DNA Manipulation and Analysis 1071630032, 9781071630037

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Methods in Molecular Biology 2633

Garry Scarlett  Editor

DNA Manipulation and Analysis

METHODS

IN

MOLECULAR BIOLOGY

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

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

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-by step 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.

DNA Manipulation and Analysis Edited by

Garry Scarlett Biophysics Laboratories, School of Biological Sciences, University of Portsmouth, Portsmouth, UK

Editor Garry Scarlett Biophysics Laboratories School of Biological Sciences University of Portsmouth Portsmouth, UK

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-3003-7 ISBN 978-1-0716-3004-4 (eBook) https://doi.org/10.1007/978-1-0716-3004-4 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This 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 If there is such a thing as a single secret to the mystery of life, then that secret is probably DNA. Since 1953 that has been a very badly kept secret, detailed descriptions and images of its structure and function abound in everything from learned articles to pop culture. However, we now not only know the structure of DNA but can decipher its code and even modify that code. We have learned to read and write on a molecular and biological level, the technologies that allow us to read and write DNA form the focus of this book. This book is designed for scientists moving into recombinant DNA technologies, covering techniques that are important for conducting experiments in fields as wide ranging as developmental to structural biology. The chapters are laid out as readers of the long running Methods in Molecular Biology series have come to expect, with an accessible theory section followed by a detailed method and finally the ever-useful notes and troubleshooting pages. No preface on the art of gene engineering could go without mentioning the discovery of the structure of DNA and the subsequent cracking of the code by a series of elegant experiments in the 1950s and 1960s. However, the key developments that are centrally relevant to this book were the first recombinant DNA experiments of Berg and, shortly after, Boyer-Cohen in 1971 and 1972; indeed, some of the principles in those early procedures have recognisable descendants described in the following chapters. A landmark development in the field in the mid-1970s was the arrival of Sanger dideoxy sequencing, replacing difficult and dangerous chemical methods for determining the order of bases on a piece a DNA, with a much easier enzymatic approach. The following decade of the 1980s was dominated by two major breakthroughs, the polymerase chain reaction (PCR) and the less mentioned but equally important phosphoramidite chemistry approaches to making short single-stranded oligonucleotides. Without these developments, many of the techniques in this book would simply not be possible, they have enabled not only de novo generation of DNA fragments but also the rapid amplification, site selection and targeted mutation of sequences. Initially, much of the early recombinant DNA work undertaken was on sequences of DNA inserted into plasmids, replicating circular extra-chromosomal DNA found in bacteria that encode useful attributes for the cell. Indeed, plasmids remain the work horses of the DNA laboratory, and the first few chapters detail a variety of different methods of inserting DNA sequences into them, each with specific strengths and weaknesses. Following on from these, the next three chapters discuss how to manipulate and create DNA sequences, which in conjunction with the molecular cloning methods discussed in the earlier part of the book provide powerful options in the laboratory for the would-be molecular biologist. The last decade of the twentieth century and the dawn of the new millennium saw increasingly rapid strides in the complex field of genomic modification, not only bacterial genomes but also the genomes of eukaryotic cells, including those in multicellular organisms. Early attempts at eukaryotic genomic modification suffered from significant issues with targeting the correct location within the genome, a problem that became worse the bigger the genome to be manipulated. The first technology that really tackled this was based upon zinc finger nucleases; this was rapidly replaced by TALENs before the extremely powerful and easy to use CRISPR/Cas9 system rose to its current dominance. Chapters 9, 10, and 11 discuss transgenics in the model system Xenopus, with Chaps. 10 and 11 describing methods for CRISPR targeted gene knockouts and gene insertions, respectively. The book then has two

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examples of the growing field of ‘cell-free’ DNA work, dealing with the raising of aptameric sequences and high throughput array technologies. Chapters 14, 15 and 16 deal with the key underpinning technologies of all DNA work, making oligonucleotides and reading the sequence. Much of this is now available as outsourced services for starter laboratories, but in-house provision allows for flexibility and increased speed. The last chapter deals with ethical considerations. The growing importance of DNA and gene manipulation to wider society has led to increasing public and governmental scrutiny, not only of ethics but also the risks of recombinant DNA technologies. Many jurisdictions have now introduced guidelines and laws governing the making and containment of genetically modified organisms, this is in addition to legal implications of using animals in research. Recombinant technologies have developed remarkably since those first experiments in the early 1970s; the growth of manufactured kits and outsourced services have brought nucleic acid–based experiments into the range of many laboratories that previously would have struggled to set up the infrastructure. We hope this book helps provide the expertise for scientists embarking on their first forays into these types of projects. Finally, I would like to thank the many chapter authors to who have contributed and provided their time and expertise to help support the scientific community with the resources held within this book. Portsmouth, UK

Garry Scarlett

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

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1 Classical Recombinant DNA Cloning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ana Mikic´, Arqam Alomari, and Darren M. Gowers 2 A Sequence- and Ligation-Independent Cloning (SLIC) Procedure for the Insertion of Genes into a Plasmid Vector . . . . . . . . . . . . . . . . . . . . . . . . . . . . Robert A. Holland 3 Molecular Cloning Using In Vivo DNA Assembly . . . . . . . . . . . . . . . . . . . . . . . . . . Sandra Arroyo-Urea, Jake F. Watson, and Javier Garcı´a-Nafrı´a 4 Assembling Multiple Fragments: The Gibson Assembly. . . . . . . . . . . . . . . . . . . . . . Luisana Avilan 5 TA Cloning Approaches to Cloning DNA with Damaged Ends DNA . . . . . . . . . Charlotte Ayling 6 PCR-Based Assembly of Gene Sequences by Thermodynamically Balanced Inside-Out (TBIO) Gene Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Timothy J. Ragan and Helen A. Vincent 7 Random Mutagenesis by PCR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F. A. Myers 8 In Vitro Site Directed Mutagenesis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Michael J. McClellan 9 Xenopus Transgenesis Using the pGateway System . . . . . . . . . . . . . . . . . . . . . . . . . . Liliya Nazlamova 10 CRISPR/Cas9 Gene Disruption Studies in F0 Xenopus Tadpoles: Understanding Development and Disease in the Frog . . . . . . . . . . . . . . . . . . . . . . . Anita Abu-Daya and Annie Godwin 11 A CRISPR/Cas-Based Method for Precise DNA Integration in Xenopus laevis Oocytes Followed by Intracytoplasmic Sperm Injection (ICSI) Fertilization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sian Angela Martin 12 A Lambda-Exonuclease SELEX Method for Generating Aptamers to Bacterial Targets . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Robert Gowland and Darren M. Gowers 13 Generation of Functional-RNA Arrays by In Vitro Transcription and In Situ RNA Capture for the Detection of RNA-RNA Interactions . . . . . . . Helen A. Vincent, Charlotte A. Henderson, Daniela Lopes Cardoso, and Anastasia J. Callaghan

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Chemical Synthesis of Oligonucelotide Sequences: Phosphoramidite Chemistry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . John Brazier 15 Low Throughput Direct Cycle Sequencing of Polymerase Chain Reaction (PCR) Products . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . George D. Zouganelis and Nikolaos Tairis 16 Nanopore Sequencing for Mixed Samples. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Angela H. Beckett and Samuel C. Robson 17 Ethics, Legality, and Safety for Geneticists . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Simon E. Kolstoe Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors ANITA ABU-DAYA • European Xenopus Resource Centre, University of Portsmouth, Portsmouth, UK ARQAM ALOMARI • Department of Basic Sciences, College of Agriculture and Forestry, University of Mosul, Mosul, Iraq SANDRA ARROYO-UREA • Institute for Biocomputation and Physics of Complex Systems (BIFI) and Laboratorio de Microscopı´as Avanzadas (LMA), University of Zaragoza, Zaragoza, Spain LUISANA AVILAN • Centre for Enzyme Innovation, University of Portsmouth, Portsmouth, UK CHARLOTTE AYLING • Biophysics Laboratories, School of Biological Sciences, University of Portsmouth, Portsmouth, UK ANGELA H. BECKETT • Centre for Enzyme Innovation, University of Portsmouth, Portsmouth, UK JOHN BRAZIER • Reading School of Pharmacy, University of Reading, Reading, UK ANASTASIA J. CALLAGHAN • Biophysics Laboratories, School of Biological Sciences, University of Portsmouth, Portsmouth, UK JAVIER GARCI´A-NAFRI´A • Institute for Biocomputation and Physics of Complex Systems (BIFI) and Laboratorio de Microscopı´as Avanzadas (LMA), University of Zaragoza, Zaragoza, Spain ANNIE GODWIN • European Xenopus Resource Centre, University of Portsmouth, Portsmouth, UK DARREN M. GOWERS • Biophysics Laboratories, School of Biological Sciences, University of Portsmouth, Portsmouth, UK ROBERT GOWLAND • Department of Biochemistry, University of Cambridge, Cambridge, UK CHARLOTTE A. HENDERSON • Biophysics Laboratories, School of Biological Sciences, University of Portsmouth, Portsmouth, UK ROBERT A. HOLLAND • Syngenta Crop Protection Research, Berkshire, UK; Centre for Enzyme Innovation, University of Portsmouth, Portsmouth, UK SIMON E. KOLSTOE • Reader in Bioethics, University of Portsmouth, Portsmouth, UK DANIELA LOPES CARDOSO • Biophysics Laboratories, School of Biological Sciences, University of Portsmouth, Portsmouth, UK SIAN ANGELA MARTIN • European Xenopus Resource Centre (EXRC), University of Portsmouth, Portsmouth, UK MICHAEL J. MCCLELLAN • Ludwig Institute for Cancer Research Ltd, University of Oxford, Oxford, UK ANA MIKIC´ • Biophysics Laboratories, School of Biological Sciences, University of Portsmouth, Portsmouth, UK F. A. MYERS • Biophysics Laboratories, School of Biological Sciences, University of Portsmouth, Portsmouth, UK LILIYA NAZLAMOVA • Clinical and Experimental Sciences, South Academic Block, Southampton General Hospital, Southampton, UK TIMOTHY J. RAGAN • Leicester Institute of Structural and Chemical Biology, Department of Molecular and Cellular Biology, University of Leicester, Leicester, UK

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SAMUEL C. ROBSON • Centre for Enzyme Innovation, University of Portsmouth, Portsmouth, UK NIKOLAOS TAIRIS • Senior Forensic DNA Analyst & Technical Witness Expert (Greece), Augusta, GA, USA HELEN A. VINCENT • Biophysics Laboratories, School of Biological Sciences, University of Portsmouth, Portsmouth, UK JAKE F. WATSON • IST Austria, Klosterneuburg, Austria GEORGE D. ZOUGANELIS • Human SciencesResearch Centre, College of Science & Engineering, University of Derby, Derby, UK

Chapter 1 Classical Recombinant DNA Cloning Ana Mikic´, Arqam Alomari, and Darren M. Gowers Abstract Traditional molecular cloning involves a series of linked experimental steps performed with the overall goal of isolating (“cloning”) a specific DNA sequence—often a gene. The main purpose of cloning is to study either that DNA sequence or the RNA or protein product it encodes. Building on key enzymatic discoveries in the late 1960s, gene cloning was pioneered in the early 1970s. Since then, DNA cloning and manipulation have been used in every area of biological and biomedical research, from molecular genetics, structural biology, and developmental biology to neurobiology, ancient DNA studies, and immunology. It is a versatile technique that can be applied to a variety of starting DNA types and lengths, including cDNAs, genes, gene fragments, chromosomal regions, or shorter fragments such as PCR products and functional control regions such as enhancers or promoters. The starting DNA can originate from any cell, tissue, or organism. In this chapter we will cover traditional (“classic”) molecular cloning strategy. This comprises six linked stages in which (1) PCR is used to amplify a DNA region of interest that is then (2) digested with restriction enzymes, alongside a selected vector, to produce complementary ends crucial for the two molecules to be (3) ligated by an ATP-dependent DNA ligase, creating a recombinant DNA molecule. The recombinant DNA is then (4) introduced into competent bacterial cells by transformation and (5) grown on a selective agar media, followed by (6) colony-PCR for screening purposes. We provide a worked example to demonstrate the cloning of an average-size gene (in this case the 2 kb DNA ligase A gene) from E. coli into a common plasmid expression vector. We have included six color figures and two tables to depict the key stages of a classical molecular cloning protocol. If you are cloning a segment of DNA or a gene, remember that each DNA cloning experiment is unique in terms of sequence, length, and experimental purpose. However, the principles of traditional cloning covered in this chapter are the same for any DNA sequence; we have included a detailed notes section, so you should easily be able to transfer them to your own work. Some of the following chapters in this volume will cover other, more recently developed, cloning protocols. Key words Molecular cloning, Gene cloning, Restriction endonucleases, Plasmid, PCR, Transformation, Bacteria

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Introduction

1.1 Historical Perspective

Before the 1970s, the largest challenge to the progression of chromosomal and genetic research was lack of a reliable and reproducible method for isolating and analyzing DNA sequences of interest.

Garry Scarlett (ed.), DNA Manipulation and Analysis, Methods in Molecular Biology, vol. 2633, https://doi.org/10.1007/978-1-0716-3004-4_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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This changed in the early 1970s when three key discoveries were brought together to revolutionize genetics and initiate the field of modern molecular biology. The first discovery, in the late 1960s, occurred in the laboratory of Werner Arber and Stuart Linn, whose group characterized the first bacterial restriction endonucleases [1]. Restriction endonucleases are a class of bacterial DNA-binding enzymes that evolved as a defense against bacteriophage infection. They target and cleave specific recognition sequences within bacteriophage genomes, causing phage genome degradation. Although much attention was given to the characterization of these enzymes, termed “restriction factors,” their ability to cut specific DNA sequences was not fully exploited until the laboratory of Daniel Nathans first used them to map the Simian Virus 40 (SV40) genome [2]. Soon after, many more restriction enzymes were isolated and their recognition sequences determined, creating a way to selectively and specifically cut DNA molecules. Today, thousands of restriction endonucleases are known, with many hundreds available from commercial suppliers. The second key discovery that contributed to the development of molecular cloning was made earlier in the 1960s by several laboratories, including those of Martin Gellert, I. Robert Lehman, Charles Richardson, and Jean Weigle. This was the discovery of a class of enzymes able to perform the opposite function to restriction endonucleases, namely, to join DNA ends. These are called the DNA ligases [3, 4]. It is important to note that the formation of a phosphodiester bond between two juxtaposed DNA ends (specifically, between a 5′-phosphate and 3′-hydroxyl group) by DNA ligase is not a sequence-specific activity. Therefore, this discovery provided a means to re-join any two DNA fragments cut by restriction endonucleases. The first experiment to use restriction enzyme digestion and subsequent ligation was performed in 1972 by Paul Berg’s laboratory, successfully synthesizing the first recombinant DNA molecule [5]. Although this confirmed the concept that the DNA from any two species may be joined and a resulting recombinant DNA molecule can be created, the process was long, laborious, and had a low yield. However, this changed with the third discovery—of bacterial transformation by plasmids—that made it possible for the molecular cloning method to become a widely applicable technology. Bacterial transformation was in fact discovered by Griffith in the 1930s, in studies on lethal and non-lethal bacterial strains [6]. Transformation of bacterial cells enabled effective replication of the created recombinant DNA molecule. However, its application for the purpose of molecular cloning was not fully realized until 1972, when Stanley Cohen and Herbert Boyer treated bacterial cells with calcium chloride to induce the uptake of plasmids carrying antibiotic-resistance genes [7]. This was a ground-

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breaking study that set the stage for the first molecular cloning experiment to be performed in 1973 by Boyer, Cohen, and Chang [8]. Their method joined the individual digestion, ligation, and transformation steps to pioneer gene cloning and lay the foundation for the millions of molecular cloning experiments since. 1.2 Traditional Molecular Cloning Overview

Traditional molecular cloning consists of a set of linked experimental steps performed with the overall goal of isolating and replicating (“cloning”) a specific DNA sequence—often a gene—from a host organism, in order to study it further. The chosen gene or region of interest is combined with a plasmid to create a novel (not found in nature) recombinant DNA molecule. The traditional plasmid cloning protocol follows six steps, and these are clearly summarized in Fig. 1. Prior to the experiment, careful consideration needs to be made about the choice of plasmid vector, restriction enzyme(s), bacterial cells, recombinant selection, and screening methods. In the following short sections, we provide a little more background on each of these stages and highlight important information to consider.

1.3

A cloning vector is a small DNA molecule into which another DNA molecule can be inserted without disrupting the stability of the vector within a host organism. Cloning vectors can be of different sizes and from different hosts. The most commonly used vectors are bacterial plasmids, which are circular and typically several thousand base pairs in size (2–10 kbp). Alternatively, bacteriophages, cosmids, bacterial artificial chromosomes (BACs), yeast artificial chromosomes (YACs), or mammalian artificial chromosomes (MACs) have been developed (but are not covered further in this chapter). Regardless of the type or size of a vector, all standard cloning vectors must contain at least three structural features to allow for the recombinant DNA to be made and replicated. A vector must possess: (1) a multiple cloning site (MCS), also known as a polylinker; (2) an origin of replication (ori), and (3) a selectable marker gene. A multiple cloning site is a region on a plasmid containing a number of different restriction enzyme recognition sites, and into which the exogenous DNA is incorporated during the cloning experiment. An origin of replication allows a vector to replicate independently of a host cell (such as an E. coli cell) to obtain multiple copy numbers per cell cycle. A selectable marker gene provides host cells with a selectable phenotype that can be used to clearly distinguish between cells that contain the vector and the ones that do not. This is frequently an antibiotic resistance gene (such as for beta-lactamase), as only cells containing the vector will be able to grow on a media containing the corresponding antibiotic [9].

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Fig. 1 Recombinant DNA cloning. A pictorial overview of a typical molecular cloning experiment, in which a chosen DNA sequence is amplified by PCR and ligated into a plasmid vector; the recombinant vector is then transformed into bacterial cells. There are six distinct steps, and these are shown in Panels A and B. Panel A: Molecular biology steps. Step 1: PCR amplification of a gene or region of interest to give a DNA insert for cloning (light blue). Step 2: restriction enzyme digestion of both the DNA insert ends (red and purple) and the chosen plasmid (green), to give compatible, cohesive (“sticky”) ends. Step 3: ligation reaction between the cut DNA insert and cut plasmid; a new recombinant DNA molecule is born. Panel B: Microbiology steps. Step 4: Transformation of recombinant plasmid (blue/green) into competent bacterial cells (orange) and growth of transformed bacterial colonies. Step 5: Selection of successfully transformed cells via antibiotic resistance (positive selection). Step 6: Screening of selected cells by colony-PCR, followed by DNA sequencing of the plasmid and insert

Some vectors contain additional features that allow for specific downstream purposes. For example, expression vectors contain structural parts required for the transcription of a recombinant DNA sequence to obtain the protein it encodes. These features include an inducible promoter, a ribosomal binding site (RBS), a termination codon, the recognition sequence for polyadenylation tail addition, and often an N- or C- terminal tag for easier purification [10]. For a comparison between sub-cloning and expression vectors, see Fig. 2. As various vectors are nowadays available for a

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Fig. 2 Types of plasmid vector: cloning vectors vs. expression vectors. A pictorial comparison between the essential structural components of a standard cloning plasmid vector (such as pUC19) and an expression plasmid vector (such as pET28b). Panel A: A standard cloning plasmid must contain an origin of replication (blue), a selectable marker (ampicillin resistance in this example, purple), and a multiple cloning site, MCS (red). The MCS contains a single recognition sequence for at least a dozen different restriction enzymes (to aid flexibility when cloning) and is often found within the LacZ operon (green). Panel B: An expression plasmid must contain all the structural parts of a standard cloning plasmid vector (origin of replication, a selectable marker, and multiple cloning site) as well as additional sequences necessary for protein expression. These are: a RNA polymerase promoter (yellow), ribosome binding site (dark pink), poly-A signal (orange), and a terminator (brown) sequence. It may also contain N- or C- terminal tags, such as histidine- or glutathione-S-transferase-tag (black lines) to aid recombinant protein purification

wide range of experimental purposes, careful consideration is required to select an optimal vector for your research. For traditional cloning purposes, the most commonly used vectors are plasmids, more specifically pUC19 and pBR322, while for gene expression purposes (e.g., to overexpress the protein product of a cloned gene), pBlueScript or pET vectors are a frequent choice. 1.4 Restriction Endonucleases

Deciding on a restriction endonuclease for your cloning experiment goes hand-in-hand with the selection of a cloning vector. This is because your experiment will not be successful unless the recognition sequence for the chosen restriction endonuclease is present within its multiple cloning sites. There are many different

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restriction endonucleases available that can broadly be divided into two distinct groups depending on the ends they produce on a cut DNA molecule—blunt or cohesive. Blunt ends are made by the action of restriction endonucleases that cleave the DNA sequence at the same nucleotide position on both strands and do not produce free overhangs. Conversely, restriction endonucleases that cleave the double-stranded DNA at different nucleotide positions generate unpaired bases at the either 3′ or 5′ ends, which are called cohesive, or “sticky,” ends. Cohesive ends of the complementary sequences can associate by hydrogen bonding, and this facilitates the incorporation of the DNA insert into a vector by increasing the stability of the recombinant DNA molecule prior to the ligation [11]. Moreover, cohesive ends ensure that the target DNA is inserted into a vector in a desired orientation (directional cloning), which is especially important for protein expression purposes. Directional cloning can also be achieved by choosing two different restriction endonucleases, one to digest each end of the target DNA [12]. The gene cloning example we show in this method uses directional cloning, utilizing BamHI and NdeI enzymes. 1.5 Recombinant DNA Production

After selecting the DNA sequence, the restriction endonuclease(s), and the cloning vector to be used in your experiment, the next step of the molecular cloning method is to use the Polymerase Chain Reaction (PCR) to amplify the gene of interest and combine it with the plasmid, thus making a recombinant molecule (Fig. 1a). This involves designing a PCR primer pair that specifically flanks the region of DNA you wish to amplify. For good specificity and stringency, the forward and reverse primers are typically ~20 bases or more in length. Rarely will you find a restriction endonuclease recognition site perfectly adjacent to the DNA sequence you wish to clone. Therefore, each primer should also contain an extra (non-binding) section with the desired restriction enzyme recognition sequence, as illustrated in Fig. 1a, Step 1. As an example, the forward and reverse primers used for cloning the E. coli Ligase A gene are shown in Fig. 3. Following PCR amplification of the target DNA, the product (often referred to as the “insert”) is purified and ready for cloning. The next step is to enzymatically digest both the DNA insert and the vector you wish to insert it into, using the restriction endonuclease enzymes you chose earlier (Fig. 1a, Step 2). This step creates the complementary free ends on the two DNA molecules, which are then combined in the final step, ligation. In this step, a recombinant DNA molecule is created by the action of DNA ligase (Fig. 1a, Step 3). The most frequent choice of a DNA ligase for this purpose is the T4 DNA ligase, which uses ATP as a cofactor and is capable of ligating both sticky and blunt ends. Although ligation protocol does not vary considerably between cloning experiments, the possible resulting molecules do. A ligation reaction can produce three different output molecules, as shown in

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Fig. 3 Example of PCR primer pair for amplifying the E. coli Ligase A (LigA) gene. The target LigA gene is depicted as a heteroduplex with the top strand in black and bottom strand in grey. The regions of the forward primer (left) and reverse primer (right) that are complementary to the LigA gene are shown in blue. Hydrogen bonding between gene and primer during PCR amplification is indicated by blue dots between the strands. The recognition sequences for restriction endonucleases (NdeI, CATATG and BamHI, GGATCC) are shown in red. Black bases on the primers’ 5′-ends are random-sequence bases added to ensure restriction enzyme binding and cutting

Fig. 4: DNA insert can be joined with the cloning vector resulting in a recombinant DNA molecule, a vector can self-ligate without the inclusion of the target DNA sequence, or two target DNA sequences can be joined in tandem [13]. The undesired vector self-ligation and tandem insert DNA forms will be more likely to form if blunt-ended restriction endonucleases are used. However, the type of restriction endonuclease used is not the only contributing parameter. The output of the ligation reaction will also depend on other properties of the DNA molecules utilized, such as length/ size of the insert or vector, length of the cohesive ends (1, 2, 3, or 4 nucleotide overhangs), and the concentration ratio of the DNA insert to the vector in the reaction. The latter is an important parameter, and often ligation reactions require oversaturation with the DNA insert compared to the vector to ensure incorporation.

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Fig. 4 Potential outcomes of a ligation reaction in molecular cloning. During any ligation reaction, the DNA insert and cloning vector are first digested with the relevant restriction enzymes (top image). DNA strand joining by a DNA ligase can then produce different outcomes: (1) the two ends of the linearized plasmid re-join since the ends are close together in 3D space and more likely to meet each other and be resealed (undesired outcome); (2) an insert and linearized vector meet and are joined successfully (desired outcome), or (3) depending on the length of the inserts, free ends of the DNA insert join up to produce tandem (or more) repeats (undesired outcome). The trick to cloning is to try a few different ratios of insert: linearized vector to favor outcome (2)

1.6

Transformation

The final steps of a typical cloning protocol aim to multiply the synthesized recombinant DNA molecule in host cells (bacteria) and to exploit various methods for the selection of the obtained colonies. To multiply the recombinant DNA molecule, it must first be introduced into the selected host cells in the fourth step of the standard cloning protocol, transformation (Fig. 1b, Step 4). Similar to deciding on the appropriate restriction endonucleases and a compatible vector, the choice of host cells in which you aim to multiply your recombinant DNA molecules is also crucial for the success of the experiment. This decision should be made early on when designing the experiment and selecting the cloning vector, as not all vectors can be taken up by all host cells. In our case, using a plasmid vector, the most common choice is to transform it into a bacterial cell. The available bacterial cells are most often strains of Escherichia coli or Lactococcus lactis. These are frequently engineered for protein expression, but some strains are

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Fig. 5 Flowchart for selection of competent cells. This decision tree may help you select the best bacterial strain for your cloning reaction. Work from left to right, and you can choose the bacterial strain best suited to your downstream application. The cells used in our example method for cloning and expressing the LigA gene (BL21 (DE3)) are underlined

not, so pay attention when selecting those you wish to use in your experiment [14]. A useful flowchart to help you choose the right cells is provided in Fig. 5. Other than the genetic constitution of the bacterial cells used, the success of transformation will also depend on the transformation method performed. The two distinct methodological approaches are transformation by electroporation or chemical transformation [15]. Transformation by electroporation (also known as electropermeabilization) is achieved by applying a short, high-voltage pulse to bacterial cells. The electric shock makes the cell membrane more permeable and susceptible to the DNA uptake. This method is simple, fast, and gives high transformation efficiency for E. coli cells [16]. Alternatively, chemical transformation is achieved by heat-shocking bacterial cells at 42 °C to allow for the vector uptake. Prior to the heat shock, cells must be treated with high salt concentrations to increase the cell membrane permeability, and these are known as “competent” cells. Nowadays,

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competent bacteria can be bought pre-treated and ready for transformation. Once transformed, the host cells need to be grown on plates of nourishing media and incubated under optimal conditions before they can be examined for the presence of the recombinant DNA molecule. 1.7 Selection and Screening

Whether the host cells will take up the recombinant DNA molecule can never be guaranteed, therefore, numerous methods have been developed to assess the success of the steps carried out so far. However, even with these methods developed, testing each individual colony would be a laborious task if not for a selection step. Selection is the fifth step of the standard cloning protocol (Fig. 1b, Step 5), in which successfully transformed bacterial cells are selected for the phenotype they exhibit (positive selection) or lack (negative selection). The principle of positive selection is that cells are selected for the presence of a property encoded by the vector, as only cells containing the vector will be able to form colonies. The examples are auxotrophy and the aforementioned antibiotic selection test, where only bacterial cells that have obtained an antibiotic resistance gene carried by the vector will grow on a media containing the same antibiotic [17]. In contrast, the principle of negative selection is based on successfully transformed bacterial cells not exhibiting a property encoded by the vector and, therefore, being unable to grow and form colonies. The examples are SacB-counter selection or toxin-antitoxin system. By performing the selection step, the number of obtained colonies is limited only to the ones that have been successfully transformed and contain the vector. This enables you to focus on assessing the smaller number of colonies for the presence of the target recombinant DNA molecule [18]. The purpose of the sixth and last step of a standard cloning protocol is screening (Fig. 1b, Step 6). Often, selection and screening tests can be performed simultaneously, as in the case of the wellknown blue-white test exploiting the lactose (X-gal) metabolism pathway and β-galactosidase enzyme activity. This test offers a fast and simple way of assessing bacterial colonies for the presence of the target recombinant DNA molecule through observation of the plates with the naked eye. Alternatively, colony PCR is a rapid screening method in which primers used in the first step of the cloning experiment are repurposed for the detection of recombinant DNA molecules. However, these and many other methods (reporter gene or lethal gene assays) can be erroneous and give a false-positive result [19]. Therefore, the best practice is always to send samples of your grown colonies to be sequenced using Sanger dideoxy sequencing or other available sequencing methods. We provide an example of a standard cloning experiment in Fig. 6 for the cloning of the E. coli Ligase A and Ligase B genes.

Classical Cloning

A

M

1

2

11

B

3000 bp 2000 bp 1500 bp 1000 bp

500 bp

C

M

1

2

3

4

5

3000 bp 2000 bp 1500 bp 1000 bp 500 bp

D

Fig. 6 Example cloning results. The genes for DNA Ligase A and B were amplified from E. coli strain K-12, cloned into plasmid pET28b, transformed into BL21(DE3) cells, checked by colony PCR, and sequenced. Panel A: 1% (w/v) agarose gel with 1 kb marker (lane M) and samples of amplified Ligase A gene (lane 1, green arrow) and Ligase B gene (lane 2, blue arrow). Panel B: Agar plates showing transformants for Ligase A (left plate) and Ligase B (right plate). Panel C: 1.5% (w/v agarose gel with 1 kb marker (lane M) and samples of colony-PCR reactions for five separate colonies from the Ligase A plate in Panel B. A positive result is visible in lane 1. Panel D: Sanger dideoxy sequencing showing that the Ligase A gene is successfully cloned, without mutations; the green arrow indicates the methionine start codon for Ligase A 1.8 Current and Future Applications

Since its first application in 1973, molecular cloning has become an essential technique utilized in every area of biological studies including genetics, molecular cell biology, developmental biology, neurobiology, neuroscience, and immunology. Its importance is highlighted by the fact that nowadays, for a simple molecular cloning experiment, you may choose between 240 available protocols exploiting more than 3000 restriction endonucleases that recognize 230 different sequences, at least 1000 different cloning vectors, and hundreds of available host cells [20]. Moreover, new advances of this technology are continuously being developed, such as TA cloning, TOPO cloning, sequence and ligation independent

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cloning (SLIC), Gateway cloning, Gibson assembly, type IIS cloning such as Golden Gate and modular cloning (MoClo), and many others [21]. Some of these methods are covered in other chapters in this book.

2

Materials Ensure your laboratory is suitably registered and set up for molecular microbiology work. Wear a lab coat, gloves, and appropriate PPE at all times to avoid contamination and minimize the risk of nuclease degradation of samples. Use molecular biology-grade reagents and distilled, deionized, water for buffers.

2.1 Polymerase Chain Reaction (PCR) and Agarose Gel Electrophoresis

1. DNA containing the target sequence, region, or gene that you wish to clone. This can be in many different forms, including intact cells, purified genomic DNA, plasmid preparations, DNA libraries, mitochondrial preparations, chloroplast preparations, or previous PCR products (see Note 1). You will need a ~20 μL stock containing 10 ng of the DNA in nuclease-free water (0.5 ng/μL). 2. PCR primers that flank the DNA target sequence and that contain the recognition sequence(s) of the chosen restriction enzyme(s) (see Note 2). You will need a working stock of 10 μM in ~20 μL of nuclease-free water each. 3. PCR buffer (10× stock): 100 mM Tris–HCl, pH 8.6, 500 mM KCl, 1.5 mM MgCl2, 50% (v/v) glycerol, 0.8% (w/v) IGEPAL CA-630, 0.5% (w/v) Tween-20. Use 1× dilution. 4. Nucleotide mix: 10 mM dNTPs. Store at -20 °C. 5. Taq DNA Polymerase: 5 U/μL from a commercial supplier. Store at -20 °C. 6. Thin-wall 0.2 mL polypropylene PCR tubes. 7. PCR thermocycler set with the following program: For the selection of the annealing temperature (see Note 3). 8. Microcentrifuge tubes (1.5 mL). 9. DNA size ladder: a 500 bp—10 kb ladder provided with a 6× DNA Gel Loading Dye. Bought from a commercial supplier and stored at -20 °C. 10. A sterilized conical flask. 11. An agarose gel electrophoresis system. This includes a powerpack, gel cassette with removable end seals, combs, agarose powder (low endo-osmosis form), 10× TBE buffer stock, 10 mg/mL ethidium bromide (EtBr), and distilled water.

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13

12. TBE buffer (10× stock). For 1 L add: 108 g Tris base, 55 g boric acid, 900 mL distilled water, 40 mL 0.5 EDTA solution, pH 8.0. To obtain the 1× TBE running buffer, dilute the 10× TBE stock 10-fold in distilled water. 13. TBE running buffer (1×). For 1 L, add: 10 mL 10× TBE stock to 900 mL distilled water. Add 50 μL of 10 mg/mL ethidium bromide stock (0.5 μg/mL final concentration). 14. Small microwave oven. 15. An ultraviolet (UV) transilluminator capable of excitation at 302/312 nm (UV-B) or 365 nm (UV-A) for visualizing DNA-ethidium bromide complexes in agarose gels. It should be equipped with an LED camera for photo capture. Here we use a G:BOX gel doc. 16. A commercial spin-column PCR purification kit. Use per manufacturer’s instructions. 17. A small-volume UV spectrophotometer (e.g. a NanoDropTM or PicoDropTM device). 2.2 Restriction Enzyme Digestion

1. A cloning vector containing the same restriction enzyme recognition sites that were introduced at the ends of the DNA insert by the PCR primer pairs. Here we use the pET28b(+) 5368 bp expression plasmid vector with NdeI and BamHI recognition sites with a kanamycin resistance gene. For the guidance on vector selection (see Note 4). 2. NdeI restriction enzyme: 20 U/μL from a commercial supplier. Store at -20 °C (see Note 5). 3. BamHI restriction enzyme: 20 U/μL from a commercial supplier. Store at -20 °C (see Note 5). 4. Restriction enzyme buffer (10× stock), for example, rCutSmartTM buffer: 200 mM Tris-acetate, 100 mM magnesium acetate, 500 mM potassium acetate, 1 mg/mL recombinant bovine serum albumin (BSA), pH 7.9 at 25 °C. Store at -20 ° C (see Note 6). 5. A water bath set to 37 °C.

2.3

Ligation

1. T4 DNA Ligase enzyme: 5 U/μL from a commercial supplier. Store at -20 °C. 2. T4 DNA Ligase buffer (10× stock): 500 mM Tris–HCl, 100 mM MgCl2, 10 mM ATP, 100 mM DTT, pH 7.5 at 25 ° C. Obtained from a commercial supplier. Store at -20 °C. 3. Benchtop cooled-centrifuge capable of 13,000 rpm (15,100 × g) and chamber temperature of 0 °C.

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Transformation

Ensure your laboratory is prepared for the microbiology work. While handling bacterial cells, you should be working using aseptic techniques. 1. Competent bacterial cells. Selection depends on the downstream applications of the cloning experiment (for guidance see Note 7). Here we used BL21-DE3 competent cells from E. coli that can be used for protein expression. Cells are usually obtained from a commercial supplier; refer to the manufacturer’s instructions for storing and avoid freeze/thawing until being used. You may make your own competent cells, recipes are available online for this. 2. Ice and ice bucket. 3. A water bath set to 42 °C. 4. Freshly prepared SOC broth: 0.5% (w/v) yeast extract, 2% (w/v) tryptone, 10 mM NaCl, 2.5 mM KCl, 10 mM MgCl2, 10 mM MgSO4, 20 mM glucose, pH 7.0 (see Note 8). To prepare 1 L of SOC media combine 20 g tryptone, 5 g yeast extract, and 0.5 g NaCl. Stir the ingredients in 950 mL of distilled water until they dissolve. Add 10 mL of 250 mM KCl and adjust the pH to 7.0 with NaOH. Adjust the volume to 1 L with distilled water. Transfer 250 mL to four clean Pyrex bottles. Sterilize each by autoclaving at 121 °C (and 15 psi) for 20 min on liquid cycle with the cap loosely attached by autoclave indicator tape. Before using the media add 5 mL of sterile 2 M MgCl2 solution and 20 mL of sterile 1 M glucose solution, gently mix.

2.5

Selection

1. Freshly prepared and pre-warmed (37 °C) SOC media selection plates (see Note 9). To prepare 1 L of SOC media combine 20 g tryptone, 5 g yeast extract, and 0.5 g NaCl. Stir the ingredients in 950 mL of distilled water until they dissolve. Add 10 mL of 250 mM KCl and adjust the pH to 7.0 with NaOH. Adjust the volume to 1 L with distilled water. Add 1.5 g of bacto agar. Transfer 250 mL to four clean Pyrex bottles. Sterilize each by autoclaving at 121 °C (and 15 psi) for 20 min on liquid cycle with the cap loosely attached by autoclave indicator tape. Once cooled to ~60 °C, add 5 mL of sterile 2 M MgCl2 solution, 20 mL of sterile 1 M glucose solution, and 50 mg/mL kanamycin (see Note 10), gently mix. Pour approximately 15 mL into each labeled petri dish (final concentration of 50 μg/mL kanamycin). Label, seal, and store inverted at 4 °C until needed. 2. A water bath set to 37 °C. 3. Sterile 50-mL polypropylene tubes for liquid media growth. 4. Sterile, disposable plastic spreader. 5. Microbiology incubator-shaker set to 37 °C.

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2.6 Screening by PCR

15

1. Sterile 50-mL polypropylene tubes for liquid media growth. 2. The same number of sterile microcentrifuge tubes. 3. SOC liquid broth. 4. Nuclease-free water: high-quality molecular biology grade; stored at 4 °C. 5. Benchtop cooled-centrifuge capable of 13,000 rpm (15,100 × g) and chamber temperature of 0 °C. 6. Taq PCR master-mix (1×): 10 mM Tris-HCl, pH 8.6, 50 mM KCl, 1.5 mM MgCl2, 0.2 mM dNTPs (50 μM each), 5% (v/v) glycerol, 0.08% (w/v) IGEPAL CA-630, 0.05% (w/v) Tween20, 2 μM forward primer, 2 μM reverse primer, 25 Units/mL Taq DNA Polymerase. Prepare for the final reaction volume of 25 μL. For multiple PCR reactions (see Note 11). Keep on ice. 7. A laboratory vortex mixer. 8. PCR thermocycler set with the same program as used for PCR amplification of the DNA insert. Increase the number of cycles to 40. 9. Agarose gel electrophoresis Subheading 2.1.

equipment;

same

as

in

10. A commercial spin-column plasmid purification kit. Use per manufacturer’s instructions.

3

Methods Order and prepare all the materials and reagents in advance. Conduct all steps at room temperature unless otherwise specified. Wear lab coat, gloves, and appropriate PPE at all times to avoid contamination and minimize the risk of nuclease degradation of samples.

3.1 Polymerase Chain Reaction (PCR) and Agarose Gel Electrophoresis

1. Prepare a single 25 μL PCR reaction by adding the following components in a 0.2 mL PCR tube: 1 μL of 0.5 ng/μL DNA insert, 1 μL of 10 μM forward primer, 1 μL of 10 μM reverse primer, 2.5 μL of 10× PCR buffer, 1 μL of 10 mM dNTPs, 17.5 μL nuclease-free water. Mix gently by pipetting. 2. To the prepared PCR mix in Step 1 above, add 1 μL of the Taq DNA Polymerase. 3. Close the lid of the PCR tube and place it in the thermocycler. Run the 3-step standard PCR cycling under the previously set program outlined in Table 1. 4. Prepare 100 mL of 1% agarose gel. In a conical flask, add 1 g of powder agarose to 10 mL of 10× TBE and bring up to 100 mL with distilled water (see Note 12). Bring the solution to the boil by heating it up in the microwave for 2–3 min, checking every

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Table 1 PCR cycle conditions PCR step

Temperature (°C)

Time (s)

Denaturation

95

Melting Annealing Elongation

95 59 72

Final elongation

72

300

Hold

4

1

60

9 = ;

30 cycles

30 30 30

30 s. Take care to wear a padded glove—molten agarose can give nasty burns. Let the solution cool down to ~50 °C on the benchtop, then add 5 μL of 10 mg/mL ethidium bromide and mix by swirling. 5. Slowly pour the gel mix into a gel casting cassette and add comb(s). Leave the gel to set for ~30 min. 6. Once the gel has set, remove the end seals of the cassette and place it into the gel tank so that the comb is at the negative (black) electrode. Add 1× TBE running buffer (with 0.5 μg/ mL EtBr) to the tank until the buffer just covers the gel (see Note 13), then remove the comb(s) and clean the wells. 7. Take two microcentrifuge tubes and label them “S” for sample and “L” for the DNA size ladder. 8. In each of the two separate 0.5 mL microcentrifuge tube, add 1 μL of the 6× DNA gel loading dye and 8 μL of nuclease-free water. 9. Following the PCR, transfer 1 μL from the PCR tube into the “S” microcentrifuge tube—this is your amplified DNA insert. Mix gently by pipetting, take care not to introduce bubbles. 10. Prepare the DNA ladder by adding 1 μL of the DNA ladder to the “L” microcentrifuge tube. Mix gently by pipetting, take care not to introduce bubbles. 11. Load all of the 10 μL prepared DNA ladder and 10 μL of the PCR sample into separate wells of the gel. 12. Run the gel at 130 V for 30 min. 13. Visualize the gel using an ultraviolet (UV) transilluminator. Examine the gel to determine if the PCR amplification of your DNA insert has worked and is the correct size. Save an electronic copy of the gel image.

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17

14. Use the commercial microspin column purification kit to purify the dsDNA from the PCR samples for which a positive result was observed. Refer to manufacturer’s instructions. 15. Use a small-volume UV spectrophotometer to obtain the concentration of the dsDNA in the purified PCR sample by pipetting 2 μl of the sample onto the bottom optical surface and taking the reading. 3.2 Restriction Enzyme Digestion

1. Take a clean microcentrifuge tube and label it “Vector” or “V.” To the tube add 1 μg of the plasmid vector, 1 μL of the NdeI enzyme, 1 μL of the BamHI enzyme, 3 μL of the rCutSmartTM buffer, and bring up to 30 μL with nuclease-free water. 2. Take a clean microcentrifuge tube and label it “DNA” or “D.” To the tube, add 1 μg of the purified DNA sample from the PCR reaction, 1 μL of the NdeI enzyme, 1 μL of the BamHI enzyme, 3 μL of the rCutSmartTM buffer, and bring up to 30 μL with nuclease-free water. 3. Incubate both tubes in a water bath at 37 °C for 1 h.

3.3

Ligation

1. Take a clean microcentrifuge tube and label it “Ligation” or “L.” To the tube add 100 ng of the restriction enzyme treated vector from the “V” tube, 112 ng of the insert DNA from the “D” tube, 1 μL of T4 DNA Ligase, 1 μL of the 10× T4 DNA Ligase buffer, and bring up to 10 μL with nuclease-free water (see Notes 14–16). 2. Mix the “L” tube reaction gently, and ensure it is all at the bottom of the tube by pulsing it in a centrifuge at 11,000× g for 7 s. 3. Incubate the “L” tube at room temperature for 3 h or at 4 °C overnight (see Note 17).

3.4

Transformation

1. Thaw the competent cells on ice for 10–30 min. Thaw as many single-use competent cell tubes as transformation reactions you aim to conduct. Label them “T1,” “T2,” etc. 2. Transform the cells by adding 5 μL of the “L” reaction mix to each of the competent cell mixtures thawed (“T1,” “T2,” etc.). Mix by gently flicking the tube 4–5 times. 3. Incubate the tubes labeled “T” on ice for 30 min (see Note 18). 4. Heat shock the cells by placing the “T” tubes in water bath at exactly 42 °C for exactly 45 s (see Note 19). Do not mix or shake. 5. Immediately place the “T” tubes back on ice for another 2 min.

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Selection

1. To each “T” tube add 250 μL of the pre-warmed SOC liquid media and cap the tube. 2. Incubate the “T” tubes in water-bath at 37 °C for 1 h. Shake vigorously (225 rpm) or rotate (see Note 20). 3. Mix the cells by flicking and inverting the “T” tubes. In the clean microcentrifuge tubes, perform several 10-fold serial dilutions of the “T” tube contents in the SOC liquid media. We recommend three 10-fold dilutions. 4. Using a disposable plastic spreader, spread 50–100 μL of each dilution onto a separate SOC selection plate, labeled with the same number as the transformation reaction tube, by using a spread plate technique. Work using aseptic techniques. You should also include two control plates (see Note 21). 5. Incubate the inverted plates overnight at 37 °C. Carefully examine the plates the next day for colonies.

3.6 Screening by PCR (Colony PCR)

1. Depending on how many colonies have grown on your selection plate(s) and how many you wish to screen, take the same number of universal (20 mL) and microcentrifuge (1.5 mL) tubes. Label them “A,” “B,” “C,” etc., so that each universal tube matches one microcentrifuge tube. 2. To each universal tube add 5 mL of the liquid SOC media. 3. To each microcentrifuge tube add 10 μL of nuclease-free water. 4. Using a sterile pipette tip, pick a single colony from the selection plate. Dip the tip into the labeled microcentrifuge tube containing 10 μL nuclease-free water and swirl for 10 s. Take the pipette tip out of the microcentrifuge tube and drop it into the universal tube containing liquid SOC media with the matching label. Repeat this step for every colony you wish to screen using a different set of labeled microcentrifuge/universal tubes (see Note 22). Set the universal tubes on the side of your bench. 5. Collect all of the used microcentrifuge tubes and spin them in a centrifuge for 2 min at maximum speed (15,000× g). You should observe a pellet and supernatant. 6. Label the same number of PCR tubes to match one of the microcentrifuge tubes as “A,” “B,” “C,” etc. To each PCR tube add 2.5 μL of the supernatant from the corresponding microcentrifuge tube. 7. To each PCR tube add 22.5 μL of the previously prepared PCR master-mix. 8. Close the lid of PCR tubes and place them in the thermocycler. Run the 3-step standard PCR cycling under the previously set program outlined in Table 1 with the number of cycles increased to 40.

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19

9. To visualize the PCR samples on an agarose gel, refer to Subheading 3.1. 10. A positive colony-PCR result with a single clear band of the expected size means you have successfully cloned your gene— congratulations! For the example of results (see Note 23). 11. To confirm that the insert is correct and that there are no unforeseen mutations in it, you must send a sample of the cloned plasmid for sequencing. So, for PCR samples where a positive result was observed, collect the corresponding universal tubes (with the tip and SOC media) and incubate them overnight at 37 °C with constant shaking at 225 rpm (see Note 24). 12. Use a plasmid minipreparation kit to purify your plasmid containing the cloned DNA; use the manufacturer’s instructions.

4

Notes 1. Before any wet lab work decide on the purpose of your cloning experiment. Selection of the cloning vector, restriction enzymes, and competent bacterial cells will depend upon the further applications of the cloned sequence of interest. Here we aim to clone Escherichia coli DNA ligase A (ligA) gene (2016 kb) using directional cloning (two restriction enzymes) into an expression vector (pET-28b), which will be transformed into competent cells capable of protein production (BL21(DE3)) to obtain the 671aa monomer. For guidance on choosing a vector and competent cells appropriate for your experimental purpose (see Notes 4 and 7). 2. When designing your primers refer to the guidelines outlined in Table 2. You can also use Fig. 1 as a visual aid. 3. Annealing temperature depends on the melting temperature (Tm) of the primers used. You should aim for 5 °C below the calculated Tm and for it to be in the range of 45–68 °C. Annealing temperature can be optimized by performing an annealing screen: a temperature gradient PCR starting 5 °C below the calculated Tm. 4. When selecting a cloning vector consider the following qualities: (i) It must possess an origin of replication. (ii) A high replication rate is desirable to obtain a high copy number per cell cycle of a transformed cell. (iii) It should possess a range of restriction enzyme recognition sites to clone into. It must possess the restriction site for the selected restriction enzymes to be used for target DNA

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Table 2 Primer design instructions Recommended (Do)

Not recommended (Do not)

Make primers 15–30 bases in length

Multiple di-nucleotide repeats or long runs of single bases (maximum of 4 bases)

Aim for 40–60% GC content with uniform distribution

Long stretches of G or C repeats

Include one C or G base at the 3′ end

No more than three Cs or Gs in a row at the 3′ end

Tm of 55–70 °C, as close as possible between the Restriction site on the very end of the primer sequence: add a few bases before/after it to allow two primers and within 5 °C range. Use websites enzyme binding (such as OligoCalc [22]) to calculate the Tm of each primer Check if primers form any secondary structures (hairpins, etc.) Add restriction enzyme recognition sequences. Check that these sequences are not, by any chance, also within your DNA target

sequence insertion. Most cloning or expression vectors possess multiple cloning sites for this purpose. Selecting two different restriction enzymes ensures the DNA insert to be ligated is inserted in a predetermined direction (directional cloning). (iv) It should be comparatively small to the bacterial cells it will be transformed into. This allows for easier manipulation and cell transformation. (v) It needs to have one or more selectable marker genes, one of which must code for an enzyme (such as beta-lactamase) that inactivates the antibiotic added to the selection plates. (vi) Consider the purpose of your experiment. If you aim to express the protein encoded by the DNA insert, your vector must contain both a promoter sequence (T7 is the most common) and a ribosome binding site (RBS). Consider whether you will require a C- or N-terminal tag for later purification. 5. Both NdeI and BamHI introduce “sticky” (cohesive or asymmetric) ends, as opposed to the “blunt” (flush) ends. However, if you work with restriction enzymes that introduce “blunt” ends, dephosphorylate the vector to avoid a high background level of plasmid self-ligation. This can be done by adding CIP (calf alkaline phosphatase) or SAP (shrimp alkaline phosphatase) to the reaction following the restriction enzyme digestion step. The alkaline phosphatases can be inactivated by heating once their reaction is completed.

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21

6. If you wish to use a different buffer for the restriction digestion reactions, ensure it is compatible with both of the restriction enzymes used. 7. When deciding on the competent cells to be used, you should consider the downstream applications of the experiment, the transformation method you wish to perform (electroporation vs. chemical), the efficiency of the transformation method, and the genotype of the cell. Refer to Fig. 5 for guidance; this shows you a flowchart of which competent cells would be the most appropriate for your experiment. 8. Standard LB media can also be used, however, SOC media gives 2-fold higher transformation efficiency. 9. Selection plates can be used either warm or cold, wet or dry without affecting the transformation efficiency. However, it is easier to spread the bacterial colonies on warm, dry plates which allow for more rapid colony formation. 10. Here we add fresh kanamycin since that is the antibiotic against which pET-28b(+) vector carries a resistance gene. If you are using a different vector, add the same amount of the antibiotic that your vector is carrying the resistance gene against. 11. When preparing a PCR master-mix for multiple PCR reactions, multiply the volume of every component apart from the DNA template by the number of PCR reactions you wish to perform and add 1 μL to obtain the total volume of each component to be added in the master-mix. In this way, you will prepare a sufficient amount of the PCR master-mix for all the reactions you wish to perform and a small spare volume to allow for pipetting error. 12. To obtain a high-quality gel and avoid crystal formation in the gel, filter the 10× TBE as you are adding it to the conical flask by using a 15-mL syringe and a sterile 0.22 μM polyethersulfone (PES) filter. 13. Before pouring the 1× TBE buffer into the gel tank, add ethidium bromide to it. DNA is negatively charged, while ethidium bromide is positively charged. Therefore, EtBr will run in the opposite direction of the DNA when you run the gel. As a result, your gel will be differentially intense, with bright bands at the top (where the concentration of EtBr is high) and faded at the bottom. Adding EtBr to the buffer levels the intensity of the DNA bands on the resolved gel. If you do not add EtBr to the running buffer, you can soak the gel for an hour in the EtBr solution after it has been run and rinse with water. This will even out the staining and it can also be performed to stain the gel if you forgot to add EtBr to your gel in the first place.

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14. The recommended molar ratio of vector:insert DNA is 3:1 when cloning a typical-size insert into a plasmid vector that is larger than the insert DNA. However, the ratio varies on type of the vector and can be between 3:1 or 1:3. To calculate the amount of insert DNA to be added to achieve the desired ratio, use the following equation: ng of vector × kb size of insert insert × molar ratio of = ng of insert vector kb size of vector The recommended amount of DNA is 100–200 ng per ligation reaction. 15. Use the DNA concentration in the PCR sample reading taken by the spectrophotometer to calculate the volume of your sample that contains the target ng of the DNA insert. You can do so by applying the Beer-Lamber Law. 16. If the obtained amount of DNA insert from the PCR reaction is too low to achieve 100 ng of DNA in the 10 μL ligation reaction, scale up the reaction volume as necessary. Increase the volume of buffer added appropriately to achieve 1×. It is not necessary to increase the volume of T4 DNA ligase added, as 1 μL should be sufficient for larger volume reactions. 17. It is optional to heat-inactivate the T4 DNA ligase following the incubation step; if you do, heat it up to 65 °C for 10 min in a hot block, before cooling on ice. 18. A 2-fold loss in transformation efficiency should be expected for every 10 min you shorten this step. 19. Timing of this step is very important and can vary for the competent cells used. Refer to manufacturer’s instructions and perform this step exactly as instructed. 20. These are the optimal conditions for outgrowth to occur and achieve best cell recovery and expression of antibiotic resistance. Shaking or rotating gives a 2-fold increased transformation efficiency. For every 15 min you shorten this step, expect 2-fold loss in transformation efficiency. 21. Two control plates should be present: one that has competent cells only (which should not grow if the antibiotic is working) and another that has competent cells transformed with a positive control plasmid. A positive control plasmid is a widely used plasmid (pUC19, pBR322, or pBlueScript) that contains the same antibiotic resistance gene that is present in your media but does not contain your target DNA insert. If the competent cells transformed with a positive control plasmid grow, this confirms the cells are working.

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23

22. Performing this step creates a stock of bacterial cells containing only the cells identical to the picked colony. If the screening test shows a positive result, i.e., cells of this colony contain a plasmid with the DNA insert, you have successfully cloned your target DNA sequence and have a bacterial culture that can serve as a stock for further experiments. 23. An example of a positive colony PCR experiment conducted following the outlined protocol is shown in Fig. 6. 24. Screening the bacterial colonies by PCR amplification is the fastest method of verifying if the competent cells have successfully been transformed with a vector containing the insert. Alternatively, positive (expression of a lethal gene) or negative (blue-white screening) selection tests could be used. The most accurate screening method, and a step you should always include, is to send your cloned plasmid samples to be sequenced by Sanger dideoxy sequencing. When your sequencing results come back, carefully check every nucleotide position in your cloned insert to ensure you do not have unexpected mutations. References 1. Linn S, Arber W (1968) Host specificity of DNA produced by Escherichia coli, in vitro restriction of phage fd replicative form. Proc Natl Acad Sci U S A 59(4):1300–1306 2. Danna K, Nathans D (1971) Specific cleavage of Simian Virus 40 DNA by restriction endonuclease of hemophilus influenzae. Proc Natl Acad Sci U S A 68(12):2913–2917 3. Kellenberger G, Zichichi ML, Weigle JJ (1961) Exchange of DNA in the recombination of bacteriophage λ. Proc Natl Acad Sci U S A 47(6):869–878 4. Meselson M, Weigle JJ (1961) Chromosome breakage accompanying genetic recombination in bacteriophage. Proc Natl Acad Sci U S A 47(6):857–868 5. Jackson DA, Symons RH, Berg P (1972) Biochemical method for inserting new genetic information into DNA of Simian Virus 40: circular SV40 DNA molecules containing lambda phage genes and the galactose operon of Escherichia coli. Proc Natl Acad Sci U S A 69(10):2904–2909 6. Griffith F (1928) The significance of pneumococcal types. J Hyg 27(2):113–159 7. Cohen SN, Chang ACY, Hsu L (1972) Nonchromosomal antibiotic resistance in bacteria: genetic transformation of Escherichia coli by R-factor DNA. Proc Natl Acad Sci U S A 69(8):2110–2114

8. Cohen SN, Chang ACY, Boyer HW, Hellingt RB (1973) Construction of biologically functional bacterial plasmids in vitro. Proc Natl Acad Sci U S A 70(11):3240–3244 9. Preston A (2003) Choosing a cloning vector. Methods Mol Biol 235:19–26 10. Carter M, Shieh J (2015) Chapter 10 – Molecular cloning and recombinant DNA technology. In: Carter M, Shieh J (eds) Guide to research techniques in neuroscience, 2nd edn. Academic, New York, pp 219–237 11. Mertz JE, Davis RW (1972) Cleavage of DNA by R1 restriction endonuclease generates cohesive ends. Proc Natl Acad Sci U S A 69(11): 3370–3374 12. Green MR, Sambrook J (2020) Cloning in plasmid vectors: directional cloning. Cold Spring Harb Protoc 2020(11):485–488 13. Upadhyay A, Upadhyay K (2009) Recombinant DNA technology. In: Basic molecular biology. Himalaya Publishing House, Global Media, pp 452–506 14. Fakruddin M, Mohammad Mazumdar R, Bin Mannan KS, Chowdhury A, Hossain MN (2013) Critical factors affecting the success of cloning, expression, and mass production of enzymes by recombinant E. coli. ISRN Biotechnol 2013:1–7 15. Hanahan D, Jessee J, Bloom FR (1991) Plasmid transformation of Escherichia coli and

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other bacteria. Methods Enzymol 204 (C):63–113 16. Dower WJ, Miller JF, Ragsdale CW (1988) High efficiency transformation of E. coli by high voltage electroporation. Nucleic Acids Res 16(13):6127–6145 17. Manna S, Harman A, Accari J, Barth C (2013) Altering the selection capabilities of common cloning vectors via restriction enzyme mediated gene disruption. BMC Res 6(1):1–9 18. Nicholl DST (2002) Selection, screening and analysis of recombinants. An introduction to genetic engineering. Cambridge University Press, Cambridge, pp 132–150 19. Padmanabhan S, Banerjee S, Mandi N (2011) Screening of bacterial recombinants: strategies

and preventing false positives. In: Brown G (ed) Molecular cloning – selected applications in medicine and biology. In Tech, Rijeka 20. Sambrook J, Russell DW (2001) Molecular cloning: a laboratory manual, vol 1, 3rd edn. Cold Spring Harbor Laboratory Press, New York 21. Plasmids 101 A Desktop Resource Plasmids 101: A Desktop Resource (3rd edn) (2017). Retrieved November 13, 2021, from www. addgene.org 22. Kibbe WA (2007) OligoCalc: an online oligonucleotide properties calculator. Nucleic Acids Res 35. (Web Server issue)

Chapter 2 A Sequence- and Ligation-Independent Cloning (SLIC) Procedure for the Insertion of Genes into a Plasmid Vector Robert A. Holland Abstract Molecular cloning is a routine technique for many laboratories with applications from genetic engineering to recombinant protein expression. While restriction-ligation cloning can be slow and inefficient, ligationindependent cloning uses long single-stranded overhangs generated by T4 DNA polymerase’s 3′ exonuclease activity to anneal the insert and plasmid vector prior to transformation. This chapter describes a fast, high-efficiency protocol for inserting one or more genes into a vector using sequence- and ligationindependent cloning (SLIC). Key words SLIC, PCR, Cloning, T4 polymerase, Genetic engineering, Protein expression

1

Introduction The generation of recombinant DNA by molecular cloning is widely performed in biotechnology laboratories for applications such as heterologous protein expression and genetic engineering. Traditional cloning involves generation of “sticky ends” on the vector and insert using restriction enzymes, removal of 5′ phosphate by alkaline phosphatase, and ligation, making it timeconsuming and inefficient [1]. With the widespread use of polymerase chain reaction (PCR) beginning in the early 1990s, a number of ligase-free techniques of were developed [2–4]. Ligationindependent cloning (LIC) specifically utilizes the 3′ exonuclease (proofreading) ability of T4 DNA polymerase and complementary primers designed to lack one of dATP, dTTP, dCTP, or dGTP for a sequence of 10–12 bp. Once amplified by PCR, the vector and insert are treated separately with T4 polymerase and just the dNTP which is missing from the sequence, generating single-stranded complementary overhangs of fixed length, which can then be annealed and transformed directly into E. coli, which repair the “nicked” DNA backbone.

Garry Scarlett (ed.), DNA Manipulation and Analysis, Methods in Molecular Biology, vol. 2633, https://doi.org/10.1007/978-1-0716-3004-4_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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Fig. 1 General concept of SLIC. The gene of interest is amplified using primers with 5′ sequences (typically 15–40 bp) complementary to the insertion site within the vector. Incubation of the resulting PCR product with T4 DNA polymerase generates complementary overhangs. Similarly, the vector is linearized either by restriction digestion or PCR and overhangs generated by T4 DNA polymerase. When mixed, the insert and vector anneal to form a nicked plasmid which can be transformed directly into competent cells where gaps in the sequence are repaired

A progression of LIC procedure is sequence- and ligationindependent cloning (SLIC) [5–7], whereby specific sequences are replaced by longer complementary overhangs (15–40 bases). Following exonuclease treatment, the resulting insert-vector constructs contain gaps in the sequence which are subsequently repaired inside the host cell by recombination machinery (Fig. 1) [8, 9]. The flexibility, directionality, and high efficiency of SLIC and similar techniques have been used for the linking together of multiple fragments into chimeric sequences several kilobases long or the insertion of affinity tags for downstream protein purification [1, 6, 10]. While several proprietary enzyme mixes are available commercially, they can often be prohibitively expensive for production of large number of clones. This chapter details our costeffective and high-efficiency protocol for the cloning of a plasmid construct containing one or more inserts using 15 bp complementary overhangs.

Sequence and Ligation-Independent Cloning

2

27

Materials

2.1 Molecular Biology

1. Thermocyler. 2. Primers – (sense & antisense) for each insert and vector (see Note 1). 3. NanoDrop 2000 spectrophotometer. 4. Thin-walled 200 μL PCR tubes. 5. Deoxynucleotide triphosphate mix (dNTPs) – 2.5 mM each (see Note 2). 6. Nuclease-free H2O. 7. Phusion High-Fidelity DNA Polymerase & 5× HF buffer. 8. Template DNA for insert. 9. Plasmid vector (see Note 3). 10. MinElute PCR cleanup kit (Qiagen). 11. Fast digest DpnI (Thermo Scientific). 12. T4 DNA polymerase (NEB) (see Note 4).

2.2

Gel Analysis

1. Tris-acetate EDTA (TAE) 50× stock: dissolve 242 g tris base, 57.1 mL glacial acetic acid, and 100 mL 0.5 M EDTA pH 8.0, topped up to 1 L with deionized H2O. 2. 1% agarose gel: TAE, 10 g/L agarose. 3. Ethidium Bromide. 4. 6× DNA loading dye. 5. DNA ladder (1 kbp/100 bp). 6. Gel tank. 7. UV transilluminator suitable for agarose gel visualization.

2.3 Plasmid Preparation

1. Water bath. 2. Ultracompetent E. coli DH5α cells. 3. Lysogeny broth (LB): 10 g/L tryptone, 5 g/L yeast extract, 10 g/L NaCl, pH 7.0. 4. LB agar plates: LB, 10 g/L agar, 1/1000 dilution antibiotic stock (see Note 5). 5. Antibiotics: kanamycin (50 mg/mL) or ampicillin (100 mg/ mL) or chloramphenicol (34 mg/mL). 6. Shaking incubator. 7. Qiaprep Spin Miniprep Kit (Qiagen).

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Method

3.1 PCR Amplification of Insert and Vector

See Note that unless specified, all steps should be carried out on ice. 1. Dissolve the primers in nuclease-free H2O, aiming for a final concentration of 50–100 μM. Measure the exact concentration using the NanoDrop 2000 spectrophotometer. 2. In 200 μL PCR tubes (one for each insert or vector), prepare 150 μL reaction mixture for the insert(s): 30 μL 5× Phusion buffer, 0.5 μM forward primer, 0.5 μM reverse primer, 200 μM dNTPs (each), 0.2–0.5 ng/μL DNA template, 0.02 U/μL Phusion polymerase, topped up with nuclease-free H2O, and mix thoroughly by pipetting. Split each reaction mixture between 6× 25 μL reactions in separate PCR tubes. Reactions can be overlaid with mineral oil (see Note 6). 3. Place each of the six reaction tubes (see Note 7) containing insert or vector with equal spacing across the thermocycler and program the thermocycler as follows: 1 cycle Initial denaturation, 98 °C, 10 s, 1 cycle 25 cycles Denature, 98 °C, 10 s Anneal, 50–70 °C gradient, 30 s. Extension, 72 °C, 15–30 s per kb (see Note 8). 1 cycle Final extension, 72 °C, 5 min. 4. While the PCR amplification is running, set up the gel tank with a 1% agarose gel with TAE buffer. Load 5 μL from each reaction tube, each mixed with 1 uL 6× DNA loading dye, into separate wells of a 1% agarose gel, flanked by DNA ladder, and run at 100 V, 30 min (for 100 mL gel). Visualize the gel using a UV transilluminator and decide on the optimal annealing temperature for vector and insert (see Note 9). 5. Scale up the amplification by repeating steps 2 and 3, except with all 25 μL reactions at the optimal annealing temperature decided on in step 4. 6. Pool reactions from each amplicon and remove 10 μL for further analysis. Add DpnI and 10× fast digest buffer (5 μL each per 50 μL) and incubate in the thermocycler at 37 °C for 10 min, followed by 4 °C (see Note 10). 7. Purify the amplicons using a PCR reaction cleanup kit, following the manufacturer’s instructions, ensuring the product is eluted in nuclease-free H2O. Measure DNA concentration using a Nanodrop 2000 spectrophotometer. 8. Mix insert and vector in a molar ratio of 2.5:1 (0.0625 pmol insert & 0.025 pmol vector), 1× T4 polymerase buffer and 0.5 U T4 polymerase, topped up 10 μL with nuclease-free

Sequence and Ligation-Independent Cloning

29

H2O. Incubate for 10 min at 25 °C then place on ice. Use immediately for transformation and store the remainder at 20 °C. 3.2 Transformation and Sequencing

1. In separate 15 mL polypropylene tubes, mix 2.5 μL of each ligation reaction with 50 μL chemically competent DH5α cells and incubate 10 min on ice. Meanwhile, pre-warm a water bath to 42 °C and LB to 37 °C. 2. Heat shock the transformation mixtures by incubating at 42 °C for 45 s then immediately transfer to ice. 3. Add 1 mL warm, sterile LB to each transformation mixture and incubate in an orbital shaking incubator at 37 °C, 220 rpm, 60 min. 4. Pipette 100 μL of each transformation mixture onto LB agar plates with appropriate antibiotics and spread evenly (see Note 11). Incubate overnight (16 h) at 37 °C. 5. Count the number of colonies on the plates and calculate the transformation efficiency (see Note 12). Pick a number of colonies (at least four) from each plate and use to inoculate 10 mL sterile LB containing appropriate antibiotics. Incubate in an orbital shaking incubator at 37 °C, 220 rpm, overnight (16 h). 6. Transfer cultures to 15 mL polypropylene centrifuge tubes and pellet cells at 3000 g, 10 min, 4 °C in a benchtop centrifuge. Being careful not to disturb the pellet, pour off the supernatant. 7. Purify plasmids from the cell pellet using the QIAprep Miniprep Kit. Check concentrations using a spectrophotometer and send a sample of each for sequencing (see Note 13). 8. Run an agarose gel to compare samples from each cloning step from PCR amplification to final plasmid. Ensure plasmid and vector are linearized to properly compare sizes (Fig. 2).

4

Notes 1. Primers should be approximately 40 bp, including a minimum of 15 bp overlap with its complementary primer. To maximize annealing of insert and plasmid, ensure melting temperature (Tm) for each overhang is between 60–75 °C. Free-to-use primer design tools are available on the websites and provide information on melting temperature and GC content. 2. Aliquot dNTPs and store at -20 °C to avoid multiple freeze thaw cycles.

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Fig. 2 Example gel visualizing cloning steps. This 8 kb vector had two bands corresponding to circular and supercoiled plasmid. Vector linearization by PCR removed a 500 bp sequence, resulting in a band of a lower molecular weight. Recombination of linearized vector with a molar excess of 600 bp insert yielded a high cloning efficiency, indicated by a single band at approximately 8 kb. The low molecular weight contaminant was the same size as the insert, so it would not affect transformation

3. Vector propagated in a laboratory E. coli cloning strain such as DH5α is preferable in order for removal of the template after PCR amplification (see Note 10). 4. Kits commercially available for the annealing step however most rely on T4 polymerase exonuclease activity. 5. Check antibiotic resistance genes conferred by the chosen vector before preparing plates, e.g., KanR (kanamycin), AmpR (ampicillin), pp-cat (chloramphenicol). 6. Prior pre-linearization of the vector outside of the sequence to be amplified by restriction digest reduces the likelihood of sense and antisense primers annealing and forming a circular primer-vector PCR product. Ensure that the chosen restriction enzyme has no other sites within the vector sequence to be amplified; otherwise, the amplification will fail. An additional purification step using a reaction cleanup kit such as MinElute (Qiagen) may be required to remove enzyme activity and exchange the template DNA to an appropriate buffer for PCR. 7. Mineral oil can be used to stop evaporation; however, this can be minimized by using a thermocycler with a heated lid. 8. If not using Phusion polymerase, refer to the manufacturer’s instructions for the recommended amplification parameters.

Sequence and Ligation-Independent Cloning

31

Extension times increase with length of template to be amplified so it is crucial to perform amplifications for vector and insert(s) separately. 9. Optimal annealing temperatures may vary from the melting temperatures of each primer. Lower temperatures are more likely to result in incorrect product formation from nonspecific annealing of primers to the template or primer-dimers. As a rule of thumb, the highest temperature which yields a high level of amplification at the correct size should be used to minimize nonspecific interactions. 10. DpnI specifically digests methylated DNA. 11. If number of transformants is low, the number of transformants can be increased by plating the whole transformation mixture. Immediately prior to plating, centrifuge the transformation mixtures in a 1.5 mL microcentrifuge tube at 13,000 rpm, remove the supernatant and resuspend the pellet in a smaller volume, i.e., 100 μL LB. 12. The transformation efficiency is defined as the number of colony forming units on a plate per unit of DNA (cfu/μg DNA). 13. Sequencing is extremely important to determine whether the insert has been correctly cloned into the vector. Sanger sequencing is sufficient for inserts up to ~1000 bp, after which the quality is poor; in this case, reverse sequencing is recommended. Check with your sequencing provider that they stock a sequencing primer for your chosen plasmid, e.g., the T3 RNA polymerase promoter used in many expression vectors; otherwise, you may need to order one to send with your plasmid constructs. References 1. Celie PH, Parret AH, Perrakis A (2016) Recombinant cloning strategies for protein expression. Curr Opin Struct Biol 38:145– 154. Available from: https://linkinghub. elsevier.com/retrieve/pii/S0959440X16300 677 2. Shuldiner AR, Scott LA, Roth J (1990) PCR-induced (ligase-free) subcloning: a rapid reliable method to subclone polymerase chain reaction (PCR) products. Nucleic Acids Res 18(7):1920–1920. Available from: https:// academic.oup.com/nar/ar ticle-lookup/ doi/10.1093/nar/18.7.1920 3. Aslanidis C, de Jong PJ (1990) Ligationindependent cloning of PCR products (LIC-PCR). Nucleic Acids Res 18(20): 6069–6074. Available from: https://

academic.oup.com/nar/ar ticle-lookup/ doi/10.1093/nar/18.20.6069 4. Tillett D, Neilan B (1999) Enzyme-free cloning: a rapid method to clone PCR products independent of vector restriction enzyme sites. Nucleic Acids Res 27(19):26e–26. Available from: https://academic.oup.com/ nar/article-lookup/doi/10.1093/nar/27.1 9.e26 5. Li MZ, Elledge SJ (2007) Harnessing homologous recombination in vitro to generate recombinant DNA via SLIC. Nat Methods 4(3):251–256. Available from: http://www. nature.com/articles/nmeth1010 6. Stevenson J, Krycer JR, Phan L, Brown AJ (2013) A practical comparison of ligationindependent cloning techniques. PLoS One

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8(12):e83888. Available from: http://www. pubmedcentral.nih.gov/articlerender.fcgi? artid=3871625&tool=pmcentrez& rendertype=abstract 7. Scholz J, Besir H, Strasser C, Suppmann S (2013) A new method to customize protein expression vectors for fast, efficient and background free parallel cloning. BMC Biotechnol 1 3 ( 1 ) : 1 2 . A v a i l a b l e f r o m : h t t p s : // bmcbiotechnol.biomedcentral.com/ar ti cles/10.1186/1472-6750-13-12 8. Kostylev M, Otwell AE, Richardson RE, Suzuki Y (2015) Cloning should be simple: Escherichia coli DH5a´-mediated assembly of

multiple DNA fragments with short end homologies. PLoS One 10(9):e0137466 9. Jeong J-Y, Yim H-S, Ryu J-Y, Lee HS, Lee J-H, Seen D-S et al (2012) One-step sequence- and ligation-independent cloning as a rapid and versatile cloning method for functional genomics studies. Appl Environ Microbiol 78(15): 5440–5443. Available from: https://journals. asm.org/doi/10.1128/AEM.00844-12 10. Gibson DG, Young L, Chuang R-Y, Venter JC, Hutchison CA, Smith HO (2009) Enzymatic assembly of DNA molecules up to several hundred kilobases. Nat Methods 6(5):343–345. Available from: http://www.nature.com/arti cles/nmeth.1318

Chapter 3 Molecular Cloning Using In Vivo DNA Assembly Sandra Arroyo-Urea, Jake F. Watson, and Javier Garcı´a-Nafrı´a Abstract Here we describe the in vivo DNA assembly approach, where molecular cloning procedures are performed using an E. coli recA-independent recombination pathway, which assembles linear fragments of DNA with short homologous termini. This pathway is present in all standard laboratory E. coli strains and, by bypassing the need for in vitro DNA assembly, allows simplified molecular cloning to be performed without the plasmid instability issues associated with specialized recombination-cloning bacterial strains. The methodology requires specific primer design and can perform all standard plasmid modifications (insertions, deletions, mutagenesis, and sub-cloning) in a rapid, simple, and cost-efficient manner, as it does not require commercial kits or specialized bacterial strains. Additionally, this approach can be used to perform complex procedures such as multiple modifications to a plasmid, as up to 6 linear fragments can be assembled in vivo by this recombination pathway. Procedures generally require less than 3 h, involving PCR amplification, DpnI digestion of template DNA, and transformation, upon which circular plasmids are assembled. In this chapter we describe the requirements, procedure, and potential pitfalls when using this technique, as well as protocol variations to overcome the most common issues. Key words Molecular cloning, In vivo DNA assembly, recA-independent recombination, IVA cloning, Sub-cloning, Site-directed mutagenesis

1

Introduction Molecular cloning is a cornerstone of biomedical research and has been continuously developed over recent decades to provide simpler and more efficient methodologies. Here we describe the use of the in vivo DNA assembly approach for molecular cloning, which relies on an endogenous E. coli DNA recombination pathway capable of joining linear DNA fragments with short homology regions at their termini [1]. This pathway is independent of recA and is present in all laboratory E. coli strains. The “recA-independent” recombination pathway has been exploited for molecular cloning, facilitating cloning procedures with minimal handling but having the advantages of recombination-based methodologies (scarless, sequence-independent, single-base precision, and directional) [1–

Garry Scarlett (ed.), DNA Manipulation and Analysis, Methods in Molecular Biology, vol. 2633, https://doi.org/10.1007/978-1-0716-3004-4_3, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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7]. Using in vivo DNA assembly, all standard plasmid DNA modifications can be performed, including sequence insertions, deletions, point-mutagenesis, and sub-cloning of large fragments between vectors. When employed as a cloning tool, linear DNA fragments are generated in vitro, primarily by PCR, with homologous termini of around 15–30 bp that direct plasmid assembly in vivo. After transformation, endogenous single-stranded exonucleases (ExoIII/V) [8] degrade the termini of these linear fragments to single-stranded DNA, which allows annealing between homologous fragments in vivo, before DNA repair (LigA) assembles a circular plasmid [9, 10]. Cloning protocols generally comprise the following: (1) primer design, (2) PCR amplification, to introduce modifications and homologous sequences, (3) digestion of the parental DNA using the methylase-dependent restriction enzyme DpnI, and (4) transformation into standard laboratory E. coli before subsequent colony screening and selection. This protocol can be used to perform all types of plasmid modifications, from inserting and deleting sequences to site-directed mutagenesis and sub-cloning, each dictated by primer design. Furthermore, up to six DNA fragments can be assembled simultaneously, allowing complex cloning strategies to be achieved in a single step; however, method efficiency decreases as procedure complexity increases. Given the principal requirement for cloning using recombination is linear DNA fragments, in vivo DNA assembly can also be combined with restriction enzyme-linearized plasmids or synthesized linear double-stranded genes, which can overcome PCR amplification issues or further simplify procedures (Fig. 1). Given that there are no requirements for commercial kits or specialized bacteria, this approach is accessible to any molecular biology laboratory. Additionally, since the homology requirements are similar to other enzyme or recombination-based commercial approaches (e.g., Gibson assembly [11]), primers designed for in vivo assembly cloning can also be combined/used with such in vitro assembly methods as an alternative backup route. Here, we describe reagents and protocols to perform in vivo DNA assembly for plasmidic DNA cloning and modification as performed in our laboratory.

2

Materials

2.1 Polymerase Chain Reaction

1. Premixed PCR master solution, prepared as 23 μL premixed reactions, stored at -20 °C (see Note 1): 250 μM each dNTP nucleotide (dATP, dGTP, dCTP, and dCTP), 1 M betaine, 2.5% DMSO, Phusion Polymerase Buffer (1X), Phusion HF Polymerase (1 μL per 25 μL reaction) (see Note 2), and deionized H2O (we generally use 200 μL thin wall PCR tubes). 2. PCR Thermocycler.

Molecular Cloning Using In Vivo DNA Assembly

35

Fig. 1 General in vivo DNA assembly scheme. A DNA fragment of interest resulting from PCR amplification or gene synthesis can be assembled in vivo into a plasmid (linearized as a result of a PCR amplification or through restriction-enzyme digestion) after transformation in standard E. coli

2.2 DNA Gel Electrophoresis and Eliminating Parental DNA

1. Agarose powder (see Note 3). 2. DNA stain (e.g., SYBR™ Safe DNA Gel Stain (Thermo Fisher) or GreenSafe Premium (Nzytech)). 3. 10X TB agarose electrophoresis buffer: 440 mM Tris, 440 mM Boric acid in water. Weigh 54 g of Tris and 27.5 g Boric acid, and dissolve by stirring in 1 L H2O. We do not use EDTA in the TB buffer as compared to the commonly used TBE recipe (see Note 4). 4. DNA Gel Loading dye. 5. DNA Molecular Weight Marker Ladder. 6. DNA Gel Electrophoresis Tank and power supply. 7. UV/Blue light Transilluminator. 8. DpnI (FastDigest, Thermofisher) (see Note 5).

2.3

Transformation

1. Chemically competent bacteria: XL-10 Gold® Ultracompetent cells (see Note 6). 2. Water bath (set to 42 °C). 3. Super Optimal Broth (SOB): 20 g bactotryptone, 5 g yeast extract, 2 mL of 5 M NaCl, 2.5 mL of 1 M KCl, 10 mL of 1 M MgCl2, 10 mL of 1 M MgSO4, and distilled H2O to 1 L. 4. Lysogeny Broth (LB): 7.5 g agar, 5 g tryptone, 5 g NaCl, 2.5 g yeast extract, and distilled H2O to 500 mL. 5. LB agar plates with antibiotics as appropriate. 6. 37 °C incubator.

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3.1.1

Sandra Arroyo-Urea et al.

Methods Primer Design

Insertion

We recommend using software for visualizing both the original and target DNA sequences for the design of oligos, as well as software for the calculation of annealing temperatures (Tm). We use the freely accessible Snapgene Viewer program for primer sequence design and the OligoCalc webserver (http://biotools.nubic. northwestern.edu/OligoCalc.html) for annealing temperature calculation [12]. All Tm values reported in this chapter are calculated using this webserver. Accurate primer design is critical to the success of in vivo DNA assembly. Regardless of the modification to be made (insertion, deletion, mutagenesis, or sub-cloning), primers consist of two regions: a 3′ region that anneals to the template DNA (template binding region) and a 5′ homologous region that drives in vivo recombination. First, design the template binding region, which has the same requirements of standard PCR oligo design (at least 18–22 bp and Tm values of ~60 °C). The homologous region is included 5′ to this sequence, and it should be ≥15 bp and have a Tm ≥ 50 °C (see Note 7). As a rule of thumb, a homologous region of ~20–25 bp is sufficient to ensure efficient recombination (usually providing a Tm ≥ 50 °C), and lengths of up to 35 bp can been used to enhance efficiency when assembling ≥5 DNA fragments simultaneously. Specific primer design requirements for each DNA modification are as follows: We define an insertion as the introduction of a new segment of DNA that can be fully included within a single pair of PCR primers (independent of size) and are typically up to 200 bp. Using one pair of PCR primers, the whole vector is amplified during PCR, and recircularization occurs after transformation through a single recombination event. To insert a DNA fragment to a plasmid, design primers with the template binding regions binding astride the insertion site and add the homologous regions at the 5′ ends. If the length of the insertion is around ~20 bp, this new sequence can form the homologous region by inclusion at the 5′ end of both primers (Fig. 2a). If the insertion is significantly larger than a typical homologous region (>30 bp), the desired sequence should be divided in two, with each primer encoding half of the total insert. Insert coding regions on each primer must be designed to overlap (~20 bp) to act as the homologous region (Fig. 2b) (see Note 8). If the new sequence is significantly shorter than ~20 bp, the 5′ end of one primer will need to be extended to overlap with the annealing region of other primer in order to create a larger homologous region that guarantees an efficient recombination (Fig. 2c).

Molecular Cloning Using In Vivo DNA Assembly

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Fig. 2 Primer design to perform insertions. Primers must have template binding regions (grey) that bind either side of the insertion site (red) and a homologous region at the 5′ end (orange box) that allows in vivo recombination. Based on the length of the desired insertion (depicted in green), the homologous regions for each of the primers could be: (a) the entire insert (when the insertion is ~20 bp), (b) a sub-region of the insert (when the insertion is >25–30 bp), or (c) extended beyond the insert sequence alone (1.7 fold amplification per cycle, should this not be the case (see Note 3).

2.3 Pilot EP-PCR Experiment

Table 1 Average number of mutations per DNA template as function of length and number of doublings Template length EP-PCR doublings Mutations per nucleotide position 100 bp 200 bp 400 bp 800 bp 1600 bp 5

0.0033

0.33

0.66

1.3

2.6

5.3

10

0.0066

0.66

1.3

2.6

5.3

11

20

0.013

1.3

2.6

5.3

11

21

30

0.020

2.0

4.0

7.9

16

32

50

0.0033

3.3

6.6

13

26

53

84

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F. A. Myers

Method

3.1 Mutagenesis of Homogenous Starting Sequences

This protocol is based on the Caldwell & Joyce Method and uses a 500 bp homogenous DNA template region which is run through 15 PCR cycles. 1. Make up the following PCR reaction mixture in 100 μl PCR tube on ice: – 10 μl 100 mM Tris-HCl, pH 8.3 (10 mM final) – 2.5 μl 2 M KCl (50 mM final) – 3.5 μl 200 mM MgCl2 (7 mM final) – 4 μl 25 mM dCTP, pH 7 (1 mM final) – 4 μl 25 mM dTTP, pH 7 (1 mM final) – 4 μl 5 mM dATP, pH 7 (0.2 mM final) – 4 μl 5 mM dGTP, pH 7 (0.2 mM final) – 2 μl 100 mM 5′ primer (2 mM final) – 2 μl 100 mM 3′ primer (2 mM final) – X μl DNA template (3 nM of target site, final) (see Notes 3 and 4). – 2 μl 25 mM MnCl2 (0.5 mM final) (see Note 5) – 1 μl 5 U/μl Taq DNA polymerase (0.05 U/μl final). Make up to 100 μL with DNAse free water (Molecular Biology Grade). 2. Place the tube into the thermocycler and perform ~15 PCR cycles (see below). The cycling conditions will vary depending on the template and the primers, but generic conditions are: 94 °C for 1 min (denaturation), 60 °C for 1 min (annealing), and 72 °C for 3 min (extension) (see Note 6). 3. The final PCR product should be run on an ethidium bromide agarose gel with a suitable size marker to check that the correct sized amplicon has been achieved. 4. Clone and sequence a sample of the PCR DNA to determine frequency of mutations in the amplicon products using the TOPO T/A cloning kit and a commercial sequencing company (see Notes 7, 8, and 9).

3.2 Mutagenesis of Heterogenous Starting Sequences

Sometimes, it may be desirable to mutate a whole collection of sequences, for example, in a library, at the same time. In such cases, the method should be modified as below (see Note 10). 1. Make up the following EP-PCR reaction on ice: – 150 μl 100 mM Tris-HCl, pH 8.3 (10 mM final) – 37.5 μl 2 M KCl (50 mM final)

PCR Mutagenesis

85

– 52.5 μl 200 mM MgCl2 (7 mM final) – 60 μl 25 mM dCTP, pH 7 (1 mM final) – 60 μl 25 mM dTTP, pH 7 (1 mM final) – 60 μl 5 mM dATP, pH 7 (0.2 mM final) – 60 μl 5 mM dGTP, pH 7 (0.2 mM final) – 30 μl 100 μM 5′ primer (2 μM final) – 30 μl 100 μM 3′ primer (2 μM final) – 960 μl DNase free water 2. Divide the EP-PCR reaction mixture into 16 (90 μl aliquots) labeled PCR tubes. 3. Add 7 μl of 30 ng/μl DNA library to tube 1 to give ~2 ng/μl. Place tube 1 in the thermal cycler. Once it has reached the annealing temperature add the following and mix: – 2 μl 25 mM MnCl2 (0.5 mM final) – 1 μl 5 U/μl Taq DNA polymerase (0.05 U/μl final) 4. Run the reaction for four cycles of the EP-PCR (use conditions described in basic protocol above). During the final extension at 72 °C, place a second tube with fresh EP-PCR into the PCR block to warm. Transfer 10 μl from the first tube into the second tube and then add the following and mix to the second tube: – 2 μl 25 mM MnCl2 (0.5 mM final) – 1 μl 5 U/μl Taq DNA polymerase (0.05 U/μl final). 5. Repeat step 4 for the remaining 14 tubes. Analyze the PCR reactions on agarose gel electrophoresis after every transfer and quantitate the bands in successive PCR amplifications.

4

Notes 1. There are now many high-fidelity alternatives to standard Taq; although preferred for many molecular biology applications, these must not be used for Error-prone PCR protocols. 2. The initial optimization step can take anywhere between 1 and 2 days. The actual EP-PCR experiment takes up to 1 day to perform depending on how many cycles are being used. Cloning steps take 2–3 days. Preparing the samples for sequencing a further day and the sequencing data is obtained between 5 and 10 days, depending on the sequencing provision used. Analysis of the sequencing data takes 1–2 days. 3. Low PCR efficiency is most often the consequence of poor primer design. Both 5′ and 3′ primers should be designed with Tm’s 60 °C, a length of between 20 and 25 base pairs the 3′ end should be a C or a G.

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4. In general, higher concentrations of starting template will reduce the mutation rate. 5. The amount of template DNA should be calculated in terms of PCR target site. Assuming the template was just the target sequence, then in our example, this would equate to 100 ng total in the reaction. However, often the target sequence is part of a plasmid, and this should be taken into consideration. 6. Manganese salts can precipitate out of solution. It is critical to add the MnCl2 just prior to the initiation of the PCR reaction. 7. The oligonucletide primers should be designed so that the annealing temperature is in the region of 60 °C to avoid mis-priming, which has an enhanced likelihood in this protocol due to the high divalent cation concentration. The longer 3-min extension reduces the selection of shorter sequences produced by mis-priming. 8. There are a number of different cloning approaches; the simplest for this application is one of the commercially available TA cloning kits. We use either TOPO TA from Thermofisher or pGEM-T Easy from Promega. 9. There are numerous suppliers of out-sourced Sanger sequencing; we use either Sigma or Genewiz and costs are normally in the £5 per reaction range. Information on how to send the samples varies with providers but can be found on their specific websites. To view the raw data, you will need a chromatogram viewer such as the free download FinchTV (https:// digitalworldbiology.com/FinchTV). 10. If lower levels of mutagenesis are observed than desired, more EP-PCR cycles will be performed. If these need to go >15 cycles, a fresh aliquot of Taq should be added after the 15th cycle is complete. 11. The basic protocol requires a small volume of starting template. However, this may be insufficient to preserve the initial library complexity if this is the starting material. In this case to avoid any losses, start with a large template concentration and perform four cycles of EP-PCR. The product should then be taken and 10% of this used in a fresh EP-PCR reaction. This serial transfer approach should be performed until the desired number of doublings has taken place. References 1. Leung DW, Chen E, Goeddel DV (1989) A method for random mutagenesis of a defined DNA segment using a modified polymerase chain reaction. Technique 1:11–15 2. Wilson DS, Keefe AD (2000) Random mutagenesis by PCR. Curr Protoc Mol Biol 2000(8):3

3. Cadwell RC, Joyce GF (1992) Randomization of genes by PCR mutagenesis. PCR Methods Appl 2:28–33 4. McCullum EO, Williams BAR, Zhang J, Chaput JC (2010) Random mutagenesis by error-prone PCR. Methods Mol Biol 634:103–109

Chapter 8 In Vitro Site Directed Mutagenesis Michael J. McClellan Abstract Site-Directed Mutagenesis (SDM) allows for changes in the DNA sequence of plasmids using polymerase chain reaction (PCR). It is a reliable, accessible, and rapid method which is the common initial step of many biochemial or genetic experiments. Here we describe the various different forms of SDM before giving a detailed method for the introduction of substitutions, insertions, or deletions using a fast, ligation-free protocol, followed by colony PCR to screen for mutated sequences. Key words Site-directed mutagenesis, Polymerase chain reaction, Mutation, Genetic alteration, Plasmid manipulation, Colony PCR

1

Introduction Site-Directed Mutagenesis (SDM) describes a number of methods which modify the DNA sequence of plasmids. SDM, in one form or another, has existed from the late 70s [1], after which a wide array of modified protocols arose to introduce varying types of mutation, and improving the speed and efficiency of the protocol, such as the addition of steps to deplete unmodified DNA from the reaction as pioneered by Kunkel [2] amongst others. In common, all methods that bear the name use primers to target a specific sequence and introduce a mutation, followed by PCR cycling to fix and amplify those mutations, which can include one or multiple base substitutions, deletions, and insertions. In this regard it is a distinct method from more modern forms of site-directed mutagenesis such as CRISPR [3–5], the development of which in no way obsoletes classic SDM. The applications of SDM are truly vast. Almost any manipulation of plasmid DNA outside of restriction and cloning can be performed by SDM, from inactivating resistance genes to creation or deletion of restriction sites [6–8]. The introduction of point mutations or deletions into proteins in expression vectors is a common first step in studying the phenotype of many genetic

Garry Scarlett (ed.), DNA Manipulation and Analysis, Methods in Molecular Biology, vol. 2633, https://doi.org/10.1007/978-1-0716-3004-4_8, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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perturbations, be it structural or functional [9–14]. Outside of coding regions, the manipulation of enhancers or promoters in reporter plasmids by SDM allows the study of transcriptional control, via removing, modifying, or creating transcription factor binding sites [15, 16]. The major steps are the selection of the desired mutation, the design of primers to effect and target the mutation, the introduction and amplification of the mutation by PCR, the repair of the plasmid either by transformation into bacteria or ligation, and finally the selection of the correct sequence by cloning and sequencing. There are a number of highly reliable SDM kits provided by numerous suppliers, including NEB and Agilent, each coming with their own excellent set of instructions and aids. Here we will describe a fast SDM method which uses PCR to introduce point mutations, deletions, or insertions and staggered nicks to allow for transformation into bacteria without ligation. To improve the efficiency of the reaction, wild type template DNA carrying methylated dam sites is digested by DpnI (therefore the plasmid to mutate must be purified from dam+ bacteria). Colony PCR is used to accelerate the protocol further such that a purified, mutated plasmid can be obtained within 24 h.

2 2.1

Materials SDM PCR

1. Q5 DNA polymerase and buffer (New England Biolabs) (see Note 1). 2. SDM primers (see Subheading 3.2). 3. Plasmid purified from dam+ bacteria and eluted in water or elution buffer. 4. 10 mM dNTP mix.

2.2 Digestion and Transformation

1. DpnI and rCutsmart buffer (New England Biolabs). 2. XL-10 GOLD ultracompetant cells (Agilent) (see Note 1). 3. 14-mL snap cap Falcon polypropylene round-bottom tubes. 4. S.O.C. medium (ThermoFisher). 5. Water bath at 42 °C. 6. Agar plates with Antibiotic appropriate for plasmid.

2.3 Colony PCR and Sequencing

1. Q5 DNA polymerase and buffer (New England Biolabs) (see Note 1). 2. Primers designed to amplify mutated region. 3. 10 mM dNTP mix. 4. LB broth with appropriate antibiotic.

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Methods Carry out all procedures at room temperature unless otherwise specified.

3.1 Selection of Mutations and Design of Primers

1. Select bases to mutate; as many as three can be easily changed in one reaction, above which efficiency may start to drop off. 2. Bases to mutate should all be within 30 bp of each other. Short deletions or insertions may also be introduced; however, any more than three bases begins to reduce the yield of mutated plasmid, as does deleted/inserted bases being non-consecutive. If multiple changes of this type are required, it may be best to do sequential rounds of mutagenesis. 3. Design primers, which are the direct complements of each other, targeting the region of plasmid you wish to mutate. 4. Primers do not require 5′ phosphorylation, and standard desalting purification is sufficient. 5. The melting temperature of primers should be high, >~75°, to allow for mismatches and should possess G or C “clamps” at either end, although this is not essential (see Note 2). 6. The primers should be complimentary to the plasmid sequence except at the bases which are to be mutated (Fig. 1). No mutated base should be less than 10 bp away from one end of the primer. 7. The minimum size of the primers should be 30 bp to allow sufficient distance between staggered nicks on the plasmid (Fig. 2) to facilitate annealing. Both primers should contain all mutated bases. 8. Larger primers can be used when targeting multiple sites, although this increases the risk of secondary structure formation.

3.2

SDM PCR

1. Make a 50 μL PCR mix containing the following: 10 μL 5× Q5 buffer X μL (1–50 ng) of dsDNA template (see Note 3) 1 μL 10 mM dNTP mix (see Note 4) 0.5 μL (1U) Q5TM High Fidelity DNA Polymerase 2.5 μL of 10 μM Forward SDM primer 2.5 μL of 10 μM Reverse SDM primer X μL H2O to make up to 50 μL 2. Cycle the reaction in a thermocycling PCR machine as per Table 1.

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Fig. 1 Schematic of primer design for the three basic forms of SDM, substitution, insertion, and deletion. In each case, the primers are the exact complement of each other, and both primers contain the desired change in sequence. Primers are also long, with a high Tm, and the mutations are central

Fig. 2 Introduction of mutation by PCR and the depletion of unmutated plasmid by DpnI digestion. Plasmid, isolated from dam+ bacteria, will carry methylated adenine on both strands at all GATC sequences (blue DNA), whereas newly synthesized DNA, extended from primers carrying the mutated sequence will not possess any methylated bases (black DNA). Multiple rounds of PCR will enrich DNA carrying the mutation relative to the methylated parental DNA. Four possible products exist after the final round of extension, all, excepting plasmid with a staggered nick and carrying the desired mutation on both strands, will be digested by DpnI, which selectively targets methylated and hemi-methylated GATC sequences

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Table 1 Thermocycling conditions for SDM PCR Stage

Cycles

Temperature

Time

1

1

98 °C

30 s

15–25

98 °C 55–60 °C 72 °C

10 s 10 s 30 s/kb

1

72 °C

1 min/kb

2

3

3.3 Digestion of Template DNA and Transformation

1. Add 1 μL (20 U) of DpnI, 6 μL of rCutsmart buffer and 3 μL H2O. Mix by pipetting and incubate reaction at 37 °C for 1 h. This step eliminates plasmid that does not contain mutated bases on both strands (Fig. 2). 2. Pre chill 14 mL snap cap Falcon polypropylene round-bottom tubes (1 per reaction). 3. Gently thaw XL-10 GOLD ultracompetent cells on ice (50 μL per reaction). 4. Prewarm S.O.C. medium and Agar plates with appropriate antibiotic to 37 °C (see Note 5). 5. Mix 50 μL of XL-10 GOLD ultracompetent cells with 2–10 μL of DpnI digested DNA in a prechilled 14 mL round bottom falcon tube, mix by pipetting, and incubate on ice for 10 min (see Note 6). 6. Submerge the bottom half of each falcon tube in 42 °C water for 40 s, transfer back into ice, and incubate for 2 min (see Note 7). 7. Add 500 μL prewarmed S.O.C. medium to each falcon and incubate shaking (225 rpm) at 37 °C for 1 h to allow bacteria to recover (see Note 8). 8. To ensure that there are isolated colonies to pick the next day, add 5 μL, 50 μL, and 500 μL of the transformation mix to prewarmed Agar plates with the appropriate antibiotic for the plasmid. For 5 μL and 50 μL, pipette the transformation mix into a pool of 495 μL and 450 μL, respectively, of prewarmed S.O.C. medium to facilitate spreading (see Note 9). 9. Spread the liquid evenly across the plate using a cell spreader or plating beads and invert the plate before incubating overnight at 37 °C.

3.4 Colony PCR and Sequencing

1. Select primers to amplify (and later sequence) the region containing the introduced mutation. These should not be more than 500 bp apart, and neither primer should be within 20 bp of the mutated bases to ensure high sequencing quality at the critical point in the read (see Note 10).

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Table 2 Thermocycling conditions for colony PCR Stage

Cycles

Temperature

Time

1

1

98 °C

30 s

30

98 °C 55–60 °C 72 °C

10 s 10 s 30 s/kb

1

72 °C

2 min

2

3

2. Prepare one 25 μL PCR mix for each colony to sequence containing the following: 5 μL 5× Q5 buffer 0.5 μL 10 mM dNTP mix 0.25 μL (0.5 U) Q5TM High Fidelity DNA Polymerase 2.5 μL of 10 μM Forward sequencing primer (see Note 11) 2.5 μL of 10 μM Reverse sequencing primer (see Note 11) 39.25 μL high purity water to make up to 50 μL 3. Pick 3–6 large and isolated colonies from any of the 3 plates. Attach a pipette tip to a P20 pipette and, using the tip, scrape half of the colony off the plate and submerge in 25 μL of PCR mix, pipette up and down, and move tip around the inside of the tube to dislodge the colony into solution. Clearly annotate and circle each colony chosen on the back of the plate and store at 4 °C (see Note 12). 4. Cycle the reaction in a thermocycling PCR machine as per Table 2. 5. Either purify PCR product or directly send PCR mix for purification and sequencing depending on your sequencing provider. 6. Once the correctly mutated colony is identified, pick the other half of the appropriate colony using a pipette tip and drop it into LB broth to inoculate. Grow overnight ready for mini- or midi-scale plasmid preparation.

4

Notes 1. Many polymerases are appropriate for SDM but must lack strand displacement or 5′–3′ exonuclease activity and must use a buffer that is compatible with DpnI activity (information which can be found on the New England Biolabs website). Q5 is listed here as it is an economical and high-fidelity polymerase

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exhibiting the required characteristics. Pfu polymerases are also appropriate. Likewise, XL-10 GOLD are an example of an abundance of possible dam+ competent cells. 2. GC clamps are short sequences of G and/or C bases at the ends of primers, which, due to their higher melting temperature, have been shown to improve PCR efficiency. If it is not possible to achieve this at both ends of the primer, the 3′ end, from which extension will occur, has priority. There are also plenty of online SDM primer design tools which reliably produce working primers. 3. The amount of plasmid DNA to use can be modified depending on the size of the plasmid. Between 1 and 50 ng is appropriate for plasmids under 5 kb; for larger plasmids, up to 100 ng can be used. Multiple reactions can be set up each with a different amount of template plasmid to ensure colonies are obtained. 4. As with any PCR reaction, it is possible to optimize the amount of primer to dNTP to template ratio; however, in SDM PCR reactions, it is recommended that primers be kept in constant excess. Cycle number can also be optimized but usually between 15 and 25 cycles will produce colonies. 5. The following steps can be performed next to a Bunsen burner (to provide updraft) or in a lateral flow hood to minimize the chances of contamination; however, with antibiotic resistance in the plate, this is not strictly necessary. To control for contamination, add an extra sample to which no DpnI treated SDM PCR reaction is added. If a similar number of colonies are obtained on this plate as those that received plasmid, stop the experiment and repeat but prepare all reagents fresh. 6. To control for transformation efficiency, transform 2–20 ng of the unmodified plasmid alongside the mutated plasmids. If no colonies are obtained on SDM transformed plates and colonies are seen on the control plate, it is most likely due to inefficient PCR amplification as a result of primer mispriming or secondary structure. To avoid unnecessary delays, we usually design multiple primer pairs per mutation, varying length, and position of the primers. PCR conditions and input amount of DNA can also be altered to improve SDM PCR efficiency. 7. If the number of transformations is limited, it is possible to heat shock bacteria without a water bath. Take a thermometer and fill a beaker with hot water from the tap. Slowly run cold water into it whilst stirring with the thermometer until it reads 42 °C, stop the tap, and heat shock the bacteria in the beaker. Do not transform bacteria in a heat block as transformation efficiency will dramatically decrease.

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8. This recovery incubation step is not necessary if the resistance gene is for Ampicillin. 9. As the plasmid is nicked rather than supercoiled, transformation efficiency will be variable and much lower than usual, hence the wide range of plating volumes. Plating can also be performed next to a Bunsen or in a hood (see Note 5), but usually being quick with replacing the plate lid and the antibiotic embedded in the agar will suffice to prevent contamination. To reduce the number of plates used it is also possible to recover in a smaller volume of S.O.C. and add to a single plate then spread to single colonies by streaking. 10. It is possible that the plasmid gains additional mutations apart from the desired ones, either from unrepaired DNA damage or Polymerase error. Depending on the downstream application, it may be necessary to sequence the entire plasmid. 11. It is not necessary to use the same primers to amplify the mutated region as for sequencing, but sequencing primers must always be within the region amplified during colony PCR. 12. If the next day sequencing is available, simultaneously inoculate LB broth (plus antibiotic) with each colony chosen for colony PCR. The next day, when sequencing results arrive, a mini- or-midi prep may be performed directly from the culture grown from the appropriate colony, discarding the rest. References 1. Gillam S, Smith M (1979) Site-specific mutagenesis using synthetic oligodeoxyribonucleotide primers: II. In vitro selection of mutant DNA. Gene 8(1):99–106 2. Kunkel TA, Roberts JD, Zakour RA (1987) Rapid and efficient site-specific mutagenesis without phenotypic selection. Methods Enzymol 154:367–382 3. Jinek M, Chylinski K, Fonfara I, Hauer M, Doudna JA, Charpentier E (2012) A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity. Science 337: 816–821 4. Cho SW, Kim S, Kim JM, Kim JS (2013) Targeted genome engineering in human cells with the Cas9 RNA-guided endonuclease. Nat Biotechnol 31:230–232 5. Doudna JA, Charpentier E (2014) Genome editing. The new frontier of genome engineering with CRISPR-Cas9. Science 346(6213): 1258096 6. Marini F 3rd, Naeem A, Lapeyre JN (1993) An efficient 1-tube PCR method for internal site-

directed mutagenesis of large amplified molecules. Nucleic Acids Res 21(9):2277–2278 7. Liu H, Ye R, Wang YY (2015) Highly efficient one-step PCR-based mutagenesis technique for large plasmids using high-fidelity DNA polymerase. Genet Mol Res 14(2):3466–3473 8. Luna S, Mingo J, Aurtenetxe O, Blanco L, Amo L, Schepens J, Hendriks WJ, Pulido R (2016) Tailor-made protein tyrosine phosphatases: in vitro site-directed mutagenesis of PTEN and PTPRZ-B. Methods Mol Biol 1447:79–93. https://doi.org/10.1007/9781-4939-3746-2_5. PMID: 27514801 9. Adereth Y, Champion KJ, Hsu T, Dammai V (2005) Site-directed mutagenesis using Pfu DNA polymerase and T4 DNA ligase. Biotechniques 38(6):864, 866, 868 10. Wu D, Guo X, Lu J, Sun X, Li F, Chen Y, Xiao D (2013) A rapid and efficient one-step sitedirected deletion, insertion, and substitution mutagenesis protocol. Anal Biochem 434(2): 254–258

SDM 11. Liu H, Naismith JH (2008) An efficient one-step site-directed deletion, insertion, single and multiple-site plasmid mutagenesis protocol. BMC Biotechnol 4(8):91 12. Monchietti P, Lo´pez Rivero AS, Ceccarelli EA, Catalano-Dupuy DL (2021) A new catalytic mechanism of bacterial ferredoxin-NADP+reductases due to a particular NADP+binding mode. Protein Sci 30(10):2106–2120 13. Harrison JJEK, Tuske S, Das K, Ruiz FX, Bauman JD, Boyer PL, DeStefano JJ, Hughes SH, Arnold E (2021) Crystal structure of a retroviral polyprotein: prototype foamy virus protease-reverse transcriptase (PR-RT). Viruses 13(8):1495

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14. Emruzi Z, Aminzadeh S, Karkhane AA, Alikhajeh J, Haghbeen K, Gholami D (2018) Improving the thermostability of Serratia marcescens B4A chitinase via G191V site-directed mutagenesis. Int J Biol Macromol 116:64–70 15. Liang Q, Chen L, Fulco AJ (1995) An efficient and optimized PCR method with high fidelity for site-directed mutagenesis. PCR Methods Appl 4(5):269–274 16. Nishizawa-Yokoi A, Yamaguchi N (2018) Gene expression and transcription factor binding tests using mutated-promoter reporter lines. Methods Mol Biol 1830:291–305

Chapter 9 Xenopus Transgenesis Using the pGateway System Liliya Nazlamova Abstract Transgenic approaches using I-SceI are powerful genome modification methods for creating heritable modifications in eukaryotic genomes. Such modifications are ideal for studying putative promoters and their temporal and spatial expression patterns in real time, in vivo. Central to this process is the initial engineering of a plasmid construct containing multiple DNA modules in a specific order prior to the integration into the target genome. One popular way of doing this is based upon the pGateway system, the modular form of which described in this chapter is known as pTransgenesis. We will initially describe the protocol of obtaining the plasmid construct containing the required sequence modules, and then the process of integrating the construct into the genome of a Xenopus embryo via co-injection with I-SceI and subsequent screening for transgenics. Key words Site-specific recombination, Plasmid cloning, Transgenesis, Xenopus

1

Introduction

1.1 Transgenesis as a Tool for Reporter Gene Assays

The term transgenic was first introduced in 1981 by John Gordon and Frank Ruddle when they successfully inserted exogenous genetic material into the genome of fertilized mouse eggs [1]. Since then, transgenesis has evolved into a widely used method, across a number of species for both commercial and academic purposes. Examples include the “spider goat” for spider silk production in goat milk and golden rice, which is fortified for the precursor to vitamin A [2, 3]. The first reported transgenic frog was made in 1984 by Laurence Etkin when he introduced, via microinjection, DNA constructs into one cell Xenopus laevis embryos [4]. In these early experiments, various constructs were injected as either linear or circular pieces of DNA into model organisms, such as Xenopus, Drosophila, and Sea Urchins. Circular plasmids were found to be integrated into the embryo genome less often than the linearized genetic material, but the main problem with the method was the highly mosaic expression pattern, where the plasmid was integrated into the genomes of only patches of

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cells. Further, gene expression often displayed mis-regulation, likely due to positional epigenetic influence [4]. However, since then Xenopus transgenesis has undergone extensive development, successful examples include the introduction of the Gal4/UAS system [5], ubiquitous expression of GFP [6], and specific eye expression under control of the γ-crystallin [7]. A modern method for creating transgenic frogs utilises the restriction meganuclease, I-SceI, avoiding many of the problems associated with the early experiments. In particular, the I-SceI method is not restricted by the DNA insert size and has a high survival rates of modified embryos. 1.2 The I-SceI Technique and the pGateway System

In order to produce a transgenic frog using the I-SceI technique, one-cell stage embryos are injected with a plasmid clone (Fig. 1), containing the insert of interest and a marker gene for indicating a successful transgenesis event. I-SceI is an endonuclease isolated from the yeast Saccharomyces cerevisiae that recognizes an 18-bp non-palindromic sequence that does not occur within the genomes of animal species but will linearize the plasmid within the cell, preventing recircularization by endogenous ligases. In our hands this method can give approximately 30% efficiency in generating F0 transgenic Xenopus. Key to the I-SceI transgenesis protocol is the assembly of the final injectable plasmid. In this chapter, we use a recombinase approach based upon the commercially available pGateway system to construct the initial plasmid and subsequently use this plasmid to generate a transgenic frog for characterization of a previously unknown promoter. The recombinase approach involves four vectors, each of which carries one of the modules (such as a putative promoter, GFP reporter) that will be combined to build the final cassette (Fig. 2). These vectors are based upon the pGateway plasmids which form the pGateway system [8, 9]. This is a modular system and is transferable between different species such as Drosophila, Zebrafish, Xenopus, and mammalian cells. An interesting aspect of the pGateway plasmids is the inclusion of the ccdB gene which encodes a gyrase inhibitor that is toxic to most E. coli strains. However, after successful recombination, this gene is removed from the plasmids as a by-product. Therefore, after bacterial transformation, only the bacterial clones containing the successful recombinants will survive, drastically reducing the background. Although the pGateway system provides non-recombinant plasmids, versions of the plasmids containing the required gene modules can be obtained easily from a number of sources. The ones in our example are available from European Xenopus Resource Center (EXRC) (https://www.port.ac.uk/research/research-pro jects/european-xenopus-resource-centre) and obviate the need for insertion of genes into the base plasmid; the relationship between the base plasmids and the ones obtained from the EXRC is shown

pGateway Transgenesis

I-SceI

γ-Crys/ GFP

Putat. promoter

Katushka

99

SAR-cHS4 I-SceI

Expression vector (p4) Amp r

+ I-SceI I-SceI γ-Crys/ GFP

Putat. promoter

Katushka

SAR-cHS4 I-SceI

Injecng the mix of I-SceI and the expression vector into one-cell Xenopus embryos

n= 100

• Injected embryos le to develop • Screen for the presence of green eyes

Fig. 1 Outline of the I-SceI transgenic procedure. A mix of I-SceI and an expression clone is injected into one-cell Xenopus laevis embryos. The expression vector carries internal control modules. These include flanking I-SceI restriction sites and 5′ and 3’ insulators (Tol2). A γ-crystalline promoter driving a GFP reporter gene (indicated in green) is used as a marker for later identification of successful transgenics. A second set of modules within the expression clone consists of the putative promoter to be tested (dark blue) and a second reporter gene Katushka (red). Depending on the properties of the putative promoter, red fluorescence may be observed at developmental stages

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attL4

attP1

ccdB Cm r attP2 pDonr221 vector Kan r

attL1

Putat. promoter attL2 Entry clone p2 Kan r

Γ-crys/ GFP attR1 Entry clone p1 Kan r

I-SceI Tol2

attB1

PCR product

attB1

attR4

attR2 Katushka Entry clone p3 Kan r

attR3 ccdB Cm r Destination vector p4 Amp r

I-SceI Tol2 B4 γ-crys/ GFP B1 Putat. promoter B2

attL3

Tol2 I-SceI

Katushka

B3 Tol2 I-SceI

Expression vector Amp r

Fig. 2 MultiSite gateway 3-fragment recombination reaction. An overview of the pGateway cloning procedure where the first step is PCR-amplification of a putative promoter construct to be recombined with the pDONR vector. The resulting new plasmid designated entry clone p2 (putative promoter) is recombined with p1 (Y-crystalline/GFP), p3 (Katushka), and destination vector p4 in order to produce the expression vector. The crossed lines represent recombination events between the designated att-sites and segments B1, B2, B3, and B4 represent the positions of the resulting recombination sites. The selected recombination att-sites in all of the plasmids allow for specific order of the cassette elements

in Table 1. In our example, vectors P1(γ-crystalline/GFP), P2 (putative promoter), and P3(Katushka) are recombined with pDest by LR Clonase® II in order to produce the final expression clone, pDest(final). The final product pDest(final) will possess all the necessary modules and elements required for transgenesis. The γ- crystallin promoter driving GFP expression will act as a marker for transgenic tadpoles and is adjacent to the promoter to be characterized which is itself driving the Katushka module. In the final plasmid construct, these modules will be flanked by a SARCH4 insulator and I-SceI meganuclease sites. When this plasmid is injected into Xenopus embryos, successful transgenesis will be confirmed by the frog’s eyes glowing green, as the γ-crystallin promoter will drive GFP expression specifically in the eye lens. If the putative regulatory element is functional, it will drive Katushka expression at defined developmental stages, and therefore, red

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Table 1 Plasmid nomenclature

Non-recombinant (commercial)

Non-recombinant short designation

Derived recombinants carrying the cassette modules

pDONR™

P1

P1 (γ-crystalline/GFP)

P2

P2 (putative promoter)

pDONR™ P2R-P3

P3

P3 (Katushka)

pDEST™ R4-R3 Vector II

pDest

pDest (final)

pDONR

™

221

A list and nomenclature of the plasmids used in the pGateway system—P1 (γ-crystalline/GFP) module serves as a selectable marker to allow for screening of successful transgenesis as GFP will be specifically expressed in the eye lens. P2 (putative promoter) contains the DNA sequence to be analyzed, and P3 (Katushka) carries the reporter gene Katushka. pDest is the destination vector, which carries the elements required for successful transgenesis: I-SceI sites, Tol2 sites, SAR- cHS4 insulators. SAR-CH4 insulator will isolate the integrated cassette from nearby functional elements that could potentially non-specifically regulate the expression of the reporter genes

fluorescence will be detected using a fluorescent microscope. The transgenic embryos can be monitored throughout the developmental stages and examined for fluorescent expression to determine temporal and spatial expression patterns of the unknown promoter in vivo.

2

Materials Ensure your laboratory is suitably registered and set-up for molecular biology work and wear a lab coat, gloves, and appropriate PPE at all times to avoid contamination and minimize the risk of nuclease degradation of samples. Use of live animals is strictly regulated at a national level jurisdiction; please be aware that you will require relevant regulatory permissions. Use molecular biology grade reagents and distilled, deionized water for preparing buffers.

2.1 Plasmid Recombination and Screening

1. pGateway kit (see Note 1). 2. P1(γ-crystalline/GFP) and P3(Katushka) constructs containing γ-crystalline and Katushka, respectively, are available upon request from EXRC https://xenopusresource.org/. 3. High-fidelity PCR mastermix. 4. Primers (see Note 2). 5. Thin-wall 0.2 mL polypropylene PCR tubes. 6. DNAse, RNAse-free 1.5 mL tubes. 7. PCR thermocycler set with the following program (Table 1). 8. DNA size ladder: a 100 base DNA ladder from a commercial supplier. Store at 4 °C.

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9. Sanger sequencing (see Note 3). 10. A commercial DNA gel extraction kit. 11. Ice and ice bucket. 12. Endofree Plasmid purification kit such as those available from Qiagen (see Note 4). 13. Competent bacteria. One Shot® TOP10 and ccdB resistant strains (see Note 5)—thaw on ice. 14. Agarose gel electrophoresis apparatus, reagents, and gel imaging system. 15. Microbiology designated bench and bacterial incubator set to 37 °C. 16. Freshly autoclaved LB broth, agar plates, and appropriate antibiotics (see Note 6). 17. Sterile 50 mL polypropylene tubes for liquid media growth. 18. Microbiology incubator-shaker set to 37 °C. 19. Spectrophotometer or optical-density reader set to 600 nm. 20. Sterile plastic 3 mL cuvettes for measuring OD600. 21. Microcentrifuge tubes (1.5 mL). 22. Benchtop refrigerated cooled-centrifuge capable of 16,000 × g and chamber temperature of 4 °C. 23. Appropriate restriction enzymes. 24. Molecular biology grade, nuclease-free water. 2.2 Transgenesis in Xenopus and Selection of Transgenic Embryos

1. Ice and ice bucket. 2. 50 mL Falcon tubes. 3. Nuclease-free 1.5 mL tubes. 4. Microinjector such as the equipment made by Harvard apparatus and calibration capillary tubes. 5. I-SceI and associated buffers (see Note 7). 6. Petri dishes. 7. Fertilized Xenopus embryos (see Note 8). 8. Modified Barth’s Saline (MBS) 10× stock (see Note 9). 9. 4% Ficoll—4 g Ficoll in 100 mL 1× MBS and stir until dissolved. 10. 2% Cysteine—2 g in 100 mL distilled water, then pH to 7.8 with 5M NaOH. 11. Light and fluorescent microscope. 12. A small-volume UV spectrophotometer (for example, a NanoDropTM or PicoDropTM device).

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Methods

3.1 Plasmid Recombination and Screening

1. PCR amplify the promoter of interest to be recombined with pDONR 221. Typically, we would use 10 μM each forward and reverse primers and 50 ng fosmid template (see Note 10). 2× PCR mastermix of a high-fidelity DNA Polymerase; however, for amplifying DNA fragments ≥3 kb, we use Platinum SuperFi II PCR master mix (Thermofisher). A suggested PCR program is shown in Table 2, although this will need to be optimized for the specific DNA template and Polymerase used (see Note 11). 2. Once the PCR is completed, analyze 5 μL on 1% agarose gel and compare to a suitable sized DNA marker to confirm the size of expected amplicon (see Note 12). 3. Gel extract the PCR amplicon using a suitable gel extraction kit such as those from Qiagen or NEB following manufacturer’s instructions (see Note 13). 4. The purified DNA was again analyzed by an agarose gel electrophoresis and its concentration measured by UV spectroscopy, prior proceeding with the BP recombination set up (see Note 14). 5. The BP recombination is set up with an equimolar amount of attB PCR product and donor vector (P2) as follows: 50 fmoles P2 vector, 50 fmoles putative promoter element, 1 μL BP Clonase II, complete with TE buffer, pH 8 to 8 μL final volume and incubate overnight at 25 °C. 6. On the following day, add 1 μL Proteinase K and incubate the reaction at 37 °C for 10 min. 7. Transform 1–5 μL (approx. 50 ng/μL) of the BP reaction into DH5α competent cells and grow overnight on kanamycin (100 ng/μL) containing agar plates.

Table 2 PCR cycle conditions PCR Step

Temperature (°C)

Time (s)

Denaturation

95

60

Melting Annealing Elongation

95 55–70 °C 72

30 30

Final elongation

72

300

Hold

4

1

30

40 cycles

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8. Pick colonies and inoculate in 3–5 mL of L-Broth containing Kanamycin (100 ng/μL) and incubate overnight in a shaking incubator at 250 rpm, 37 °C for approx. 16 h. 9. Purify plasmid DNA from each of the grown bacterial colonies using a Mini-scale DNA plasmid prep kit such as Qiagen following the manufacturer’s instructions. 10. Screen for correct recombinant clones ideally using restriction enzymes which cut uniquely to the internal cloned sequence and also to the plasmid backbone. Cut in the region of 1 μg purified plasmid DNA. 11. Analyze the restriction digest reaction with Agarose gel electrophoresis. 12. Compare the observed fragment sizes to the predicted restriction fragment map using suitable plasmid analysis software such as ApE (https://jorgensen.biology.utah.edu/wayned/ ape/). 13. Once the successful recombinant is confirmed perform a Maxiscale DNA plasmid preparation kit such as Qiagen following the manufacturer’s instructions. 14. Perform sequencing analysis to validate the correct DNA sequence. 15. Once you have obtained P1(γ-crystalline/GFP) and p3, perform a Maxi-scale DNA plasmid preparation using a kit such as Qiagen following the manufacturer’s instructions. 16. Use pDest readily available from the kit. 17. The LR reaction is set up at 10 μL final volume with equal molar concentration of each recombinant plasmid-10 fmol of P1, P2(putative promoter) and P3(Katushka), and 20 fmol of the destination vector pDest. To calculate the precise DNA concentrations, use the equation shown previously (see Note 15). 18. On the next day, add 1 μL Proteinase K to the reaction and incubate at 37 °C for 10 min. 19. Transform 1–5 μL (approx. 50 ng/μL) of the LR reaction into ccdB-sensitive competent cells (such as One Shot® TOP10 supplied with the kit) and grow overnight on Ampicillin (100 ng/μL) containing agar plates. 20. Once you have the final recombinant destination vector re-transform the confirmed plasmid clone and perform an Endo-free Maxi preparation kit from Qiagen following the manufacturer’s instructions.

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1. Dilute the plasmid pDest(final) to 1 μg/μL with nuclease-free water, and confirm the concentration by UV spectroscopy. If incorrect adjust to 1 μg/μL. Then perform a second dilution with nuclease-free water to a final concentration of 100μg/μL, which is the working stock of the pDest(final). 2. Prepare a restriction digest reaction using I-SceI to linearize the plasmid as this ensures the cassette integrity prior integrating into the genome. The reaction is set up 40 min before the embryo injection begins (see Note 16). Mix 2 μL I-SceI buffer (10×), 7 μL plasmid DNA (100 ng/μL) (so total of 700 ng), 2 μL 10× BSA, and complete to 18 μL with nuclease-free water. 3. Transfer the 18 μL mix to a tube containing 2 μL I-SceI aliquot taken fresh out from the -80-storage freezer. 4. Incubate the reaction at 37 °C for 40 min. 5. Prepare 0.3× MBS buffer and set up microinjector needle, look at reference [10]. Adjust the microinjector to inject 5 nL total per injection over a 0.5–1 s range. 6. Dejelly(remove) embryos’ vitaline membrane with approx. 40 mL 2% L-Cysteine solution, pH 7.8–8 in a 50 mL falcon tube. Mix by very gentle inversion of the tube until the embryos pack tightly together for 5 to 10 min. 7. Wash with 40 mL 0.3× MBS at least 3 times and then return to a fresh petri dish filled with 0.3× MBS. 8. Pour 4% Ficoll in MBS to a dish layered with indented agarose or plastic mesh in order to hold the embryos in evenly arranged position for injections. 9. Transfer between 20 and 30 of the dejellied embryos using a Pasteur pipette with the end cut off (see Note 17) to the dish prepared in step 7. 10. Inject 5 nL linearized plasmid solution, into 1-cell stage Xenopus embryos. To obtain non-mosaic transgenesis, it is critical to start injecting as soon as possible before the first cell division and definitely before 2-cell stage. 11. After completing the injections, keep the embryos in a fresh petri dish with 4% fresh Ficoll solution at 12 °C for 2–4 h to allow them to recover. 12. Transfer embryos to a fresh petri dish containing 0.3× MBS and keep at 18 °C until the stage of interest. 13. Use a fluorescent microscope to check for Katushka fluorescent reporter expression. If positive, red glow should be observed within the appropriate tissue. 14. Final confirmation of a successful insertion can be obtained by tissue restricted green fluorescence in the eye at stage 40.

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Notes 1. The pGateway kit is available from Life Technologies and provides the required enzyme mixes, plasmids, and chemically competent cells. However, in the example given, P1 contains both γ-crystalline and GFP and P3 contains Katushka, which are available separately from the European Xenopus Resource Center upon request (https://xenopusresource.org/). Store LR and BP Clonases™ at -80 °C. After use return immediately back to -80 °C storage. 2. Gene-specific complementary regions of the primers can be designed according to standard rules. However, at the 5′ end, they would need to contain the correct recombination sites (tails) for each plasmid. In our example, successful recombination with entry vector pDONR221 would require primer tails -aatB1 (forward) and attB2 (reverse) sequences. pDONR221 specific primers: attB1 GGGGACAAGTTTGTACAAAAAAGCAGGCTNN[gene specific sequence] attB2 GGGGACCACTTTGTACAAGAAAGCTGGGTN[gene specific sequence]

3. Sequencing can be done in house if you have access to capillary electrophoresis equipment. If not a number of commercial companies provide this service, for example, such as Source Bioscience, UK. 4. Endofree maxi prep kit contains an additional step where you treat the plasmid sample with a reagent that neutralizes any residual endotoxin carry over from E.coli. Endofree maxi prep kit is required for increased survival of the injected Xenopus embryos. This is strongly recommended when introducing plasmids into live model organisms. 5. The ccdB gene is toxic for most E.coli competent bacterial strains; to propagate the ccdb containing vectors, a ccdB resistant E.coli strain is required, which must be purchased separately from Life technologies. However, this is only required if you wish to make stocks of the non-recombinant entry and destination plasmids, as opposed to using the finite quantities of plasmid supplied. Once the ccdB gene is replaced by the target sequence after successful recombination, standard One Shot® TOP10 top or DH5α can be used. 6. Weight out 25 g of LB broth powder (10 g/L tryptone, 5 g/L yeast extract, 10 g/L NaCl) in l L beaker. Make up to 1 L with distilled water and stir until fully dissolved. Transfer 250 mL to

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four clean Pyrex bottles. Sterilize each by autoclaving at 121 °C for 15 min with the cap loosely attached by autoclave indicator tape. Once cooled to ~60 °C, add appropriate antibiotic to final concentration. To grow bacterial clones after BP recombination, add Kanamycin to a final concentration of 50 μg/mL. To grow bacterial clones after LR recombination, add Ampicillin to a final concentration of 50 μg/mL. Flame the neck of each bottle when used. Label and store at 4 °C until needed. 7. To preserve the integrity of I-SceI, upon delivery aliquot into 2 μl stocks and keep at -80 °C until needed. Do not keep enzymes on ice for long periods of time. 8. Xenopus colonies require specialist husbandry, information and protocols or use of Xenopus for embryological experiments can be found in reference [10]. 9. To make 10× MBS solution, mix the following chemicals to the concentration shown below: – 88 mM NaCl – 1 mM KCL – 10 mM HEPES – 2.4 mM NaHC03 – 0.82 mM MgS04.7H2O – 0.33 mM Ca (N03)2.2H2O – 0.41 mM CaCl2.6H2O First weigh and dissolve NaCl, KCl, and HEPES in 800 mL distilled water. Then adjust the pH to 7.6 with 5 M NaOH. Finally, add the salts—MgS04.7H2O, Ca (N03)2.2H2O, and CaCl2.6H2O. Keep MBS 10× stock at 4 °C up to 30 days and check for salts precipitation before use. When needed dilute with distilled water to 0.1× MBS working solution and add 1 mL Pen/strep to final concentration of 5 U/mL penicillin and 5 μg/mL streptomycin. MBS salt concentrations are critical to embryo survival and proper development; the salt concentrations must be weighed precisely, and care should be taken when making MBS solution. 10. In this protocol, fosmid, containing the Xenopus putative promoter of interest, was used as a PCR template. European Xenopus Resource Center stores stock of fosmid Xenopus genome libraries which can be supplied upon request. 11. PCR primers need to be optimized to find optimal annealing temperatures according to standard PCR protocols. A range of polymerases are available, it is important to select a highfidelity polymerase, and if the target sequence is greater than 3 kb, it must also possess high processivity.

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12. Set up two 50 μL PCR reactions to be able to recover sufficient amount of PCR product after subsequent agarose gel extraction. Once confirmed PCR has produced correct size amplicon, load the left over 45 μL PCR reactions + gel loading buffer on a new 0.8–1% Agarose gel. To load ≥45 μL of reaction volume in a single well, use a 2 mm thick gel comb when casting 10 mm thick gel. 13. After melting the gel pieces, mix the 2 samples prior loading on the purification column from the gel extraction kit. Alternatively, tape 2 wells of the gel and load the 90 μL PCR in a single well composed of two taped wells. 14. DNA concentration and molar ratios are critical for the success of all the recombination reactions in this protocol. Extra care must be taken in the calculations and pipetting at this step. Do not exceed 250 ng total DNA mass per 10 μL of BP recombination reaction. Return BP Clonases™ back to -80 °C (ideally temporarily store Clonases on dry ice once you finish using them to preserve enzyme integrity). 15. Adjust the plasmid concentrations so you can add 1 μL of each plasmid per LR reaction. Do not exceed more than 300 ng of total plasmid mass in 10 μL LR reaction. Return the LR Clonases™ back to -80 °C storage or keep on dry ice. 16. Make sure the reaction is set up immediately before the embryo injections, this is a critical step and timings must be exact. Ideally start the I-SceI reaction 5 min before obtaining the embryos from the female frog. The best integration of DNA into the genome occurs during the first 20 min after fertilization; hence, the injections were done during this window of time. 17. Cut off the end of a 1 mL (blue) tip with a sterile sharp scalpel, do not use scissors as this will leave a jagged edge. Alternatively, you can use glass Pasteur pipette. References 1. Gordon JW, Ruddle FH (1981) Integration and stable germ line transmission of genes injected into mouse pronuclei. Science (New York, NY) 214(4526):1244–1246 2. Tang G et al (2009) Golden rice is an effective source of vitamin A. Am J Clin Nutr 89(6): 1776–1783 3. Williams D (2003) Sows’ ears, silk purses and goats’ milk: new production methods and medical applications for silk. Med Device Technol 14(5):9–11

4. Etkin L et al (1984) Replication, integration, and expression of exogenous DNA injected into fertilized eggs of Xenopus laevis. Differentiation 26(3):194–202 5. Hartley KO, Nutt SL, Amaya E (2002) Targeted gene expression in transgenic Xenopus using the binary Gal4-UAS system. Proc Natl Acad Sci 99(3):1377–1382 6. Kroll KL, Amaya E (1996) Transgenic Xenopus embryos from sperm nuclear transplantations reveal FGF signalling requirements during

pGateway Transgenesis gastrulation. Development (Cambridge, England) 122(10):3173–3183 7. Smolich BD et al (1993) Characterization of Xenopus laevis γ-crystallin-encoding genes. Gene 128(2):189–195 8. Hartley JL, Temple GF, Brasch MA (2000) DNA cloning using in vitro site-specific recombination. Genome Res 10(11):1788–1795

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9. Love NR et al (2011) pTransgenesis: a crossspecies, modular transgenesis resource. Development 138(24):5451–5458 10. Guille M (1999) Chapter 10: Molecular methods in developmental biology: Xenopus and Zebrafish. Zebrafish 127

Chapter 10 CRISPR/Cas9 Gene Disruption Studies in F0 Xenopus Tadpoles: Understanding Development and Disease in the Frog Anita Abu-Daya and Annie Godwin Abstract CRISPR/Cas9 has become the favorite method for gene knockouts in a range of vertebrate model organisms due to its ease of use and versatility. Gene-specific guide RNAs can be designed to a unique genomic sequence and used to target the Cas9 endonuclease, which causes a double-stranded break at the desired locus. Repair of the breaks through non-homologous end joining often results in the deletion or insertion of several nucleotides, which frequently result in nonsense mutations. Xenopus frogs have long been an excellent model organism in which to study gene function, and they have proven to be useful in gene-editing experiments, especially the diploid species, X. tropicalis. In this chapter, we present our protocols for gene disruption in Xenopus, which we regularly use to investigate developmental processes and model human genetic disease. Key words CRISPR/Cas9, Xenopus tropicalis, Xenopus laevis, Disease modeling, Gene knockout, MicroCT

1

Introduction Gene knockouts are a powerful tool for loss of function studies in development and disease. A mutation in a specific gene can be achieved by a double stranded break in a unique DNA sequence, which triggers cellular DNA repair mechanisms. Although homology-directed repair will reproduce the wild-type state [Chapter 11: A CRISPR/Cas-Based Method for Precise DNA Integration in Xenopus leavis Oocytes Followed by Intracytoplasmic Sperm Injection (ICSI) Fertilization], non-homologous end joining is far more common and frequently causes insertions or deletions (indels) of several bases, resulting in frame shift mutations and the degradation of the transcribed mRNA by nonsense-mediated decay [1–3]. In the past decade, several methods for digesting a specific genomic sequence in vivo have been developed, including

Garry Scarlett (ed.), DNA Manipulation and Analysis, Methods in Molecular Biology, vol. 2633, https://doi.org/10.1007/978-1-0716-3004-4_10, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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zinc-finger nucleases [4–6], transcription activator-like effector nucleases [7, 8], and CRISPR/Cas9 [9, 10]. Whereas the first two methods rely on designing a nuclease that specifically binds to the target DNA sequence, CRISPR/Cas9 uses the same endonuclease, the Cas9 protein, which is targeted to the desired genomic locus by specific guide RNAs. As it is significantly easier and cheaper to synthesize guide RNAs rather than proteins specific to the region, CRISPR/Cas9 has become the dominant technique for genome editing. Xenopus embryos are particularly well suited for gene-editing experiments. Injection of mRNA or DNA into the large, robust, externally developing eggs has long been used for overexpression experiments, and injections of Cas9 protein and guide RNAs is just as efficient [11, 12]. One potential complication of studying the F0 crispant generation is that cell division begins two hours after fertilization and goes on at the same time as the Cas9 endonuclease function, potentially resulting in highly mosaic mutant embryos. However, we have found that the method is so efficient in Xenopus that knockout phenotypes can be analyzed in founder embryos. This approach has recently been applied to understanding the function of genes involved in development and human disease [10, 13–15], as reviewed in [1, 16–18]. Two Xenopus species are commonly used in laboratories, X. laevis and X. tropicalis. Although the former is more established and easier to work with, its allotetraploidy genome complicates genetic analysis. X. tropicalis is a true diploid with one of the smallest tetrapod genomes and is the preferred model system for genetic analysis [19–24]. However, not all of X. laevis genes are duplicated, some are singletons, making genetic analysis of certain loci possible in the larger, more robust species [25]. The genomes of both species have been sequenced and are well annotated [25, 26]. The methods described here work in either species. In this chapter we describe the complete process involved in creating gene knockouts in Xenopus embryos. We will explain how to design guide RNAs to target the Cas9 endonuclease to the gene of interest, how to prepare single guide RNAs (sgRNAs) from oligonucleotides, and the injection of sgRNAs and Cas9 protein into fertilized eggs. We provide a rapid method for genotyping embryos to assess whether gene-editing was successful. Finally, we describe the application of MicroCT using the contrast reagent Phosphotungstic Acid to quickly assess tadpole morphology, as a first step in phenotyping mutant embryos.

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Materials Ensure your laboratory is suitably registered and set-up for molecular biology work and wear a lab coat, gloves, and appropriate PPE at all times to avoid contamination and minimize the risk of nuclease degradation of samples. Use of live animals is strictly regulated at a national level jurisdiction, please be aware that you will require relevant regulatory permissions. Use molecular biology grade reagents and distilled, deionized water for preparing buffers.

2.1 Target Identification, Oligonucleotide Design and Guide RNA Synthesis

1. Software facilitating the planning, visualization, manipulation, and simulation of DNA fragments (for example, SnapGene). 2. Oligonucleotides for sgRNA synthesis: the CRISPR Universal (AAA AGC ACC GAC TCG GTG CCA CTT TTT CAA GTT GAT AAC GGA CTA GCC TTA TTT TAA CTT GCT ATT TCT AGC TCT AAA AC) and Target specific oligonucleotide stocks, made up to 100 μM in nuclease-free water (see Note 1). Store at -20 °C. 3. Taq PCR mix (2×): 10 mM Tris–HCl, pH 8.6, 50 mM KCl, 1.5 mM MgCl2, 0.2 mM dNTPs, 5% (v/v) glycerol, 0.08% (w/v) IGEPAL CA-630, 0.05% (w/v) Tween-20, 25 Units/ ml Taq DNA Polymerase. It is usually bought from a commercial supplier (for example, GoTaq G2 DNA Polymerase, Promega). Store at -20 °C. 4. Nuclease-free water, molecular biology grade. 5. Thin-wall 0.2 mL polypropylene PCR tubes. 6. PCR thermocycler. 7. A commercial spin-column PCR purification kit, the SigmaSpinTM Sequencing Reaction Clean-Up Kit (Sigma-Aldrich) is recommended. Use as per the manufacturer’s instructions and store at 4 °C. 8. A small-volume UV spectrophotometer (for example, a NanoDropTM device). 9. DNA size ladder: a 1 kb plus DNA ladder from a commercial supplier (for example, New England BioLabs Ltd) is recommended. Store at 4 °C. 10. Heat block set to 37 °C. 11. 1× Tris-Borate-EDTA (89 mM Tris-HCl, 89mM Boric Acid, 2mM EDTA, pH 8.3). 12. Agarose, from a commercial supplier. 13. Ethidium bromide or an alternative DNA gel stain for electrophoresis.

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14. A gel-visualization system with image acquisition software for visualizing DNA in agarose gels, for example, the G:BOX F3 gel doc system (Syngene, Synoptics Ltd.) with GeneSys image acquisition software. 15. An in vitro transcription kit, the MEGAshortscriptTM T7 Transcription Kit (Invitrogen—ThermoFisher Scientific) is recommended. Store at -20 °C. 16. DNase and RNase free 1.5 mL microcentrifuge tubes. 2.2 Injection Delivery of CRISPR/Cas9 Constructs

1. Fertilised Xenopus eggs. The in vitro fertilization of eggs for microinjection can be achieved using either fresh crushed testes or frozen sperm. 2. A controlled temperature incubator (ideal range: 14–28 °C). 3. 2% L-Cysteine solution (Recommended: non-hydrochloride (168149), Sigma-Aldrich), pH 7.8 for dejellying eggs. Store the solution at 4 °C and use until precipitates are visible. 4. A 1× stock of Marc’s Modified Ringers (MMR) solution (0.1 M NaCl, 2 mM KCl, 1 mM MgSO4, 2 mM CaCl2, 5 mM HEPES, and pH 7.4) is used to prepare the 0.05× MMR (X. tropicalis) and 0.1× MMR (X. laevis) working stocks. OPTIONAL: Commercially available penicillin (10,000 U/mL) and streptomycin (10 mg/mL) solution can be added to improve the survival of early staged embryos (Nieuwkoop and Faber (NF) stages 1–41) [27]. Store at 4 °C. 5. Plastic petri-dishes lined with 1% agarose (in 0.05× MMR, Store at 4 °C) or glass dishes should be used to house earlystaged (NF1 – NF10) X. tropicalis embryos. It is further recommended at these stages that glass Pasteur pipettes are used for embryo manipulation. 6. A picoinjector (for example, Medical Systems PLI-100 Picoinjector (Harvard Apparatus)), air compressor (for example, PT5 Bambi Air Compressor), a stereomicroscope (for example, 2× magnification Nikon Stereomicroscope (SMZ800, C-W 10× B Eyepieces)), with reticule micrometer and a cold light source (for example, KL 2500 LED Lightsource (Schott)) are required for microinjection. 7. Borosilicate glass capillaries (Harvard Apparatus), used to make needles for microinjection. 8. Micropipet puller (for example, Sutter instrument, Model P-87 Flaming Brown Micropipette Puller) with an appropriate program (for example, 801 ms (Heat), 250 ms (Pull), 190 ms (Velocity), 160 ms (Time), adjust as required) to prepare microinjection needles from capillaries. 9. Fine watchmaker’s forceps (for example, Dumont no.5) to cut the end of needles.

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10. It is recommended users have an injection grid. These injection grids are created in 60 mm Petri dishes, with 8 mm nylon mesh sealed using chloroform. 11. 3% (w/v) Ficoll® (GE Healthcare) in 0.05× MMR (for Xenopus tropicalis) or 0.1× MMR (for Xenopus laevis). Store the solution at 4 °C, for up to 1 week. 12. Cas9 protein obtained from a commercial supplier (recommended: EnGen® Spy Cas9 NLS protein (M0646), New England Biolabs Ltd.). Store at -20 °C. 13. Nuclease-free water, molecular biology grade. 14. DNase and RNase free 1.5 mL tubes. 15. Tabletop microcentrifuge. 2.3 Analysis of Crispant Tadpoles: Genotyping

1. DNase and RNase free 1.5 mL tubes. 2. Heat block set to 56 °C. 3. Lysis Buffer: 50 mM Tris-HCl (pH 8.5), 1 mM EDTA, 0.5% [v/v] Tween-20 and Store at 4 °C. Add 100 μg/mL Proteinase K prior to each use. 4. Tabletop microcentrifuge. 5. Thin-wall 0.2 mL polypropylene PCR tubes. 6. PCR thermocycler. 7. Taq PCR mix (2×): 10 mM Tris-HCl, pH 8.6, 50 mM KCl, 1.5 mM MgCl2, 0.2 mM dNTPs, 5% (v/v) glycerol, 0.08% (w/v) IGEPAL CA-630, 0.05% (w/v) Tween-20, 25 Units/ ml Taq DNA Polymerase. Usually bought from a commercial supplier (for example, GoTaq G2 DNA Polymerase, Promega). Store at -20 °C. 8. Nuclease-free water, molecular biology grade. 9. Target specific primers, designed in steps 2.1-2.2 and 3.1-3.6, made up to 100 μM in nuclease-free water. Store at -20 °C. 10. 1× Tris-Borate-EDTA (89 mM Tris-HCl, 89 mM Boric Acid, 2 mM EDTA, pH 8.3). 11. Agarose, from a commercial supplier. 12. Ethidium bromide or an alternative DNA gel stain for electrophoresis. 13. A gel-visualization system with image acquisition software for visualizing DNA in agarose gels. 14. T7 Endonuclease I enzyme and reaction buffer commercially available from New England BioLabs Ltd.: 10 U. Store at -20 °C.

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2.4 Analysis of Crispant Tadpoles: Phenotyping

1. 0.2% (w/v) Ethyl-m-aminobenzoate (also known as tricaine or MS222), pH 7. 2. Plastic petri-dishes lined with 1% agarose (in 0.05x MMR, Store at 4 °C). 3. MEMFA (0.1 M MOPS pH 7.4, 2 mM EGTA, 1mM MgSO4, 4% Stabilized Formaldehyde (Acros Organics)). 4. Laboratory grade Methanol. 5. Nuclease-free water, molecular biology grade. 6. Phosphotungstic acid (PTA), commercially available, used at a working concentration of 1% PTA in nuclease-free molecular grade water. 7. Agarose, from a commercial supplier. 8. Hot melt glue and glue gun. 9. A stereomicroscope (for example, 2× magnification Nikon Stereomicroscope (SMZ800, C-W 10× B Eyepieces) and a cold light source (for example, KL 2500 LED Lightsource (Schott)). OPTIONAL: A high-resolution microscope (for example, Zeiss Axio Zoom.V16 Stereomicroscope and CL9000LED light source (Carl Zeiss Microscopy)). 10. Zeiss Xradia Versa 520 (Carl Zeiss Microscopy), imaged using the manufacturer’s software (Scout and Scan Reconstructor, Carl Zeiss Microscopy) and visualized using TXM3DViewer (Carl Zeiss Microscopy).

3

Methods Ensure all work complies with local ethical and animal scientific requirements. Conduct all steps at room temperature and use double-distilled water in solutions unless otherwise specified. Wear lab coat, gloves and appropriate PPE at all times to avoid nuclease degradation of samples and injury. The following protocol is broken down into a series of steps to guide the creation and analysis of mutant Xenopus tadpoles: 1. Designing guide RNAs. 2. Generating guide RNA from ssDNA templates. 3. Microinjection of guide RNA into Xenopus embryos. 4. Genotyping strategies to understand indel formation in crispant embryo genomic DNA samples. 5. General phenotyping strategies for gross morphological changes in crispant tadpoles.

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The first and most important step in CRISPR/Cas9 gene disruption studies is designing guide RNAs which will target Cas9 to the sequence of interest. There are many approaches to standardize the design and improve the efficiency of prospective gene-editing constructs; we successfully use the freely available CRISPRscan and inDelphi browser applications. CRISPRscan identifies appropriate sgRNAs in a given sequence, estimates the efficiency of cutting in vivo, and checks that the sequence is unique to minimize the possibility of off-target effects [28, 29]. On the other hand, the inDelphi mESC model has been shown to reliably predict frameshift frequency and precision events (range of indels) resulting from CRISPR/Cas9 genome editing [30–32]. 1. The gene of interest is first identified through the gene-pages presented on Xenbase [33] using the most up to date genome assembly (presently, X. tropicalis: v10 and X. laevis: v9.2). The features of each gene’s structure can be identified in JBrowse and modeled in a DNA analysis software facilitating annotation (for example, SnapGene®5.2.4). 2. Any chromosomal or gene duplication should be considered in the experimental design. See Note 2 for more details including how to approach rare situations where the gene is poorly annotated. When considering the CRISPR/Cas9 target sequence it is best to choose a coding region not located within 20 bp of the end of the exon due to the potential for splicing effects. If many sgRNAs are possible, we choose those predicted to have high mutagenic activity (CRISPRscan score >30), with no viable off-target events (test in Xenbase blast using the latest genome assembly version to make sure the sequence is unique), and a high frameshift frequency (inDelphi frameshift score >75% (mESC model)) [28, 30]. See Notes 3 and 4 for how to approach small exon genes, splice variants, highly repetitive regions of the genome, large deletions, and for recommended CRISPR/Cas9 experimental controls. 3. Species conservation (between Xenopus sp. and for example, H. sapiens) and key protein domains can be mapped from protein sequences stored in the National Centre for Biotechnology Information (NCBI) databases using a freely available multiple sequence alignment tool (for example, Clustal Omega (EMBL-EBI)). This information may guide targeting an appropriate region in the protein (5′UTR, first exon, early exon, key protein domain, patient variant site). See Notes 3 and 4 for more details. 4. It is recommended that three sgRNAs are designed from genomic DNA sequences encompassing the target region (c. 500 bp) to test in a preliminary experiment. This approach allows the selection of the most effective sgRNA and minimizes the likelihood the phenotype is due to off-target changes.

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5. The next point for consideration in the experimental design is the biological target. It is possible to gene-edit both the Xenopus egg (generation of mosaic homozygotes, as described in this chapter) and the Xenopus oocyte (if generation of non-mosaic heterozygotes is required, Chapter 11: A CRISPR/Cas9-Based Method for Precise DNA Integration in Xenopus leavis Oocytes Followed by Intracytoplasmic Sperm Injection (ICSI) Fertilization). Further, gene-editing in the egg makes it is possible to mutate the whole embryo or restrict editing to individual blastomeres (using the fate-map) up to the 32-cell stage (see Note 5). 6. Primers to amplify 500–800 bp of the genomic DNA surrounding the target region must be designed, using a freely available online tool, for example, Primer3 or PrimerBlast (see Note 6). These primers will be used in genotyping the potential mutant embryos. 3.2 Guide RNA Synthesis

For sgRNA synthesis we have had good success using the method described in Nakayama et al. [34] This relies on annealing two single-stranded oligonucleotides, the gene-specific nucleotide suggested by CRISPRscan, which contains a 5′ T7 promoter, and a Universal CRISPR Oligonucleotide, which contains the Cas9 binding sequence. The two oligonucleotides have an overlapping segment and can be annealed and extended by Taq polymerase to generate a DNA template from which the sgRNA can be transcribed with T7 polymerase. 1. Set up the annealing and extension reaction to convert ssDNA to dsDNA by mixing a 50 μL Taq PCR mix, 46 μL nucleasefree water, 2 μL Universal CRISPR oligonucleotide (100 μM), and 2 μL target-specific oligonucleotide (100 μM) in a thinwalled PCR tube on ice. 2. Mix gently, remove 3 μL of the mixture and keep on ice as a negative control. Run the annealing and extension reaction in a thermal cycler as detailed in Table 1. 3. Following the annealing and extension reaction, take a 3 μL sample from each tube and run each sample alongside the negative control on a 1.2% agarose gel to assess the formation of the dsDNA template (Fig. 1). The ssDNA and dsDNA templates visualized using the intercalating reagent Ethidium Bromide. dsDNA templates show no increase in size following the annealing and extension reaction, however the dsDNA bands are much brighter and tighter than ssDNA, indicating that the procedure was successful. 4. sgRNA can now be transcribed from the dsDNA template using the MEGAshortscriptTM T7 Transcription Kit (Invitrogen – ThermoFisher Scientific) or a similar kit.

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Table 1 CRISPR annealing and extension reaction Stage

Temperature (°C)

Time (s)

1

Denaturation

95

300

13

Denaturation Annealing Elongation

95 65 68

20 20 15

30

Denaturation Annealing Elongation

94 58 68

20 20 15

1

Final elongation

68

300

1

Hold

4

1

Fig. 1 ssDNA and dsDNA templates visualized using the intercalating reagent Ethidium Bromide

5. MEGAshortscriptTM T7 Transcription Kit: Add 2 μL T7 10× Reaction Buffer, 2 μL T7 ATP Solution (75 mM), 2 μL T7 CTP Solution (75 mM), 2 μL T7 GTP Solution (75 mM), 2 μL T7 UTP Solution (75 mM), 8 μL template dsDNA template, and 2 μL T7 Enzyme mix, into a single 1.5 mL RNase and DNase free tube. It is important to set up the reaction at room temperature and add the reagents in the order specified to prevent precipitation of the DNA by components in the transcription buffer. 6. Incubate the transcription reaction at 37 °C for at least 2 h. In our hands overnight (12–16 h) incubation leads to significantly improved yield. 7. To remove any residual DNA template, add 1 μL TURBO DNase I to each tube and gently mix by pipetting.

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Fig. 2 sgRNA constructs visualized by agarose gel electrophoresis

8. Incubate all tubes on the heat block for a further 15 min at 37 °C. 9. Use the SigmaSpinTM Sequencing Reaction Clean-Up Kit (Sigma-Aldrich) or an equivalent commercial micro spin column purification kit (see Note 7) to purify the RNA. Prespin the SigmaSpin columns for 2 min at 750 × g to remove excess buffer. Load the RNA mixture into the raised column. Centrifuge for 4 min at 750 × g to collect the purified RNA. The final elution volume is typically 20–30 μL. 10. Measure the concentration and purity of each sgRNA preparation on a small-volume UV spectrophotometer. The RNA concentration can range from 2000 ng/μL (typically expect around 1000 ng/μL from an overnight incubation); to proceed with this protocol, it is recommended that the RNA concentration be at least 300 ng/μL. 11. It is important to visualize the sgRNA by running 200–500 ng of the sgRNA preparation on a 1.5% agarose gel to ensure it is not degraded, a smear will indicate degradation. Due to secondary structure RNA often runs as two bands on non-denaturing gels (Fig. 2). 12. Store sgRNAs as single-use aliquots at -80 °C at a concentration of 300–500 ng/μL. 3.3 Injection of CRISPR/Cas9 Constructs into Xenopus Embryos

The third step in CRISPR/Cas9 gene disruption studies is generating the crispant animals. The protocol presented is an adaptation of Molecular Methods in Developmental Biology—Chapter 10: Microinjection into Xenopus Oocytes and Embryos [12]. 1. Prepare fertilized eggs for injection by removing the outer jelly coats in 2% L-Cysteine (non-hydrochloride solutions should be used when handling X. tropicalis embryos) solution (pH 7.8–8.0) with gentle rocking.

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2. When satisfactorily de-jellied, remove the Cysteine solution over five 30-s washes in MMR (0.05× MMR for X. tropicalis or 0.1× MMR for X. laevis). 3. Transfer batches of 100 embryos, into a petri dish containing an injection grid and 3% Ficoll solution. 4. Calibrate needles by droplet size, using nuclease-free water. Needles should be cut, and injection apparatus set up to deliver 4 nL injections over a 50 ms period. If 4 nL droplets are produced in less than 50 ms the needle is too big and will damage the eggs, this is especially important for X. tropicalis eggs which are smaller and less robust. The droplet volume is measured using a graticule or by drawing the liquid into a glass capillary by surface tension and measuring the length of the fluid column. Once the apparatus is set up satisfactorily, expel the water from the needle and draw in, or backfill, with the CRISPR injection mix. 5. CRISPR injection mix: Prepare 300–1500 pg sgRNA (see Note 8 for steps to improve efficiency), 2.6 ng Cas9 protein in a 1.5 mL DNase- and RNase-free tube, make up to 4 μL using nuclease-free water. Mix by pipetting up and down, centrifuge (1 min, RT, 1000 × g) and maintain on ice. 6. Using the manipulation settings on the picoinjector, gently lower the needle into the animal pole of a single cell embryo or the blastomere(s) of interest (see Note 5 for more details concerning cellular targets). 7. Following RNA injection, embryos should be transferred into fresh 3% Ficoll solution and maintained in agarose-lined Petri dishes at a species-specific temperature (14–18 °C X. laevis and 21–28 °C X. tropicalis) until NF stage 1027. 8. At late blastula stages, it is recommended that embryos are washed in MMR to remove the Ficoll solution and transferred into a fresh dish containing either 0.05× (X. tropicalis) or 0.1× MMR (X. laevis) solution. 9. Culture embryos/tadpoles in petri dishes until the desired stage, removing dead embryos daily. At free-feeding stages, move tadpoles to large dishes and incorporate 50% media changes alternate days (see Note 9 for further details) until tadpoles reach the stage of experimental interest. 3.4 Genotyping Embryos

The next step in CRISPR/Cas9 gene disruption studies is genotyping the crispant animals. The success of genome editing can be judged using the T7 endonuclease I assay, or by Sanger sequencing of amplicons containing the target site and tracking of indels by decomposition (TIDE or Synthego ICE).

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1. Collect whole embryo or partial (tail, toe, organ-specific) Xenopus tissue samples (see Note 10) in 1.5 mL DNase and RNase free-tubes and freeze the samples at -80 °C. 2. Thaw samples at room temperature and add 60 μL Lysis buffer, vortex the samples, and incubate for 2 h at 56 °C. 3. Using a heat block, incubate samples at 95 °C for 15 min to inactivate Proteinase K. 4. Centrifuge samples briefly (1000 × g for 1 min, RT) and proceed with PCR amplification. Alternatively, gDNA samples can be stored at -20 °C (see Note 11 for sample preparations requiring high-purity gDNA extracts). 5. Prepare a PCR master mix to amplify control and test samples. Into a single tube, add 12.5 μL Taq PCR mix, 8.5 μL nucleasefree water, 50–80 ng gDNA extract, 1 μL Target-FWD primer (10 μM), and 1 μL Target-REV primer (10 μM). It is recommended that an initial temperature gradient (54–64 °C) be performed, particularly in instances where primer pairs are located in non-coding regions. 6. Mix samples gently by pipetting up and down and maintain the mixture on ice before running the Genotyping PCR reaction detailed in Table 2. 7. Following the Genotyping PCR reaction, take a 3 μL sample from each tube and run each sample on a 1.2% agarose gel to assess the amplicon. 8. The efficiency of each sgRNA can be rapidly assessed by the T7 endonuclease I assay [35]. Mix 200 ng PCR product from step 3.4.(6.), 2 μL NEB Buffer 2, and nuclease-free water to 19 μL in a thin-walled PCR tube. Denature the DNA, at 95 °C for 5 min, then re-anneal by ramping down the temperature to

Table 2 PCR amplification of genomic target region Stage

a

Temperature (°C)

Time (s)

1

Denaturation

95

300

40

Denaturation Annealing Elongation

95 58a 72

30 30 60b

1

Final elongation

72

360

1

Hold

4

1

Annealing temperature is primer specific. bExtension times for products over 1000 bp are adjusted, the extension time increased 30 s for every additional 500 bp

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Table 3 Thermocycler conditions for T7 endonuclease I assay Stage

Temperature (°C)

Time (s)

1

Denaturation Ramp to Ramp to Hold

95 85 25 4

300 -2 °C/s -1 °C/s 1a

1

Incubation

37

900

1

Hold

4

1

a

Add T7 Endonuclease I after this step. NB Please note the denaturation ramp may vary between different thermocyclers

Fig. 3 Genotype analysis of F0 mosaic tadpoles. The target locus is amplified from genomic DNA preparations of injected (crispant) and uninjected (control) tadpoles. Amplicons are digested with T7 Endonuclease I, which reveals a second band unique to crispant tadpoles that correspond to the location of the CRISPR target site (A). Sanger sequencing is used to confirm the presence of indels in these crispant samples. Analysis of the Sanger sequencing trace files by Synthego ICE revealed the editing efficiency of the target domain is around 50% (B)

85 °C at -2 °C/s then to 25 °C at -1 °C/s (Table 3). Add 1 μL T7 endonuclease I (10 U) and incubate at 37 °C for 15 min. The T7 Endonuclease I assay cleaves DNA mismatches. The CRISPR/Cas9 process occurring at the same time as rapid cell division in the embryo usually results in high mosaicism of indels, and if the gene-editing process was successful, the re-annealed amplicons should form heteroduplexes which will be cleaved by the nuclease. 9. Digested amplicons are assessed on a 1.2% agarose gel. DNA fragments corresponding to cleavage at the CRISPR cut site should be visible (Fig. 3a). See Note 12 for troubleshooting. 10. Sequence gene-edited samples to confirm the presence of indels. If the CRISPR/Cas9 was successful, the sequence should become mixed around the Cas9 cleavage site due to mosaicism. The resulting chromatogram/ab1 trace files can be uploaded to freely available online platforms (for example, TIDE [36] or Synthego ICE [37]) to examine the predicted indel burden of samples. See Fig. 3b, for example, and Note 13 for troubleshooting and additional analyses of gene-edited samples.

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Fig. 4 Detailed structural differences are examined in Xenopus tadpoles using high-resolution Micro Computed Tomography. The imaged volumes show a whole tadpole with a 3D reconstruction of the cardiovascular system and gut visualized using TXM3DViewer (Carl Zeiss Microscopy) that can be exported as cross-section high-resolution TIFF image files

3.5 Phenotyping Mutant Embryos

The final step in CRISPR/Cas9 gene deletion studies is phenotyping the crispant animals. There are many strategies to explore the effects of knockouts, here we report MicroCT as a broad and extremely useful technique to investigate comprehensive or unexpected phenotypes (Fig. 4). 1. Gross morphological assessment can be performed under standard bright-field microscopy. For best results, image embryos/ tadpoles on agarose lined dishes. See Note 14 for considerations where an unexpected phenotype presents in the F0 crispant model. 2. Detailed structural differences can be examined using highresolution Micro Computed Tomography (MicroCT) in fixed specimens contrast stained in Phosphotungstic acid (PTA, Sigma-Aldrich) [38]. The recommended stage of analysis is NF42-50. Resolution below NF30 is poor and penetrance of the contrast reagent, using this methodology, above NF50 is highly variable. 3. Fix terminally anaesthetized (0.2% MS222) tadpoles in glass vials containing 5 mL MEMFA for 2 h and dehydrate in a series of Methanol washes: 25%, 50%, 75%, and 100%. Replace the 100% Methanol once more, and store embryos at -20 °C. 4. Rehydrate fixed samples in a series of methanol washes: 100%, 75%, 50%, and 25% and replace the final methanol concentration with 1% PTA for 48 h (NB it is essential that no salts are introduced after this step). 5. Remove the solution containing 1% PTA and wash all tadpoles in nuclease-free water. 6. Embed specimens in 0.7% agarose. Dissolve the agarose in nuclease-free water and maintain it at 60 °C on a heat block until use. Cool the agarose solution to 30 °C, immerse the tadpole in agarose and quickly transfer the tadpole into the

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imaging vessel. It is recommended that X. tropicalis tadpoles are embedded in a 20 μL pipette tip, and X. laevis tadpoles are embedded in a 200 μL pipette tip. After the agarose has solidified, seal the open ends with glue. 7. Run all samples in line with the recommended settings: The Zeiss Xradia Versa 520 (Carl Zeiss Microscopy) is set to operate at a voltage of 50 kv and a current of 75 mA. The 4× objective lens is used to provide an effective isotropic voxel size of 3.1 μM, with 1601 projections collected over 360° under an exposure time of 2.0 s per projection. 8. Reconstruct the tomograms to 16-bit grey-level images using the manufacturer’s software (Scout and Scan Reconstructor, Carl Zeiss Microscopy) which employs a filtered backprojection algorithm. 9. Visualise the imaged volumes using an image viewer capable of opening microscope image files (for example, TXM3DViewer (Carl Zeiss Microscopy)). MicroCT analysis of this nature provides a reference for crispant models and can guide downstream analysis.

4

Notes 1. It is possible to buy commercial sgRNAs as an alternative to generating the template in-house. 2. Identify the number of homologs in X. laevis, any gene duplication events (affecting both X. laevis and X. tropicalis) and where relevant, the number of isoforms of each gene. The integrated RNAseq data available in v9 of the Xenopus genome can guide understanding of the expression pattern of each gene of interest. Where genes are not well annotated or cannot be found in Xenopus, blast the H. sapiens protein sequence to search the Xenopus genome and consider looking at synteny of the surrounding genes (using, for example, Genomicus or PANTHER). 3. The overall success of CRISPR/Cas9 experiments depends on the experimental design and target identification. Historically, initial CRISPR/Cas9 gene-editing experimental designs targeted the first exon of a gene. Following the discovery that cryptic promoters in the first non-coding region hold the potential to rescue protein expression, this approach quickly moved to target early exons (exon 2 or exon 3) in the gene. Creating premature stop codons near the beginning of the transcript often results in nonsense-mediated decay of the transcript. Genotyping analysis revealed that many experiments resulted in a high proportion of in-frame mutations, which

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are often silent. The implementation of machine learning algorithms can minimize the proportion of in-frame mutations (for example, the inDelphi mESC model can predict frameshift frequency, precision, and indel locations). In recent years, research using CRISPR/Cas9 has focused on targeting an essential functional domain of the protein. Here the occurrence of both in-frame and out-of-frame mutations facilitates the study of protein dysregulation by interrupting protein function. Building on this idea, variant-directed crispant models of human disease are founded on the understanding that deleterious changes in the human sequence indicate a site within the protein that is not tolerant to change due to functional or structural consequences. Always check previously published oligonucleotide templates against the most recent predictive algorithms and blast sequences against the latest version of the Xenopus genome. 4. In cases where the exons of a gene are small and prove difficult to target, consider removing the coding region within a key functional domain by targeting CRISPR to the relevant splice donor or splice acceptor. Unless intending to disrupt the splice site, do not situate CRISPR/Cas9 constructs within 20 bp of the end of the exon. Similarly, unless intentional, avoid targeting regulatory elements. In cases where genomic regions are known to contain large, highly repetitive regions consider using a dual nicking approach and design two sgRNAs to use with Cas9 nickase. In addition, two sgRNAs can be injected simultaneously with Cas9 protein to remove a large region of the gene (>500 bp). Unless using the Cas9 nickase or deletion approach, it is not recommended that researchers co-inject multiple sgRNAs simultaneously within one embryo, as it complicates the downstream analysis of the line. Further, to circumvent limitations of the sgRNA position, different Cas enzymes possessing different PAM specificities can be considered. When gene-editing an allotetraploid X. laevis locus (or a duplicated gene), it is recommended that a conserved region of both genes is targeted using a single sgRNA. There are increasing reports of the unaccounted effects of CRISPR/Cas9 off-target consequences, making it important to implement effective experimental controls. In CRISPR/Cas9 experiments, the recommended controls for the delivery of constructs vary greatly, from non-coding or scrambled sgRNA templates to the use of positive control genes, including housekeeping genes or those with known phenotypes. In addition, it is possible to incorporate the use of multiple sgRNAs targeted to one region within the experimental design and demonstrate phenotype specificity or rescue the effects of gene knock-out experiments with the re-introduction of the WT allele.

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5. Taking advantage of the well-defined Xenopus fate map, geneediting can be restricted to specific blastomeres to target the desired organ systems of interest (for example, the kidney primordia). This technique is largely implemented to circumvent lethality in the F0 generation. Building on this, the most recent approach implementing this technique to generate a genetically altered line targets the four vegetal cells at 32-cell stage to generate a highly mosaic animal bearing edited germ cells. 6. Online tools do not always hold the most up to date version of the Xenopus genome assemblies. It is recommended that all primer sequences be blasted using the local blast tool located within Xenbase. Where possible, it is recommended that primers are located within coding regions or the first 20 bases of the intron, and the CRISPR/Cas9 cut site is centrally located within the amplicon. Here, it is also suggested that the ideal amplicon size should be approximately 500–800 bp. Larger amplicons (>1 kb) can prove problematic as increasingly genomic regions that are associated with the disease are found to occur in regions of the genome where there is a lot of secondary DNA structures, and smaller amplicons ( 3′ fashion that forms phosphodiester bonds. These principles were used by Sanger in a process known as a Sanger Dideoxy-sequencing [3]. In this process nucleotide analogs are added that obey the complementarity rules but have the deoxyribose 3′ OH removed, thus not allowing the addition of another nucleotide and serving as DNA polymerase terminators. The Sanger sequencing process involves terminators that are labeled with radioactivity. A set of four sequencing reactions are setup and the products are separated via slab gel electrophoresis, resulting into separated fragments at a resolution of one base, and the results are detected on a radiographic plate. DNA sequencing has evolved by replacing radioactivity with fluorescent terminators that are incorporated in a single reaction that undergoes steps of denaturation, annealing, and extension, creating products that are terminated by one of the four nucleotides (Fig. 1) [4]. Every terminator is labeled with a particular fluorescent dye with distinct excitation and emission properties. The characterization of the resulting fragments is automated, and it involves separation of the extension reaction in an electrophoretic matrix, which is housed within a special capillary. The capillary contains a tiny window that allows the exposure of the fluorescent extension products to laser excitation while the emission signals are recorded by an autodetector and are translated to peaks with the use of appropriate software (Figs. 2 and 3). The cycle sequencing technology via capillary electrophoresis is predominantly available from Applied Biosystems (Thermo Fisher Scientific). There are five systems currently available by the company as summarized in Table 1. This protocol focuses on the use of

Fig. 1 Fluorescent cycle sequencing overview

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Fig. 2 Sanger sequencing vs. cycle sequencing

Fig. 3 Capillary electrophoresis principle of operation

the latest instrument in capillary electrophoresis technology, the Seqstudio™ by Thermo Scientific [6]. It is a low throughput benchtop instrument featuring an all-in-one removable cartridge that contains the capillary array, polymer reservoir, and the anode buffer. This feature is very advantageous for a small to mid-size University laboratory as it helps avoiding the situation of capillary deterioration as a result of occasional usage. Finally, an important feature of Seqstudio™ is the flexibility of operation as can be operated though a PC and can be monitored remotely through cloud-based applications. Here, we present a protocol the Seqstudio™ is treated as a stand-alone instrument, and it is a part of a multistep process such as the sequencing of PCR products.

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Table 1 Capillary electrophoresis systems (Life Technologies) [5] Seqstudio genetic 3500 series genetic analyzers 3730 genetic analyzer analyser (3500, 3500 xL) (3730, 3730 xl) Number of capillaries

4

8, 24

48,96

Maximum number of 6 dyes

6

5

Capillary cartridge

Yes

No

No

Polymer type

POP-1

POP-6, POP-7, POP-4*

POP-6, POP-7

Minimum run time

30 min

30 min

20 min

850b

900b

2 × sample plates 96 or 384 – well

16 × sample plates 96 or 384 well

Maximum sequencing 800b read length Sample capacity

2

12 × 8-strip 1 × 96-well plate

Materials Ensure that the work takes place in a dedicated laboratory and that PPE is worn at all times in order to avoid contamination. Use deionized water for buffer dilution and PCR grade molecular biology water (12 Ω) for cycling reactions. When planning your work, primers should be ordered first as they may take longer time to arrive as they are customary synthesized.

2.1

Equipment

1. Microcentrifuge (see Note 1). 2. Horizontal Gel electrophoresis Tank and Powerpack (see Note 2). 3. Gel Documentation (see Note 3). 4. Thermal Cycler (see Note 4). 5. Nanodrop™ 2000. 6. Seqstudio™ (Applied Biosystems). 7. Seqstudio™ Cartridge v2 (Applied Biosystems). 8. Seqstudio™ Cathode Buffer (Applied Biosystems). 9. Intergrated Capillary protector (Applied Biosystems). 10. Reservoir Septa (Applied Biosystems). 11. PC for data storage and analysis (see Note 5). 12. Vacuum Centrifuge. 13. PCR hood.

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2.2 Kits, Reagents, and Consumables

199

1. Qiaquick PCR Purification Kit (Qiagen). 2. Big Dye™ Terminator v3.1 Cycle Sequencing kit (Applied Biosystems). 3. DyeEx 2.0 Spin kit (Qiagen). 4. Sequencing primers (as described in the methods section). 5. MicroAmp™ Reaction Tubes or MicroAmp™ 96-well plates. 6. Agarose Low EEO. 7. 10 X TBE Buffer. 8. DNA Ladder (see Note 6). 9. Gel Electrophoresis Loading Dye (see Note 7). 10. Gel Staining agent (see Note 8). 11. Hi-Fi™ Formamide (Applied Biosystems).

3

Methods Order all materials in advance and store them according to manufacturer’s instructions. DNA Cycle Sequencing should take place in a dedicated area to avoid carry over contamination, and procedures should be conducted in room temperature. Wear PPE to avoid contamination and to minimize degradation due to nuclease exposure.

3.1 PCR Product Purification and Clean up

Prior to cycle sequencing, PCR products are purified from the PCR Master mix and primer dimers with the use of Qiaquick PCR Purification Kit (Qiagen) [7]. In case that PCR results in nonspecific products, a special protocol that involves gel excision is required.

3.2 Specific PCR Product

1. To 1 volume of each PCR sample, add 5 volumes of PB buffer and mix well. 2. Apply the mixture to each spin column which is placed in a collection tube. 3. Spin the columns in a microcentrifuge at 13000 rpm at room temperature for 30–60 s. 4. Discard the flow through. Place the spin column in the collection tube and add 750 μL of PE Buffer to the spin column (see Note 9). 5. Spin at 13000 rpm for 30–60 s. Discard flowthrough, place back to collection tube, and repeat spin to dry the spin column. 6. Remove spin column and add it to a 1.5 mL microcentrifuge tube. Add 30–50 μL EB buffer or water. 7. Spin at 13000 rpm for 30–60 s to elute DNA.

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3.3 Excision Gel Protocol

1. Excise the gel band with a new scalpel and place in a microcentrifuge tube. Minimize the amount gel at a maximum of 400 mg. 2. Add 300 μL of buffer QG per 100 mg of gel for 1% agarose. For >2% agarose gels, add 600 μL QG per 100 mg of gel. 3. Incubate the microcentrifuge tube with the QC buffer and the gel slice at 50 °C for 10 min with periodical vortexing until melting. 4. Add 100 μL of isopropanol per 100 mg of gel slice if the fragment in question is ≤500 bp and 4Kb > (see Note 10). 5. Mix the resulting liquid well and place it to the spin column, centrifuge for 1 min at 13000 rpm and discard the flowthrough. If the liquid is more than 700 μL repeat step until all liquid flows through. 6. Follow steps 4–7 as described in Subheading 2.1.

3.4 PCR Product Quality and Quantity Assessment 3.4.1 Measurement of Concentration with Nanodrop™ 2000

Purified PCR products can be readily tested for concentration using the Nanodrop™ 2000 instrument (Thermo Fisher) [8], which is an accurate and versatile instrument that requires minimum amount of sample (1–2 μL) and lower detection limit of 2 ng/μL. Template concentration is critical for the downstream sequencing reactions. 1. Double click on the desktop NanoDrop™ 2000 software icon and select the “Nucleic Acids” icon. 2. Select “Add to report” so the measurements will be saved in the report. 3. Use appropriate buffer to establish a blank. An appropriate blank would be the solution that the PCR product is eluted or dissolved in. 4. Pipette 1–2 μL of the blank solution onto the bottom pedestal, lower the arm, and click the “Blank” button. Ensure that the arm is always down for all measurements. 5. Clean the bottom pedestal with a Kimwipe, enter the sample ID, and then click the “Measure” button.

3.4.2 Agarose Gel Electrophoresis

Prior to downstream sequencing applications, the PCR products should be checked with agarose gel electrophoresis to ensure: (a) the integrity of the template and (b) the absence of primer dimers that may interfere with downstream sequencing reactions. For gel electrophoresis, the materials used are listed in Subheading 2.1, and an established methodology is followed [9].

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3.5 Cycle Sequencing 3.5.1 Considerations for Sequencing Primer Selection and Design

201

In cases where the amplification product is less than 800 bp, one direction sequencing reaction may be sufficient to obtain the necessary data. However, to obtain maximum data, two reactions in forward and reverse fashions are advised. Forward and reverse sequencing is invaluable not only for improved coverage but also as a means of cross-referencing and validating sequencing results. Very often the primers used in either direction could be the original primers used for fragment amplification. Nevertheless, in cases where resequencing is required or the fragments are too large (> 1500), nested primers must be employed in order to maximize coverage. The criteria for primer selection and design are as follows [10, 11]: 1. Primers should be 20–30 bp in length. 2. Close to 50% GC content. 3. The primers should include a C or G at the 3′ of their sequence. 4. Thymidine at 3′ and 5′ and four or more repeated bases should be avoided. 5. Primers should not ideally form secondary structures or hybridize with each other forming hairpins and loops. This issue can be checked with the use of appropriate applications available from primer synthesis vendors. 6. Melting temperature range is 55–65 °C. 7. Primers should be checked for specificity to the target with the use of nBlast program and or annealing with the use of an alignment program such as MEGA7. 8. As cycle sequencing reaction is often unreadable for the first 30–40 bp, the primer should be designed upstream of the area of interest. Finally, there is the option of including tag sequences to the amplification primers. These tag sequences have to complements corresponding to universal primer sequences listed in Table 2 that are known to be reliable for DNA sequencing.

Table 2 Recommended universal primers for cycle sequencing [12] Primer name

Sequence (5′ to 3′)

M13–21

TGT AAA ACG ACG GCC AGT

M13–47

CGC CAG GGT TTT CCC AGT CAC GAC

M13-REV4

TCA CAC AGG AAA CAG CTA TGA C

T7

TAA TAC GAC TCA CTA TAG GG

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3.5.2 Cycle Sequencing Reaction

The cycle sequencing reaction is setup with the use of Big Dye™ Terminator v 3.1 Cycle Sequencing (Applied Biosystems) [13]. It is recommended that the reaction is setup in a dedicated PCR cabinet to avoid cross contamination. In addition, frequently calibrated pipettes dedicated for sequencing should be used for the setup cycle sequencing and other procedures. 1. Thaw the contents of the Big Dye™ Terminator v 3.1 kit and keep in ice. 2. Prepare 3.2 μM solutions for sequencing primer(s) and keep in ice. 3. Label 1.5 mL microcentrifuge tubes accordingly noting sample and primer used, and add the following components as described in Tables 3 and 4. The total volume of the reaction is 20 μL. Mix well, centrifuge and keep on ice. 4. There is an option of diluting the sequencing reaction. For that purpose, the Big Dye™ Terminator v 1.1 & v 3.1 5X Sequencing Buffer should employed as shown in the example in Table 5. Diluted reactions are not recommended to run without optimization as the resulting sequence may be compromised. The volume of Buffer is given by the formula: Buffer Volume = 0.5 * [(Total Reaction Volume)/2.5 – Reaction Mix].

Table 3 Components required per cycle sequencing reaction Component

Quantity

Example volume

Big dye terminator 3.1 ready reaction mix

8 μL

8 μL

Sequencing primer

3.2 pmoles

1 μL

Water (molecular biology grade)

Variable

9 μL

Template

Variable depending on concentration of template (see Table 6)

11 μL

Total

20 μL

Table 4 Recommended DNA quantities per reaction PCR product size

Quantity

100–200 bp

2–6 ng/μL

200–500 bp

6–20 ng/μL

500–1000 bp

10–40 ng/μL

1000–2000 bp

20–80 ng/μL

>2000 bp

40–100 ng/μL

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Table 5 Example of a diluted sequencing reaction

Component

Quantity

Example volume

Big dye™ terminator 3.1 ready reaction mix

4 μL

4 μL

Big dye™ terminator v 1.1 & v 3.1 5X sequencing buffer

2 μL

2 μL

Sequencing primer

3.2 pmoles

1 μL

Water (molecular biology grade)

Variable

11 μL

Template

Variable depending on concentration of template (see Table 2)

2 μL

Total

20 μL

Table 6 Cycle sequencing reaction parameters Parameter

Temperature

Time (sec)

Ramp rate

Incubation

96 °C

60

–

Denaturation Annealing Extension Hold

96 °C 50 °C 60 °C 4 °C

10 25 cycles 5 240 Until purification

1s

5. The reaction mixtures can be transferred now to PCR tubes or appropriate 96 plates; gently mix and proceed to cycle sequencing. 6. Place the tubes in a thermal cycler such as the Proflex PCR system and perform the cycle sequencing reaction following the conditions listed in Table 6. 7. The samples can now undergo purification or stored in containers that will minimize exposure to light at -20 oC. 3.6 Cycle Sequencing Product Purification

After completion of the cycle sequencing, the excess dye terminators are required to be removed. For this purpose, we employ the DyeEx 2.0 Spin kit (Qiagen) [14], which uses gel filtration technology. 1. The column is gently vortexed in order to resuspend the resin. 2. The cap on the column must be slightly loosened (about one quarter turn) so vacuum within the tube is avoided.

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3. The bottom closure of the should be broken and the spin column should be placed in the provided 2 mL tube. 4. Centrifuge the spin column for 3 min at 750 × g. 5. The spin column is transferred now to a microcentrifuge tube and apply with care 20 μL of the cycle sequencing reaction. The application should be done in the middle of the spin column and without disrupting the gel filtrate matrix. 6. Remove the spin column from the microcentrifuge tube. The eluate contains the purified cycle sequencing product. The sample is dried in a vacuum centrifuge. 3.7 Sample Preparation

1. Thaw a fresh aliquot of Hi-Fi™ Formamide. Make sure that it has not been through more than two freeze-thaw cycles. 2. Resuspend the samples in 10–20 μL Hi-Fi™ Formamide. Do not resuspend in water as the sample stability will be decreased. 3. Transfer the samples in MicroAmp™ Optical 96-Well Reaction Plates or MicroAmp™ Reaction Tubes. The tubes are provided in strips of 8 and should be kept in place by the MicroAmpTM 96-well tray and tray retainer. 3. Cover the plates or tubes with the appropriate septa by aligning the holes of the septa with the wells or the tubes. Ensure that the septa are in position by appropriate pressing. 4. Centrifuge briefly the plates or reaction tube assemblies to ensure that the contents are at the bottoms of their tubes. 5. Run the samples as soon as possible after resuspension and setup.

3.8 Running Sequencing on the Capillary Electrophoresis System (Seqstudio™)

The instrument electrophoresis run can be setup from a connected PC through a mobile device or from the instrument touch screen [15]. Here, we describe the setup from the touch screen. Prior to starting the run, it is important to check the storage history of the cartridge (see Note 11). 1. From the start up menu, touch the Eject button and carefully open the instrument door according to the screen prompt. 2. Open the lid by pressing the release button on the autosampler. 3. Ensure that the Cathode Buffer Container is full above the fill line. Replace it accordingly (see Notes). 4. Place the plate or the tube assembly firmly in the autosampler and close the lid until it clicks shut. 5. Select “Retract Plate” from the “Start Menu” and close the instrument door. 6. To run the system go to the home screen and setup run and follow the instructions in the previous section or select.

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Table 7 Run modules, read lengths, and appropriate run times Run module

Continuous read Length (CLR)

Approximate run time

ShortSeq

≥350 bp

30 min

MediumSeq

>500 bp

45 min

LongSeq

>800 bp

120 min

7. In the home screen touch “Setup run.” 8. Next, touch “Create New Setup.” 9. At the top right of the Plate Properties, select the tab Properties. In the Applications box, select Sequencing. 10. Save location by selecting Instrument. This will save your setup in case you need to repeat the experiment. 11. At the “Plate Properties” menu select “Plate.” The wells are organized in 4 well injection groups. The default loading is A1-D1, E1-H1, A2-H2, etc. 12. From the “Plate Menu” Select “Edit.” Select the appropriate Run module based on the information displayed in Table 7. 13. To name the samples in the wells, select “Sample name” from the “Edit” menu. 14. Press “Done” and the “Plate properties” will appear. To run the samples, choose “Run.” You may need to select “Save” if you are planning to run the samples later. 15. During the run the status dials for every injection appear as colored circles. If the color is green, all quality control checks have passed. If the color is yellow, there is at least 1 warning quality alert. If the color is red, there is at least 1 failing quality alert. 16. You may need to stop the run if there are a failing quality alerts. To do that choose “Actions”.and from that menu “Cancel Remaining Injections.” 17. When a run is completed, a “Run Complete” message will appear on the screen. To view the results, select the “Results” box. To get a quality report, touch “List View.” The report is colored coded as described previously. 18. At the end of the run, touch the eject button on the screen, select “Eject” plate and after the instrument prompt opens. 19. To retrieve the results, attach an “instrument dedicated USB.” From the Home screen, select Setttings > Run History. Touch Export and select “Storage Location.” The files down loaded should be in ab1. Format.

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3.9 DNA Sequence Viewing and Analysis

Sequence can be viewed on the SeqStudio instrument. However, we find that the viewing and manipulation of sequences is more convenient on a PC with the aid of freeware. For routine use, the most cost efficient approach is to use the light version of Chromas™ (Technelysium Pty LTd) that allows visualization of the sequence and export to FASTA format which is useful when further bioinformatics processing is necessary. Access to sequence traces with Chromas™ is very simple as the user simply have to down load and install the program in the PC. Simply clicking on AB1 files will open the traces in Chromas. To analyze and process the sequence, take the following steps: 1. Without scrolling inspect the first 50 bp. You should see peaks of large intensity that are not distinctly separated. This is typical as primer binding affects the reading. Click on the sequence letters in the region of the first 50 bp. At the top right of your screen a quality value will appear. An acceptable quality value should be Q > 20. The majority of the first 40–50 bp usually are below 20. Based on this the region should be excluded. 2. Scroll the sequence the to the left. In a good quality sequence (Fig. 4b) the nucleotide peaks are very well separated from each other and the Q > 20 for all the nucleotides. On the contrary, a problematic sequence appears with very high noise or a large number of nucleotides with Q < 20 (see Troubleshooting). In that case, the sequence is rejected from further processing. 3. If you identify spurious nucleotides (i.e., 5 in a sequence of 500 bp) in the middle region with Q < 20, this may be due to heterozygosity in particular loci (see Subheading 3.9 Troubleshooting). 4. While scrolling to still to the left there is a point where the signal drops. This is the end of the sequence (see Fig. 4d). 5. When you are satisfied with the quality of sequence, select from the “Edit” menu the “Trim Low-quality” function and then “Export” the sequence as a “FASTA” file for further processing.

3.10 Troubleshooting

In this section we list some examples of problematic cycle sequencing of PCR products with the Seqstudio system. Troubleshooting advice is listed below every listed example.

3.10.1 Unincorporated Dye Blob

A common artifact that appears as between 70 bp and 120 bp is a large peak that obscures the real sequence (Fig. 5). This can be attributed to unincorporated Dye Blob and usually does not interfere with the reading of sequence that appears underneath. This can be solved with the addition of more DNA template or less BigDye in the reaction.

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Fig. 4 Typical images of a sequence as shown in Chromas™ lite. (a) The beginning of the sequence is affected by binding primer. (b) The middle part of a good quality sequence exhibits high Q values (Q > 20), and the peaks are well separated from each other. (c) The middle part of a poor quality sequence where there are stretches of multiple nucleotides (yellow boxes) with low Q values (Q < 20). (d) The end of the sequence where there is no coherent laser signal

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Fig. 5 Example of unincorporated dye blob

Fig. 6 Example of a poor start followed by a weak signal 3.10.2 Poor Start Followed by a Weak Signal

When there is a poor start followed by a weak signal, this may attributed either to primer binding to itself or there is other primer present (Fig. 6). This can be addressed either by redesigning a new sequencing primer or by checking the efficiency of PCR purification via gel electrophoresis.

3.10.3 Peaks

Overlapping

In the case of overlapping peaks in sequencing data, the sequence might exhibit multiple priming sites or the sequencing primer might not be sufficiently purified by the manufacturers and the impure mixture may be giving shadowing sequences (Fig. 7). In both cases, the use of newly synthesized primers is advised. There is a possibility that there is heterozygosity in the form of single nucleotide polymorphisms (SNPs) in which case cross-referencing with the sequence of the complementary strand is strongly advised for confirmation.

3.10.4

Failed Sequence

When the run fails to produce sequencing data, the probable causes might be the suitability of the sequencing primer (Fig. 8), the freshness of the primers stock, the quantity and quality of DNA, and the presence inhibitors within template DNA. It is advisable to repeat reaction first with a fresh aliquot of sequencing prime, measure the concentration of template DNA, and ensure that a

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Fig. 7 Example of overlapping peaks

Fig. 8 Example of a failed sequence

good quality PCR band is evident during electrophoresis. If the problem persists, a second round of PCR purification maybe required. Lastly, the synthesis of new sequencing primers may be required. 3.10.5 Secondary Structure

4

In some cases, sequence may start well but the signal may weaken rapidly (Fig. 9). This is due to repeat regions such as GT or CT repeats that may cause the sequencing signal to be depleted due to formation of DNA loops or secondary structures. It is advisable to add 1 μL of DMSO in the reaction or design nested sequencing primers close to the secondary structure.

Notes 1. A compact microcentrifuge 14,000 rpm is recommended.

with

maximum

speed

at

2. Electrophoresis tanks from recognized manufacturers with appropriate safety features are recommended. Powerpacks from recognized manufacturers that are compact with capabilities for 300 V, 400 Ma, and 60 W are sufficient.

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Fig. 9 Example of secondary structure issues

3. Entry level gel documentation systems that allow for the use of safe gel staining dyes are sufficient for this protocol. 4. It is recommended to use an Applied Biosystems™ thermal cyclers as Big Dye™ chemistry kits are optimized with the use of these instruments. Alternatively, cycle sequencing reaction optimization might be necessary. 5. The PC used for data visualization and storage in this protocol should be run on Windows™ 8 or 10 operation systems. 6. A wide range ladder should be used (100–10,000 bp) as the PCR products may vary in size. 7. Premade loading dye (6X Loading Dye) in order to ensure consistency during loading is recommended. 8. For staining of DNA in agarose we prefer to use dyes that are safer than ethidium bromide such as SYBRSafe™ and Noveljuice™. 9. Before the use of PE Buffer, add the appropriate (as recommended on the bottle) amount of ethanol (96–100%). 10. Fragments between 500b and 4 Kb are not affected by the addition of isopropanol. 11. The cartridge can be stored up to 4 months on or off the instrument. If the cartridge is stored off the instrument, it should be kept at 2–8 °C and in an up-right position with an integrated capillary protector and an optical cover installed. References 1. Saiki RK, Scharf S, Faloona FA, Mullis KB, Horn CT, Erlich HA, Amheim N (1985) Enzymatic amplification of β-globin genomic sequences and restriction site analysis for diagnosis of sickle cell anemia. Science 230:1350– 1354

2. Bevan SI, Rapley R, Walker MR (1992) Sequencing of PCR amplified DNA. PCR Methods Appl 1:222–228

Sanger Sequencing 3. Sanger F, Nicklen S, Coulsen AR (1977) DNA sequencing with chain terminating inhibitors. Proc Natl Acad Sci 74:5463–5467 4. DNA Sequencing by Capillary Electrophoresis 2nd Edition (2009) Applied biosystems, Foster City 5. Instruments for Sanger Sequencing and Fragment Analysis by Capillary Electrophoresis. Thermo Fisher Scientific. Available on: https://www.thermofisher.com/uk/en/ home/life-science/sequencing/sangersequencing/sanger-sequencing-technologyaccessories.html 6. Seqstudio Analyser Application Guide (2019). Applied biosystems 7. Qiaquick PCR & Gel Cleanup Kit Handbook. Qiagen. Available on: https://www.qiagen. com/zh-us/products/discovery-and-transla tional-research/dna-rna-purification/dna-puri fication/dna-clean-up/qiaquick-pcr-purifica tion-kit?catno=28506 8. Nanodrop 2000/2000c Spectophotometer User Manual (2009) Thermo scientific 9. Serwer P (1983) Agarose gels: properties and use for electrophoresis. Electrophoresis 4:375– 382

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10. De Bellis G, Manoni M, Pergolizzi R, Vezzoni P, Luzzana M (1990) Primer design in fluorescent DNA sequencing. Nucleic Acids Res 18(16):4951–4952. https://doi.org/10. 1093/nar/18.16.4951. PMID: 2395669; PMCID: PMC332014 11. Rychlik W (1993) Selection of primers for polymerase chain reaction. Methods Mol Biol 15:31–40. https://doi.org/10.1385/089603-244-2:31 12. Standard Primers (2021) MRC PPU DNA sequencing services. Available on: https:// www.dnaseq.co.uk/resources/primers/stan dard-primers 13. Big Dye Terminator v3.1 Cycle Sequencing kit. Pub No. MAN0015666 (2016) Applied Biosystems 14. DyeEx 2.0 Spin Kit Handbook (2002) Qiagen. Available on: https://www.qiagen.com/cn/ resources/download.aspx?id=c1992325-e41 f-4a78-8b5a-248cc91d2c2e&lang=en 15. Seqstudio™ Genetic Analyser Instrument and Software User Guide (2019) Applied Biosystems

Chapter 16 Nanopore Sequencing for Mixed Samples Angela H. Beckett and Samuel C. Robson Abstract Long read Nanopore sequencing can be utilised to determine the quality and accuracy of genetically engineered changes in animals, which often produce heterogenous samples. The protocol presented in this chapter can be used for a range of both low and high throughput sequencing applications. DNA must be repaired, barcoded and ligated to sequencing adapters prior to sequencing. Quality of sequencing data produced is dependent on stringent adherence to the protocol. However, nanopore sequencing is a fast moving field, therefore it is worth considering using the most up to date chemistry available. Key words Next-generation sequencing, Third-generation nanopore sequencing, Library preparation, Oxford Nanopore Technologies

1

Introduction Genetically engineered DNA modifications in microorganisms and animals must be confirmed and validated using sequencing technologies [1]. After the modified region is amplified by PCR, the amplicon is subcloned into a plasmid, transformed into bacteria and resultant colonies are sequenced [2]. Sanger sequencing has typically been used for detecting and testing efficiency of sequencing clones [3, 4]. This first-generation sequencing technique utilises a chain termination method. The target sample is divided into 4 PCR reaction pools; each pool contains deoxynucleoside triphosphates (dNTPs: A, T, G and C), polymerase, fluorescently tagged primer and one type of dideoxy-dNTP (ddNTP) per pool (A, T, G or C). ddNTPs are lacking a 3′ hydroxyl group which halts the action of polymerase during DNA extension, terminating the reaction. As ddNTP concentration is so low, each time extension occurs, differing amounts of dNTPs are added before a ddNTP is incorporated; thus, many-sized fragments are generated. After PCR, the product DNA from each PCR pool is loaded into individual wells of a denaturing urea PAGE gel. The size separation of the fragments is used to determine the reverse complement of the sequence. For

Garry Scarlett (ed.), DNA Manipulation and Analysis, Methods in Molecular Biology, vol. 2633, https://doi.org/10.1007/978-1-0716-3004-4_16, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2023

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example, if the smallest fragment on the gel is from Pool 1(A), and the second smallest fragment is from Pool 2 (C), the first nucleotide in the reverse complement of the sequence is G, the second is T and so on. In recent years, this process has been automated through the use of capillary electrophoresis and base-specific fluorescent tags. Second- and third-generation sequencing technologies have since been developed, such as short- and long-read high-throughput sequencing methods developed by Illumina and Oxford Nanopore Technologies (ONT), respectively [5]. The increased throughput, speed and quality of these technologies have led them to become widely adopted. Illumina utilises a bridge amplification technique; a sequencing adapter is bound to the end of DNA fragments, which are complementary to a lawn of oligos bound to the glass surface of the Flow Cell. Polymerase creates a complement of the hybridised strands, after which the double strand is denatured and the original template is washed away. The remaining strand folds over and binds to a second type of oligo bound to the Flow Cell surface forming a bridge, where it is replicated by polymerase and denatured into single strands (Bridge Amplification). The replication is repeated through multiple cycles, creating clusters of identical DNA oligos. Fluorescently tagged nucleotides are added one by one, with the signal released after each addition indicating which base has been incorporated. This occurs across all fragments on the Flow Cell simultaneously. Emitted fluorescence is detected, and the sequence for forward strands is calculated, after which the reverse strands are transcribed and the process is repeated. This massively parallel system greatly sped up sequencing efforts. However, this method is limited as it only generates short sequencing reads, thus resulting in potential gaps in the aligned sequence or difficulty resolving highly repetitive regions. This limitation has been addressed in third-generation long read sequencing, including nanopore-based sequencing from Oxford nanopore Technology (ONT) and Single Molecule Real-Time (SMRT) approaches from Pacific Biosciences (PacBio), capable of sequencing long fragments of DNA. In this chapter, the methodology for library preparation and sequencing using nanopore sequencing is described. ONT have developed a small sequencing machine (MinION), roughly the size of a standard stapler, which is portable and inexpensive (90%. 6. Tris-EDTA Buffer Solution pH 8.0. 7. NEBNext Ultra II End repair/dA-tailing Module (New England Biolabs - NEB). 8. NEBNext Quick Ligation Module (NEB). 9. Blunt/TA Ligase Master Mix (NEB). 10. Native Barcode Expansion Kit (ONT) (see Note 1). 11. Ligation Sequencing Kit 109 (LSK-109) (ONT). 12. Flow Cell Priming Kit (included with LSK kit) (ONT). 13. Flow Cell Wash Kit (ONT).

2.3

Equipment

1. Qubit Fluorometer. 2. PCR Hood. 3. PCR Thermal Cycler. 4. Mini Plate Spinner Centrifuge. 5. MicroAmp™ Cap Installing Tool.

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6. Microcentrifuge. 7. Vortexer. 8. Flow Cell MIN 109D (ONT). 9. 96-well cooling block. 10. Centrifuge tube Mini-cooler. 11. ONT Sequencer (see Note 2). 12. Magnetic Rack (see Note 3). 13. Timer. 14. P2, P10, P20/P100, P200, P1000 Single Channel Pipette. 15. 8x P10 Multichannel Pipette. 16. Repeat Pipettor (see Note 4).

3

Method Here we assume the appropriate region of the genomic DNA from the engineered animal has been PCR amplified, and the resulting amplicons will act as the starting material. Where possible, all work should be performed in a PCR hood, which limits contamination occurrence. All thawed reagents should be stored on cool blocks or ice unless otherwise stated.

3.1

Setting Up

1. Include a negative control, such as nuclease-free water (NFW), and a known positive control per sequencing library (i.e., per Flow Cell). If positive and negative controls have been added to the plasmid PCR, carry them through to library preparation along with the samples. 2. Create an experiment plan which includes sample ID and sample order. Up to 96 samples, including 2 controls, can be added to a single Flow Cell, depending on coverage needed per sample. 3. In a clean PCR hood/Laminar Flow, sub-aliquot NFW and other NEB reagents into single use volumes (see Note 5). 4. Thoroughly clean a PCR hood and pipettes with DNAse ZAP cleaning spray. 5. If possible, wear a different lab coat for library prep (post-PCR amplification) than worn for the PCR preparation (pre-PCR amplification) (see Note 6).

3.2 Post-PCR Quality Control

1. Quantify DNA from all samples, the positive control and the NFW negative control using the Qubit Fluorometer (ng/μL). 2. Label the required number of Qubit, 0.5 mL tubes to test the samples, including two additional tubes for the calibration standards (see Note 7).

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3. Prepare respective calibration standards by adding 190 μL of Qubit working solution and 10 μL of the respective standard (Standard 1 and Standard 2). 4. Add 199 μL of Qubit working solution and 1 μL of sample (see Note 8). 5. Mix contents by inverting three times, and perform a swift wrist flick to pool the contents at the bottom of the tube (see Note 9). 6. Allow tubes to incubate at room temperature for 2 min. 7. On the Home screen of the Qubit Fluorometer, select “1xdsDNA High Sensitivity,” followed by “Read Standards” (see Note 10). 8. Insert the tube containing Standard 1 into the sample chamber, close the lid and “Read Standard.” When the reading is complete (~3 s), remove Standard 1 (see Note 11). 9. Repeat process with Standard 2 (see Note 12). 10. Select “Run Samples” to proceed to sample quantification. 11. On the assay screen, select the sample volume (1 μL) and units required (ng/μL). 12. Insert a sample tube into the sample chamber, close the lid and select “Read Sample.” When the reading is complete (~3 s), remove the sample tube. 13. Repeat until all samples have been read. Record sample concentrations (see Note 13). 14. Check purity of DNA on a micro-spectrophotometer, and run blank before testing samples. If 230/260 and 260/280 are between 1.8 and 2.0, then continue with the experiment. If samples are not within range, perform appropriate clean-up before proceeding (see Note 14). 3.3 Sample Dilution, End Repair and dATailing

1. Thaw Mastermix 1 (MM1) reagents (Table 1) on ice. 2. For each sample, add 500 ng of DNA (≤24 samples) or 150 ng of DNA (>24 samples) and make up to 15 μL using NFW. Put each sample in an individual well on a 96-well plate. Keep 96-well plate on cool block throughout the process (see Note 15). 3. Prepare MM1 (Table 1): In a 1.5 mL DNA Lo-bind tube, combine Ultra II End Prep Reaction Buffer and Ultra II End Prep Enzyme Mix (see Note 16). 4. In clean wells, on the same 96-well plate used for sample dilution (if space, otherwise on a new 96-well plate), dispense MM1 into 3 μL aliquots (see Note 17).

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Table 1 Mastermix 1 reagent volume per sample Component

x1 sample volume (μL)

130–400 ng sample DNA

12

Ultra II End Prep Reaction Buffer

1.75

Ultra II End Prep Enzyme Mix

0.75

5. Using an 8x P10 Multichannel pipette (see Note 18), transfer 12 μL of diluted sample (containing either 400 ng or 130 ng of sample DNA) from the previous step to the corresponding well-containing MM1 (Fig. 3). 6. Centrifuge plate to collect contents at bottom of the well (see Note 19). 7. Incubate the reaction in a thermal cycler at 22 °C for 15 min, followed by 65 °C for 15 min (see Note 20). 8. When incubation is complete, cool on ice or on a cool block for 1 min. 3.4

Barcode Ligation

1. Meanwhile, prepare Mastermix 2 (MM2): In a 1.5 mL DNA Lo-bind tube, mix Blunt TA/Ligase Master Mix and NFW (Table 2) (see Note 16). 2. In clean wells on the same plate (if space, otherwise on a new 96-well plate), dispense MM2 into 8 μL aliquots (see Note 17). 3. Add 1.25 μL of barcode per sample, ensuring that each sample has a unique barcode and sample/barcode order has been recorded (see Note 21). 4. Add 0.75 μL of the previous reaction mixture (MM1 after incubation on thermal cycler) to each corresponding well, seal plate with Flat 8-cap strips using the capping tool and centrifuge plate (see Note 22). 5. Incubate the reaction in a thermal cycler as follows at 22 °C for 20 min, 65 °C for 10 min and 4 °C for 1 (see Note 23). 6. At the start of the incubation time, remove AMPure (SPRI) beads from the fridge and leave at room temperature, remove Short Fragment Buffer (SFB) from either the LSK kit or SFB Expansion Pack and thaw on ice/in a cool block. 7. After incubation, cool the 96-well plate containing MM2 on ice/in a cool block for 1 min.

3.5 Up

Barcode Clean-

1. In a new 1.5 mL DNA Lo-bind Eppendorf tube, pool all 10 μL barcoding reactions together (per library, e.g., 960 μL for 96-barcoded samples).

NGS

Diluted sample

MM1

MM2

01

09

01

09

01

09

02

0

02

10

02

10

03

11

03

11

03

11

04

12

04

12

04

12

05

13

05

13

05

13

06

14

06

14

06

14

07

15

07

15

07

15

08

16

08

16

08

16

12.0 µl

221

0.75 µl

Fig. 3 Example 96-well plate layout for 16 samples. As indicated by arrows, 12 μL of diluted sample DNA (containing 400 ng or 130 ng) with a total volume of 15 μl from samples 01–08 can be transferred to Mastermix 1 (diluted sample from Column 1: A1-H1 transported to Column 3: A3-H3) simultaneously using an 8x P10 Multichannel pipette. After Mastermix 1 has finished incubation and Mastermix 2 has been added to the plate, 0.75 μl of Mastermix 1 can be transferred by multichannel to Mastermix 2 (Column 3: A3-H3 transported to Column 5: A5:H5). (Figure created using BioRender.com)

Table 2 Mastermix 2 reagent volumes. ×24-×96 sample volumes have overage included

Component

×1 sample ×24 sample ×48 sample ×72 sample ×96 sample volume (μL) volume (μL) volume (μL) volume (μL) volume (μL)

Previous Reaction Mixture

0.75

NBXX Barcode

1.25

Blunt/TA Ligase Mastermix

5

135

265

395

525

Nuclease Free Water

3

81

159

237

315

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Fig. 4 Demonstration of pipetting technique whilst using magnetic rack. Magnetic rack is held at an angle whilst pipetting to better visualise the beads and improve pipetting accuracy

2. Mix beads thoroughly by vortex and mix-pipetting, until homogenous. Add 4 μL of AMPure beads per sample, i.e., 96 μL AMPure beads for an × 24 pooled sample reaction or 384 μL for 960 μL sample reaction (see Note 24). 3. Pulse centrifuge to collect tube contents at the bottom of the tube (see Note 25). 4. Incubate for 5 min at room temperature. 5. Whilst the library is incubating, prepare ethanol ahead of time. Dilute absolute ethanol to 70% by adding 700 μL ethanol to 300 μL of NFW (see Note 26). 6. Place library tube(s) onto a magnetic rack and incubate for 2 min (see Note 27). 7. Holding the magnetic rack at an angle, carefully remove and discard the supernatant, being careful not to touch the bead pellet (Fig. 4) (see Note 28). 8. Add 250 μL short fragment buffer (for x24-plex pool), 500 μL SFB (x48-plex pool), 1000 μL SFB (x96-plex pool) and re-suspend beads completely by pipette mixing. 9. Pulse centrifuge to collect tube contents at the bottom of the tube (see Note 29). 10. Place sample tube(s) onto magnetic rack and incubate for a minimum of 2 min.

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11. Carefully remove supernatant and discard (see Note 30). 12. Repeat SFB wash, pellet and discard. 13. Pulse centrifuge and remove any residual SFB using a P10 pipette (see Note 29). 14. Leave the tube in the magnetic rack, and add 200 μL of roomtemperature 70% ethanol to bathe the pellet. Do not resuspend or disturb the pellet. 15. Carefully remove and discard ethanol, being careful not to touch the bead pellet. 16. Pulse centrifuge to collect all liquid at the bottom of the tube, and carefully remove as much residual ethanol as possible using a P10 pipette. 17. With the tube lid open, incubate for 1 min or until the pellet loses its shine (see Note 29). 18. Resuspend the pellet in 30 μL NFW/Tris-EDTA buffer pH 8.0/Elution Buffer, mix gently by flicking/pipetting and incubate for 2 min (see Note 30). 19. Place on a magnetic rack and incubate for 2 min. 20. Transfer the library to a clean 1.5 mL Eppendorf tube ensuring no beads are transferred into this tube (see Note 31). 3.6 Sequencing Adapter Ligation and Clean-Up

1. Quantify 1 μL of barcoded amplicon pool using the Qubit Fluorometer, as described in Subheading 3.2, step 2 (see Note 32). 2. Set up the AMII adapter ligation reaction (Table 3). 3. Incubate at room temperature for 20 min. 4. Meanwhile, remove the required number of Flow Cells from fridge and allow to come to room temperature. 5. Mix AMPure beads thoroughly by vortex and mix-pipetting, until homogenous. 6. After the 20 min adapter incubation, add 50 μL (1:1) of AMPure beads to the library tube(s) and mix gently by inverting/pipetting. Table 3 Adapter ligation mix Component

Volume (μL)

Library (pooled & barcoded samples)

29

NEBNex 5X Quick Ligation Reaction Buffer

10

AMII Adapter Mix (LSK109 kit)

5

NEB Quick T4 DNA Ligase

5

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7. Incubate for 5 min at room temperature. 8. Place library tube(s) onto magnetic rack and incubate for 2 min. 9. Carefully remove and discard the supernatant, being careful not to touch the bead pellet. 10. Add 250 μL SFB and re-suspend beads completely by pipetting. 11. Pulse centrifuge and place library tube(s) onto magnetic rack and incubate for 2 min. 12. Discard supernatant. 13. Repeat SFB wash steps. 14. Pulse centrifuge and remove any residual SFB. 15. Add 15 μL Elution buffer (LSK) and re-suspend beads by pipette mixing. 16. Incubate at room temperature for 2 min. 17. Place on magnetic rack and incubate for 2 min. 18. Transfer final library to a new 1.5 mL DNA Lo-bind Eppendorf tube (see Note 33). 3.7 Sequencing Preparation

1. Quantify 1 μL of barcoded amplicon pool using the Qubit Fluorometer as described in Subheading 3.2, step 2 (see Note 34). 2. Thaw sequencing reagents at room temperature (Table 4) before placing on ice; a fresh tube of Flush Buffer is required per Flow Cell. 3. Open the ONT MinKnow Software. 4. Insert configuration cell into the instrument by sliding the cell under the clip and pushing down gently. Run the hardware check from the “Start” menu. Check system messages to see if hardware check was successful (see Note 35). Remove configuration cell.

Table 4 Sequencing reagents required to prepare library for loading onto flow cell Kit

Reagent

Ligation Sequencing Kit

Sequencing Buffer

Ligation Sequencing Kit

Loading Beads

Flow Cell Priming Kit

Flush Buffer (FB)

Flow Cell Priming Kit

Flush Tether (FLT)

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5. Load Flow Cell into the instrument by sliding it under the clip and pushing down gently. 6. If using Flow Cell containing storage buffer (first use Flow Cells and Flow Cells which have been stored after the second use), go to the “Start” menu and run a “Flow Cell Check.” A Flow Cell check takes ~20 min, so continue with the next step while waiting for it to complete. The number of pores available will display on the “sequencing overview” screen. Any value over 800 pores passes the ONT warranty threshold. If the number of available pores is below 800 on a new Flow Cell, and it is less than 3 months old; set it aside, inform ONT and request a replacement (see Note 36). 7. Meanwhile, add 30 μL FLT to the FB tube and mix well by vortex to create Flush buffer-mix. 8. Once the Flow Cell check is complete, open the priming port and take a P1000 pipette with the volume to 800 μL. Holding the pipette completely upright, place the empty pipette tip into the priming port and remove any air from the inlet port by turning the volume dial anti-clockwise (Fig. 5). Remove ~30 μL of the buffer from the Flow Cell (see note 37). 9. Load 800 μL of Flush buffer-mix into the Flow Cell via the priming port (Fig. 5), dispense slowly and smoothly to minimise shear forces on the library and avoid the introduction of any air bubbles (Fig. 5) (see note 38). 10. Wait for 5 min, in the meantime move to the next step.

1. Slide open the priming port cover by 90˚ to reveal the priming port

2. Place tip vertically into priming port and dial anticlockwise to remove air bubble

3. Load 800µl of Flush Buffer-mix slowly through the priming port

4. After 5 minutes, open the SpotON port

5. Load 200µl of Flush Buffer-mix slowly through the priming port

6. Load the library dropwise into the SpotON port

Fig. 5 Priming and loading a Flow Cell. Flow Cell is primed with flush buffer-mix prior to the library being loaded via the SpotON port. (Image adapted from © 2022 Oxford Nanopore Technologies plc)

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Table 5 Sequencing reagent quantities required to prepare library for loading onto flow cell

3.7.1 Library Preparation and Loading

Component

Volume (μL)

Sequencing Buffer

37.5

Loading Beads

25.5

Library

12

1. Dilute library in NFW so that 12 μL contains 40 ng of DNA (3.3 ng/μL) for x24–47 sample pools or 50 ng of DNA for ≥48 sample pools (4.2 ng/μL). If concentration of DNA is less than 3.3 ng/μL, do not dilute. If concentration of DNA is >15 ng/μL, perform a 1:1 dilution (see Note 39). 2. In a new tube prepare the library for sequencing (Table 5), ensuring that loading beads are mixed thoroughly by vortex and mix pipetting until homogeneous (see Note 40). 3. After the 5-min incubation of Flush buffer-mix on the Flowcell has elapsed, gently lift the SpotON cover to reveal the SpotON port (Fig. 5). 4. With the SpotON port open, load another 200 μL of Flush buffer-mix into the Flow Cell via the priming port. Dispense slowly and smoothly, leaving a small amount of liquid remaining in the pipette tip to avoid the introduction of any air bubbles (Fig. 5) (see Note 41). 5. Load 75 μL of the library onto the Flow Cell via the SpotON sample port, using a P200 pipette suspended above the port, dispensing in a dropwise fashion. Ensure each drop siphons into the port before adding the next (Fig. 5) (see Note 42). 6. Gently replace the SpotON sample port cover, making sure the bung enters the SpotON port. Close the priming port and close the instrument lid. 7. On the ONT MinKnow software go to the “Start” menu and “Start Sequencing.” 8. Enter experiment name and sample name (see Note 43). 9. Select kits used (i.e., LSK109), and relevant barcoding kit (i.e., NBD104, 114 or 196). 10. Set run options: either set time to 72 h and stop when there is a sufficient amount of data or set for a shorter amount of time depending on coverage required (e.g., 24 h for 20 ng/μl and NFW is