutation yields a new diameter of 10.5 A˚ [6]. Other mutants K375C and K375C/Y110G were also engineered. It is crucial to validate the properties of new Wza mutants. Therefore, in the case of WzaK30, the introduction of the K375C mutation at the entrance of the pore is to allow the identification of the current level, signal/noise ratio, and stability of the WzaK30 channels [6]. 9. When the two-tube PCR approach does not afford the desired constructs, an alternative design of PCR primers for a TA ligation followed by site-specific digestion and ligation can be carried out. Generally, the two-tube approach is quick and efficient for simple constructs. 10. Alternatively, nonradioactive L-Methionine can be used in the IVTT reaction. After running the gel, stain the gel using


Lingbing Kong

standard Coomassie methods. After the gel is stained, excise the desired band containing Wza proteins and extract the Wza proteins as outlined in Subheading 3.3, steps 8–10. This method avoids the use of radioactive materials and there is no need to establish relevant facilities and regulations. 11. Limiting the applied voltage in SDS-PAGE to 35–50 V increases the yield of functional octameric WzaK30. A lower voltage than 35 V tends to trigger errors and lead to failures in power supply at a later stage. Increasing the voltage may be required after several hours; otherwise, the power supply may stop because of too low current. 12. A simplified approach for small gel piece drying is to use vacuum Bu¨chner funnel connected to an oil pump. In either way, the gel or gel piece is placed on the filter paper, which is covered by a small layer of cling film and a silicone rubber sheet that covers the whole area of the filter paper or any area necessary to prevent the vacuum from leaking. 13. It is important to leave the crushed gel pieces in a buffer on bench for at least an hour to allow sufficient diffusion of protein into the buffer. 14. EDTA has no effect on the channel properties, so there is no need to include it in the buffers. 15. The reaction of the cysteine residues with MTSES to form disulfide bonds is usually very rapid, within a few minutes. The cleavage of the disulfide bonds by addition of DTT takes much longer. It could take over 10 min for all the disulfide bonds to be cleaved, as indicated by the current level returning to the value before the addition of MTSES. 16. Blockers differ in binding constants. Dendrimers that cause irregular and irreversible blockades are not considered as an ideal blocker. 17. A charged channel blocker satisfies the Woodhull’s model for voltage-dependent binding of a charged channel blocker. 18. The repeats refer to individual Wza pores, i.e., three repeats of the blocker concentration test must be performed by three independent pores. Therefore, all independent repeats of the Wza channel should be included equally in the initial evaluation of the channel properties. 19. On-gel carbohydrate staining offers an efficient way to quantify the capsular polysaccharides (CPS) extracted from live bacteria grown in the presence or absence of the Wza blockers. The pro-Q Emerald 300 Glycoprotein Stain Kit (Molecular Probes) is suitable staining for CPS moieties with at least two vicinal hydroxyls in each repeating unit. This is because the oxidant in the pro-Q Emerald 300 Glycoprotein Stain Kit (Molecular

Wza Single Channel Studies


Probes) cleaves the –C(OH)–C(OH)– bond to generate aldehyde groups that will react with the primary amine groups of the fluorescent reagent to generate fluorescent signals at the CPS band on a SDS-PAGE gel. The K30 CPS repeating unit, for example, has two pairs of these adjacent hydroxyl groups, both at the glucuronic acid, i.e., the C2 and C3 hydroxyls and the C3 and C4 hydroxyls. Other bacteria with different CPS characteristics should be re-evaluated to determine the best conditions for carbohydrate staining. 20. M9 minimal medium is much cleaner than LB medium. The inhibitory effects of the blockers obtained with M9 minimal medium match well with those in complement-mediated killing. The results gained in the LB medium are meaningful at the qualitative level; however, attempts to obtain quantitative data in LB medium encountered difficulties, probably due to the complex mixture of the yeast extract. LB medium is therefore not used for quantitative experiments to determine the inhibitory concentration of a blocker. 21. Use caution when handling phenol. Perform this step in a chemical hood, as phenol is an irritant. 22. The high-speed centrifuge tubes used here should be made of polypropylene plastic instead of polycarbonate. The latter tends to become dissolved in the dichloromethane and leads to loss of samples.

Acknowledgements Professor Kong is currently a tenure-track Assistant Professor and Principal Investigator at Kagawa University and the International Institute of Rare Sugar Research and Education (IIRSRE). He is grateful for the start-up funds supported by Kagawa University and IIRSRE, which enabled the purchase of a patch clamp system and essential accessories for the establishment of the single-molecule electrical channel recording studies in the lab. The JSPS grants (Startup grant 17H06912 and Basic Research (B) grant 18H02109) for Kong are also highly appreciated, which are supporting the establishment of a multidisciplinary lab for the studies of the function and inhibition of Wza homologs other than WzaK30 that are causing more bacterial infections. Professor Kong also is grateful to Professor Hagan Bayley at the University of Oxford for his generous gift of all the Wza plasmids as well as some basic accessories for the planar bilayer system developed during Professor Kong’s PhD and postdoctoral research.


Lingbing Kong

References 1. Whitfield C (2006) Biosynthesis and assembly of capsular polysaccharides in Escherichia coli. Annu Rev Biochem 75:39–68 2. Dong C, Beis K, Nesper J, BrunkanLamontagne AL, Clarke BR, Whitfield C, Naismith JH (2006) Wza the translocon for E. coli capsular polysaccharides defines a new class of membrane protein. Nature 444:226–229 3. Nestorovich EM, Bezrukov SM (2012) Obstructing toxin pathways by targeted pore blockage. Chem Rev 112:6388–6430 4. O’Neill J (2016) Tackling drug-resistant infections globally: final report and recommendations. Review on Antimicrobial Resistance. Government of the United Kingdom

5. Kong L, Vijayakrishnan B, Kowarik M, Park J, Zakharova AN, Neiwert L, Faridmoayer A, Davis BG (2016) An antibacterial vaccination strategy based on a glycoconjugate containing the core lipopolysacchride tetrasaccharide Hep2Kdo2. Nat Chem 8:242–249 6. Kong L, Harrington L, Li Q, Cheley S, Davis BG, Bayley H (2013) Single-molecule interrogation of a bacterial sugar transporter allows the discovery of an extracellular inhibitor. Nat Chem 5:651–659 7. Kong L, Almond A, Bayley H, Davis BG (2016) Chemical polyglycosylation and nanoliter detection system enables single-molecule recapitulation of bacterial sugar export. Nat Chem 8:461–469

Chapter 7 Protein Analyte Sensing with an Outer Membrane Protein G (OmpG) Nanopore Monifa A. V. Fahie, Bib Yang, Christina M. Chisholm, and Min Chen Abstract Nanopore sensing is a powerful lab-on-a-chip technique that allows for the analysis of biomarkers present in small sample sizes. In general, nanopore clogging and low detection accuracy arise when the sample becomes more and more complex such as in blood or lysate. To address this, we developed an OmpG nanopore that distinguishes among not only different proteins in a mixture but also protein homologs. Here, we describe this OmpG-based nanopore system that specifically analyzes targets biomarkers in complex mixtures. Key words Outer membrane protein G, Sulfhydryl-maleimide chemistry, Protein-protein interactions, Nanopore gating, Extracellular loops, Loop dynamics, Biotin, Antibody, Streptavidin, Avidin, Electrophysiology


Introduction Nanopore sensing is an attractive analytical tool in the field of diagnostics. It has been used to identify small molecules [1, 2] and biomolecules [3–8]. Nanopores have been engineered from either pore-forming proteins, synthetic materials, or even 3D-DNA structures [9–23]. One of the most successful applications of nanopore sensing has been in nucleic acid sequencing [24]; however nanopore studies for protein sensing have gained considerable interest and development recently [24–29]. Proteins can be detected by (1) translocation or interaction within the walls or constriction site of the nanopore, (2) binding to external adaptors where a change in the adaptor is transduced to the nanopore, and (3) binding directly to the outer surface of the nanopore. For a translocating protein, the ionic blockade that is generated in the nanopore is sensitive to a myriad of features on the protein such as its folding state, monomeric/multimeric state, posttranslational modifications, or distribution of surface charge [8, 30– 33]. Thus, the same protein could generate different ionic

Monifa A. V. Fahie (ed.), Nanopore Technology: Methods and Protocols, Methods in Molecular Biology, vol. 2186,, © Springer Science+Business Media, LLC, part of Springer Nature 2021



Monifa A. V. Fahie et al.

blockades as it passes through a nanopore depending on its state. This can make detection specificity challenging. Specialized nanopores are those decorated with high affinity molecules that make them specific to only a few analytes. These types of nanopores have been mainly used to detect the native/ active state of protein analytes. Once the protein is bound to the high affinity molecule this can induce a change in the nanopore via an adaptor [5] or via direct binding [34–37]. The current method described in this chapter is an example of a specialized nanopore that discriminates among protein variants through direct interaction with analyte. Recently, we engineered a nanopore based on outer membrane protein G (OmpG) from Escherichia coli (E. coli). This nanopore has a unique feature where it generates a fluctuating current unlike other common nanopores. We exploited the dynamic nature of the seven flexible extracellular loops of OmpG to capture specific analytes (Fig. 1). Analytes are distinguished through changes in loop dynamics and protein-protein interaction with the OmpG loop surface [33, 36, 38, 39]. We observed that a single OmpG construct could distinguish among several proteins even structural homologs (Fig. 2). Because of the sensing power of OmpG, it is an exciting platform on which to build nanopore sensors for sensitive and specific analysis.



2.1 Cloning of Cysteine Mutant OmpG

1. pT7-OmpGwt plasmid. 2. Oligonucleotides for mutagenesis. 3. High fidelity polymerase. 4. Restriction enzymes such as DpnI. 5. Chemically competent DH5α cells. 6. Ampicillin (100 mg/mL).

2.2 Protein Expression of OmpG-Cysteine Mutant

1. Chemically competent BL21 DE3 cells.

2.3 Preparation of OmpG-Biotin or OmpG-Sulfonamide Nanopore

1. Storage buffer: 50 mM Tris–HCl, pH 8, 150 mM NaCl, 1 mM EDTA.

2. LB media (Miller version): Tryptone 10 g/L, yeast extract 5 g/ L, sodium chloride 10 g/L. 3. Isopropyl β-D-1-thiogalactopyranoside (IPTG).

2. Lysis buffer: 50 mM Tris–HCl, pH 8, 150 mM NaCl, 200 μg/ mL lysozyme, 1 mM EDTA, 3 mM TCEP. 3. Inclusion Body (IB) wash buffer: 50 mM Tris–HCl, pH 8, 1.5 M urea, 1 mM TCEP.

OmpG Nanopore Detection of Protein Analytes

Extracellular side Loop 4 Loop 3 T T Loop 2 N G E P Loop 1 G L G D Y G E A K K D Y A D S R V D Y G M Y G N D T A T E D D T H A Y R F V V W G G N N E I E Y M A G I N F H W D N R E E





Loop 5





Loop 6

D D Q I Loop 7 W E E D R D G W E H D N G D S





Outer membrane envelope




Periplasmic turns

extracellular loops




open C224

loop 6 loop 6 10pA

0 pA 1s

periplasmic turns

Fig. 1 Outer membrane protein G structure and characteristic gating pattern. (a) Flat representation of the OmpG protein amino acid sequence. The peptide sequence of the seven loops are highlighted in color and the residues used to mutate into cysteine are highlighted in white bold lettering. (b) Top and side view of the OmpG crystal structure. (c) Characteristic OmpG gating signal in the current recording

4. Binding buffer: 50 mM Tris–HCl, pH 8, 3 mM TCEP, 8 M urea. 5. Probe Sonication system. 6. 0.45 μm filters. 7. Anion exchange columns (Q sepharose). 8. Desalting columns (5 mL, GE). 9. Chromatography gravity/centrifuge columns (15 mL). 10. Falcon tubes (50 mL).


Monifa A. V. Fahie et al.







I (pA)


no binding

I (pA)

I (pA)

I (pA)


30 20 10 0 30 20 10 0 30 20 10 0

no binding


mAb binding

30 20 10 0





Fig. 2 Detection of different antibodies binding with OmpG-biotin construct. (a) Schematic of OmpG and antibody reversible interactions and corresponding current recording traces of no binding and binding signals. (b) Concentration-dependent binding of monoclonal antibody (mAb) with OmpG-biotin construct

11. Wash buffer: 50 mM Tris–HCl, pH 8, 3 mM TCEP, 8 M urea, 75 mM NaCl. 12. Elution buffer: 50 mM Tris–HCl, pH 8, 3 mM TCEP, 8 M urea, 200 mM NaCl. 13. High salt buffer: 50 mM Tris–HCl, pH 8, 3 mM TCEP, 8 M urea, 500 mM NaCl. 14. Labeling buffer: 50 mM HEPES, pH 7.0, 150 mM NaCl, 8 M urea. 15. Labeling reagents: Maleimide-PEG2-Biotin for preparing OmpG-biotin, Maleimide-PEG4-sulfonamide for preparing OmpG-sulfonamide. 16. Refolding buffer: 50 mM Tris–HCl, pH 9.0, 3.25% octyl glucoside (OG). 17. Trypsin enzyme. Trypsin activity buffer: 50 mM Tris–HCl, pH 8, 1 mM CaCl2. 18. Iodoacetamide. 19. Chymotrypsin enzyme. Chymotrypsin activity buffer: 100 mM Tris–HCl, pH 8, 10 mM CaCl2. 20. Bradford assay reagents. 21. SDS-PAGE materials and electrophoresis equipment. 2.4 Current Recording of OmpG-Biotin or OmpG-Sulfonamide Nanopore

1. Phospholipids: 10 mg/mL diphytanoylphosphatidylcholine (DPhPC) dissolved in pentane. 2. Oil: 10% v/v hexadecane dissolved in pentane. 3. Polytetrafluoroethylene film (25 μm thickness) for the aperture.

OmpG Nanopore Detection of Protein Analytes


4. Biotin-binding analytes—(a) streptavidin, (b) avidin, (c) mouse anti-biotin monoclonal antibody clone BTN.4, (d) mouse antibiotin monoclonal antibody clone SB58C. 5. Sulfonamide-binding analytes—(a) human carbonic anhydrase VII, (b) human carbonic anhydrase II. 6. Human blood (10%). 7. Equipment: two chamber chip, Ag/AgCl electrodes, Digidata 1320A/D board (Axon Instruments), Axopatch 200B integrating patch clamp amplifier (Molecular Devices). 8. Software: latest versions of pClamp and Clampfit.



3.1 Substitution of a Single Cysteine Mutation on OmpG Protein

1. Introduce single cysteine substitutions on either of OmpG’s seven loops by site-directed mutagenesis. The plasmid template is pT7-OmpG (see Note 1). The mutagenesis primers to mutate residues on OmpG loops are described in Table 1 (see Note 2). 2. Use a standard polymerase chain reaction protocol from a commercially available high-fidelity enzyme such as Phusion to generate the PCR product with the desired mutation (see Note 3).

Table 1 List of the oligonucleotides used to generate the OmpG mutants OmpG mutations

Mutagenic oligonucleotides

L1 (E24C)


L2 (S58C)


L3 (E101C)


L4 (T143C)


L5 (S182C)


L6 (D224C)


L7 (G264C)



Monifa A. V. Fahie et al.

3. Digest the template DNA in the PCR reaction mix with DpnI enzyme (see Note 4). 4. Transform the DpnI-digested PCR mix into competent E. coli DH5α cells (or an equivalent cell strain) onto LB-Agar plates with 100 μg/mL ampicillin. 5. Isolate single clones and identify the desired mutant construct with restriction enzyme digestion and DNA sequencing. 3.2 Expression and Purification of Denatured OmpG Cysteine Mutant from Inclusion Bodies 3.2.1 Expression of OmpG Proteins

1. Transform the validated pT7-OmpG cysteine mutant plasmid into BL21 DE3 E. coli competent cells and plate onto LB-Agar plates supplemented with 100 μg/mL ampicillin (LB-AgarAmp). Incubate LB-Agar-Amp plate(s) for 14–18 h at 37  C (see Note 5). 2. Isolate one or more colonies from the BL21 transformation and inoculate a starter culture of LB media supplemented with 100 μg/mL ampicillin. Allow the starter culture to grow 37  C with agitation at 200 rpm for 12–16 h. 3. Inoculate a larger volume (at least ten times volume of starter culture) of LB + Amp media with the starter culture. Allow cells to grow at 37  C, 200 rpm shaking until the OD600 reaches between 0.5 and 0.8 (see Note 6). 4. Add inducer isopropyl β-D-1-thiogalactopyranoside (IPTG) to culture at a final concentration of 0.5 mM to induce the protein expression and continue to shake culture at 37  C (see Note 7). 5. Harvest cells after 3–4 h of IPTG induction by centrifugation at 3000  g for 20–30 min. Remove LB supernatant and resuspend cell pellet in either storage buffer or lysis buffer (see Note 8). 6. If the cell pellet is resuspended in storage buffer, keep pellet in 20  C freezer for approximately 2 months or until ready to purify (Subheading 3.2.2, step 1). Once cell pellet is resuspended in lysis buffer, this lysate cannot be stored or frozen; the inclusion body must be prepared immediately (Subheading 3.2.2, step 2).

3.2.2 Isolation of OmpG Inclusion Body

1. Thaw OmpG cell suspension at room temperature if previously frozen in storage buffer (see Note 9). Add lysozyme to a final concentration of 200 μg/mL and reducing agent (DTT or TCEP) to a final concentration of 3 mM. Incubate the OmpG-containing lysate at room temperature for 20 min or 37  C for 10 min to allow the lysozyme to break down the cell wall. 2. Sonicate the cell mixture on ice to further break the bacterial membranes and shear the nucleic acid to reduce the lysate’s viscosity. Pulse 5 s on/5 s off for a total of 7 min (see Note 10).

OmpG Nanopore Detection of Protein Analytes


3. Centrifuge the lysate at 20,000  g for 20 min. Discard the supernatant. 4. Resuspend the pellet in inclusion body (IB) wash buffer. Incubate at room temperature for 10 min. 5. Centrifuge pellet again at 20,000  g for 15 min. Discard IB wash supernatant. The resulting inclusion body pellet can be stored at 20  C for up to 4 months. 3.2.3 Anion Exchange Purification of Denatured OmpG Cysteine Proteins

1. Solubilize the OmpG-containing inclusion body in binding buffer (see Note 11) for at least 30 min at room temperature or until pellet is ~90% solubilized (see Note 12). 2. Centrifuge the solubilized inclusion body at 20,000  g for 15 min. Discard the pellet and pass the supernatant through a 0.45 μm filter before purification. 3. Equilibrate anion exchange Q beads with at least 5 column volume of binding buffer. 4. Apply the filtered inclusion body solution onto the equilibrated Q beads to allow the OmpG to bind. Collect the flow-through. 5. Wash the beads with 5 column volume of binding buffer. Collect wash. 6. Wash the beads with 5 column volume of wash buffer. Collect second wash. 7. Elute protein from beads with 3 column volume of elution buffer into clean centrifuge tubes. 8. Wash beads with 5 column volume of high salt buffer to remove any bound proteins or nucleic acids. Collect third wash. 9. Analyze protein content for each wash or elution collection with SDS-PAGE. A 12% or 15% gel is suitable for OmpG analysis. 10. For short-term storage (48 h or less), place eluted protein in 4  C (see Note 13). 11. Clean the Q beads: Wash beads with 10 column volume of distilled water. Check the manufacturer’s instructions on appropriate cleaning solvents and protocols for the beads. Store beads in 20% ethanol at 4  C for future use.

3.3 Ligand Labeling and Refolding of OmpG Proteins 3.3.1 Conjugation of Maleimide Ligand to OmpG Cysteine Mutant

1. Determine the protein concentration of the eluted OmpG fraction(s) with a Bradford assay (see Note 14). 2. Use about 0.5–1.0 mg of protein for the labeling reaction. Buffer exchange the OmpG using size exclusion chromatography (SEC) column equilibrated with labeling buffer. Refer to the manufacturer’s recommendations for appropriate volume of protein to purify with the column. For example, if using a


Monifa A. V. Fahie et al.

5 mL HiTrap desalting column (GE), no more than 0.75 mL protein may be injected onto column in order to efficiently exchange the buffer (see Note 15). 3. To label the OmpG cysteine mutant with small ligands like maleimide-PEG2-biotin or maleimide-PEG4-sulfonamide, incubate the protein with excess ligand in a molar ratio of at least 1:5 (protein to ligand) at room temperature (~23  C) for 1–2 h with moderate shaking (~100 rpm) (see Note 16). 4. Add DTT to a final concentration of 10 mM to quench the excess maleimide ligand. 5. Buffer exchange in elution buffer without reducing agent by SEC to remove excess free ligand. 1. Dilute the labeled OmpG mixture with refolding buffer until the final concentration of urea is approximately 3 M (see Note 17).

3.3.2 Refolding of Labeled OmpG Protein

2. Incubate samples at 37  C for 48–72 h. 3. Test the refolding efficiency of the labeled OmpG proteins by heat-denatured gel shift assay. Take one replicate of the refolded OmpG protein (at least 2 μg) and prepare with SDS-sample buffer. Take a second replicate of the OmpG protein, mix with SDS-sample buffer, and boil for 10–15 min at 95  C. 4. Run both boiled and unboiled proteins on an SDS-PAGE gel and analyze the shift in apparent molecular weight of the boiled vs. unboiled sample (Fig. 3). 5. Store refolded OmpG protein in small aliquots (10–20 μL) at 80  C for up to 1 month (see Note 18). 1. After diluting denatured OmpG into refolding buffer, mix OmpG and streptavidin in a 1:1 molar ratio and incubate at 23  C for 2–5 min (see Note 19).

3.4 Testing Labeling Efficiency of Refolded or Denatured OmpG Proteins

2. Add SDS-sample buffer and run samples on a 12% or 15% SDS-PAGE gel (Fig. 4). As a negative control, run the labeled OmpG minus the streptavidin to compare.

3.4.1 Labeling Efficiency of Biotin-Labeled OmpG (Gel Shift Assay) Loop 1 kDa 34.6-



Loop 2 -


Loop 3 -


Loop 4 -

Loop 5 +



Loop 6

Loop 7





heat -denatured -refolded

Fig. 3 Refolding efficiency gel analysis of OmpG-ligand constructs. Constructs were denatured at 95  C for 10–15 min and compared with refolded construct. Proteins were loaded onto a 12% or 15% SDS-PAGE gel and protein mobility in gel was analyzed as an indicator for their folding status

OmpG Nanopore Detection of Protein Analytes Loop 1 -

Loop 2 Loop 3



kDa 55.642.734.6-




Loop 4

Loop 5





Loop 6 -



Loop 7 -


streptavidin -streptavidin-OmpG complex

-denatured OmpG

Fig. 4 Labeling efficiency of OmpG-biotin loop constructs. OmpG-biotin constructs were heat denatured in SDS-sample buffer at 95  C for 10–15 min. Thereafter, one sample was incubated with 1:1 mole ratio of streptavidin protein for 5 min at ambient temperature, while no streptavidin was added to the other sample. The constructs, streptavidin, were run on SDS-PAGE gels and gel mobility shifts in the presence of streptavidin were analyzed as a marker for labeling efficiency

3. Using an imaging software, such as Image J, perform a densitometric analysis on the OmpG only sample vs. the labeled OmpG + streptavidin mixture to determine the labeling efficiency estimate. 3.4.2 Proteolytic Digestion of Labeled OmpG-Sulfonamide for Mass Spectrometric Analysis

1. Digest the labeled denatured OmpG, i.e., OmpG in 8 M urea with trypsin or chymotrypsin based on the manufacturer’s protocol for the specific protease. Trypsin digestion is suitable for analysis of peptides from loops 2, 3, 4, and 5. To analyze loops 1, 6, and 7 digest the OmpG with chymotrypsin instead. 2. Dilute denatured OmpG in trypsin activity buffer so that the urea concentration is 1 M or lower. Reduce any existing disulfide bonds with DTT at a final concentration of 10 mM (see Note 20). Incubate at 23  C for 30 min, then add iodoacetamide to a final concentration of 50 mM. Incubate at 23  C in the dark for 1 h with gentle agitation. 3. Use a trypsin:OmpG mole ratio range of 1:20 to 1:75 and incubate digestion mixture at 23  C for 12–16 h. 4. Alternatively, for cleavage with chymotrypsin, add DTT to a final concentration of 5 mM to reduce any unlabeled OmpG dimers in ~8 M urea. Incubate for 5 min. 5. Add 15 mM iodoacetamide and incubate for 15 min at room temperature in the dark. Dilute the sample with chymotrypsin activity buffer to obtain