PCR Mutation Detection Protocols

Methods

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PCR Mutation Detection Protocols Second Edition

Edited by

Bimal D.M. Theophilus Department of Haematology, Birmingham Childrens Hospital NHS Foundation Trust, Birmingham, UK

Ralph Rapley Department of Biosciences, University of Hertfordshire, Hatfield, Hertfordshire, United Kingdom

Editor Dr. Bimal D.M. Theophilus Birmingham Childrens Hospital NHS Foundation Trust Department of Haematology Steelhouse Lane B4 6NH Birmingham United Kingdom [email protected]

Dr. Ralph Rapley University of Hertfordshire Department of Biosciences College Lane AL10 9AB Hatfield, Herts. United Kingdom [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-60761-946-8 e-ISBN 978-1-60761-947-5 DOI 10.1007/978-1-60761-947-5 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2010937422 © Springer Science+Business Media, LLC 2011 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)

Preface In the preface to the first edition of PCR Mutation Detection Protocols, we indicated that it was an exciting time for molecular genetics and, in particular, the use of molecular diagnostic techniques for the analysis of DNA and mutations. A number of years on and this has proven to be true and has been reflected in the numerous developments and methods that allow mutation analysis to be currently undertaken. The explosion of DNA sequence information, the access to bioinformatics and mutation databases coupled with the ability to readily detect and confirm mutations has cemented the role of molecular diagnostics in medicine. In particular, mutation detection by the PCR has stood the test of time and has been increasingly adopted by many laboratories and now forms the cornerstone of many DNA diagnostic techniques. Indeed, it is now very difficult to think of molecular diagnostics without the PCR, and it is interesting to see the many ways in which the PCR has been adapted and has evolved over the years. However, the methods that one would think of as having been replaced by the PCR are still very much with us and fulfill an important role. It is a testament to the developers of methods such as Southern blotting that they are part of the panel of techniques with which to identify mutations in DNA. It is, of course, important to include these techniques in this second edition of PCR Mutation Detection Protocols as well as the PCR and its many various incarnations such as SSCP, CSGE, and dHPLC. One theme that is increasingly running through clinical diagnostic laboratories nowadays is demand for accurate diagnostics with high throughput. The increasing casework in many ways reflects the success of molecular diagnostic techniques and their adoption into the diagnostic armory of the clinician. With this in mind, the inclusion of a number of these methods in the second edition of PCR Mutation Detection Protocols is an important one. The increase in DNA analysis requests will no doubt lead to further developments especially in the PCR; however, it is the development of affordable, durable, and accurate microarray systems and their adoption into the clinical laboratory that will ultimately be able to cope with this demand, and a collection of such methods are presented in this edition. As with the first edition of PCR Mutation Detection Protocols, each chapter includes the underlying basis of the method and allows the reader to select and undertake the method successfully. The notes sections with each of the chapters provides the often hard to find information that may mean the difference between success and failure of the method. PCR Mutation Detection Protocols is aimed at postgraduate scientists, researchers, and clinicians already engaged in the area. However, it may also provide an important first step for those wanting to adopt a new technique in their laboratory.

Birmingham, UK Hertfordshire, UK

Bimal D.M. Theophilus Ralph Rapley

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

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1 Conformation-Sensitive Capillary Electrophoresis . . . . . . . . . . . . . . . . . . . . . . . . Emma Jane Ashton 2 Conformation Sensitive Gel Electrophoresis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Marian Hill 3 Denaturing HPLC for Mutation Screening . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mike Mitchell and Jacqueline Cutler 4 In Situ Detection of Human Papillomavirus DNA After PCR-Amplification . . . . . Gerard J. Nuovo 5 LATE-PCR and Allied Technologies: Real-Time Detection Strategies for Rapid, Reliable Diagnosis from Single Cells . . . . . . . . . . . . . . . . . . . . . . . . . . Kenneth E. Pierce and Lawrence J. Wangh 6 Long-PCR Amplification of Human Genomic DNA . . . . . . . . . . . . . . . . . . . . . . Stephen Keeney 7 Human Papilloma Virus Strain Detection Utilising Custom-Designed Oligonucleotide Microarrays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Duncan Ayers, Mark Platt, Farzad Javad, and Philip J.R. Day 8 Multiplex Ligation-Dependent Probe Amplification (MLPA®) for the Detection of Copy Number Variation in Genomic Sequences . . . . . . . . . . Petra G.C. Eijk - Van Os and Jan P. Schouten 9 Screening for Genomic Rearrangements by Multiplex PCR/Liquid Chromatography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Claude Houdayer, Catherine Dehainault, Marion Gauthier-Villars, and Dominique Stoppa-Lyonnet 10 Mutation Surveyor: Software for DNA Sequence Analysis . . . . . . . . . . . . . . . . . . Jayne A.L. Minton, Sarah E. Flanagan, and Sian Ellard 11 Non-invasive Prenatal Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cathy Meaney and Gail Norbury 12 Automated DNA Sequencing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yvonne Wallis and Natalie Morrell 13 Phylogenetic Microarrays for Cultivation-Independent Identification and Metabolic Characterization of Microorganisms in Complex Samples . . . . . . . Alexander Loy, Michael Pester, and Doris Steger 14 Prenatal Detection of Chromosome Aneuploidy by Quantitative-Fluorescence PCR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kathy Mann, Erwin Petek, and Barbara Pertl

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15 Use of Robotics in High-Throughput DNA Sequencing . . . . . . . . . . . . . . . . . . . Stephen Keeney 16 Detection of Factor V Leiden and Prothrombin c.20210G>A Allele by Roche Diagnostics LightCycler® . . . . . . . . . . . . . . . . . . . . Peter C. Cooper 17 RT-PCR for the Detection of Translocations in Bone and Soft Tissue Tumours in Formalin-Fixed Paraffin-Embedded Tissues . . . . . . . Ann Williams and D. Chas Mangham 18 Detection of Minimal Residual Disease in Leukaemia by RT-PCR . . . . . . . . . . . . Joanne Mason and Mike Griffiths 19 Mutation Detection by Southern Blotting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gillian Mellars and Keith Gomez

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Erratum. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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

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Contributors Emma Jane Ashton • DNA Laboratory, Great Ormond Street Hospital NHS Trust, London, UK Duncan Ayers • Manchester Interdisciplinary Biocentre, University of Manchester, Manchester, UK Peter C. Cooper • Department of Coagulation, Sheffield Haemophilia and Thrombosis Centre, Royal Hallamshire Hospital, Sheffield Teaching Hospitals NHS Foundation Trust, Sheffield, South Yorkshire, UK Jacqueline Cutler • Molecular Genetics, Centre for Haemostasis & Thrombosis, St. Thomas’ Hospital, Guy’s & St Thomas’ NHS Foundation Trust, London, UK Philip J. R. Day • Manchester Interdisciplinary Biocentre, University of Manchester, Manchester, UK Catherine Dehainault • Service de Génétique Oncologique, Institut Curie Hôpital, Paris, Paris, France Petra G. C. Eijk - Van Os • Microbiology Research Centre Holland, Amsterdam, The Netherlands Sian Ellard • Department of Molecular Genetics, Royal Devon & Exeter NHS Foundation Trust, Exeter, UK; Peninsula Medical School, Institute of Biomedical and Clinical Science, Exeter, UK Sarah E. Flanagan • Peninsula Medical School, Institute of Biomedical and Clinical Science, Exeter, UK Marion Gauthier-Villars • Service de Génétique Oncologique, Institut Curie Hôpital, Paris, France Keith Gomez • Katharine Dormandy Haemophilia Centre, Royal Free Hospital, Hampstead, London, UK Mike Griffiths • West Midlands Regional Genetics Laboratory, Birmingham Women’s NHS Foundation Trust, Edgbaston, Birmingham, UK Marian Hill • Department of Clinical Pathology, Queens Medical Centre, Nottingham University Hospitals, Nottingham, UK Claude Houdayer • Service de Génétique Oncologique, Institut Curie Hôpital, Paris, Paris, France; Université Paris Descartes, Paris, France Farzad Javad • Manchester Interdisciplinary Biocentre, University of Manchester, Manchester, UK Stephen Keeney • Molecular Diagnostics Centre, Central Manchester University Hospitals Foundation Trust, Manchester, UK Alexander Loy • Department of Microbial Ecology, Faculty of Life Sciences, University of Vienna, Wien, Austria

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D. Chas. Mangham • Department of Musculoskeletal Pathology, Royal Orthopaedic Hospital NHS Trust, Birmingham, UK; Department of Musculoskeletal Pathology, Robert Jones & Agnes Hunt Orthopaedic Hospital NHS Trust, Oswestry, Shropshire, UK Kathy Mann • Cytogenetics Department, Guy’s Hospital, London, UK Joanne Mason • West Midlands Regional Genetics Laboratory, Birmingham Women’s NHS Foundation Trust, Edgbaston, Birmingham, UK Cathy Meaney • State Registered Clinical Scientist (SRCS), Regional Molecular Genetics, Great Ormond Street Hospital, London, UK Gillian Mellars • Katharine Dormandy Haemophilia Centre, Royal Free Hospital, Hampstead, London, UK Jayne A. L. Minton • Department of Molecular Genetics, Royal Devon & Exeter NHS Foundation Trust, Exeter, UK Mike Mitchell • Molecular Genetics, Centre for Haemostasis & Thrombosis, St. Thomas’ Hospital, Guy’s & St Thomas’ NHS Foundation Trust, London, UK Natalie Morrell • West Midlands Regional Genetics Service, Birmingham Women’s NHS Foundation Trust, Edgbaston, Birmingham, UK Gail Norbury • State Registered Clinical Scientist (SRCS), Regional Molecular Genetics, Great Ormond Street Hospital, London, UK Gerard J. Nuovo • Comprehensive Cancer Center, Ohio State University Medical Center, Columbus OH, USA Barbara Pertl • Department of Obstetrics and Gynaecology, Medical University Graz, Graz, Austria Michael Pester • Department of Microbial Ecology, Faculty of Life Sciences, University of Vienna, Wien, Austria Erwin Petek • Department of Obstetrics and Gynaecology, Medical University Graz, Graz, Austria Kenneth E. Pierce • Department of Biology, Brandeis University, Waltham MA, USA Mark Platt • Manchester Interdisciplinary Biocentre, University of Manchester, Manchester, UK Jan P. Schouten • Microbiology Research Centre Holland, Amsterdam, The Netherlands Doris Steger • Department of Microbial Ecology, Faculty of Life Sciences, University of Vienna, Wien, Austria Dominique Stoppa-Lyonnet • Service de Génétique Oncologique, Institut Curie Hôpital, Paris, France; Université Paris Descartes, Paris, France; INSERM U830, Institut Curie Centre de Recherche, Paris, France Yvonne Wallis • West Midlands Regional Genetics Service, Birmingham Women’s NHS Foundation Trust, Edgbaston, Birmingham, UK Lawrence J. Wangh • Department of Biology, Brandeis University, Waltham MA, USA Ann Williams • Department of Musculoskeletal Pathology, Royal Orthopaedic Hospital NHS Trust, Birmingham, UK

Chapter 1 Conformation-Sensitive Capillary Electrophoresis Emma Jane Ashton Abstract Conformation-sensitive capillary electrophoresis (CSCE) is a rapid, high-throughput screening method that can be applied to any region of a genome for detection of sequence variants. Slab gel-based conformation-sensitive gel electrophoresis was first described by Ganguly et al., and the transfer from slab gels to capillaries for higher throughput was reported by Rozycka et al. CSCE is based on the principle that DNA homoduplexes and heteroduplexes migrate at different rates during electrophoresis under mildly denaturing conditions. Fragments showing an altered peak morphology compared to the wild type are then sequenced to determine the precise nature of the sequence variant detected. Key words: Fluorescent multiplex conformation-sensitive capillary electrophoresis, Heteroduplex formation, Fluorescent multiplex polymerase chain reaction, Mutation detection, Capillary electrophoresis

1. Introduction CSCE is based on the principle that DNA homoduplexes and heteroduplexes migrate at different rates during electrophoresis and sequence changes within a fragment can, therefore, be visualized as up to four different peaks (two different homo- and heteroduplexes). The use of fluorescent tags means that multiple fragments can be analyzed simultaneously in one capillary using size and color to differentiate PCR products. Primers are designed to include the sequence of interest plus at least 50 bp either side of this region to maximize detection of sequence variants. Fragments are arranged into multiplexes according to size and color and PCR is carried out, followed by a heteroduplex formation step. The products are then run on a genetic analyzer under mildly denaturing conditions to allow separation of homo- and

Bimal D.M. Theophilus and Ralph Rapley (eds.), PCR Mutation Detection Protocols, Methods in Molecular Biology, vol. 688, DOI 10.1007/978-1-60761-947-5_1, © Springer Science+Business Media, LLC 2011

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heteroduplexes. Fragments showing altered peak morphology compared to the wild type are then sequenced to determine the nature of the variant detected. CSCE can be applied for highthroughput mutation screening (1–3) or SNP typing (4).

2. Materials 2.1. PCR Product Preparation

1. Fluorescently labelled primers (FAM, VIC, NED) – light sensitive. Store stock primers at −20°C or below. Store primer dilutions at 4°C. 2. Immolase DNA polymerase (Bioline, London, England). 3. 10× PCR buffer (Bioline, London, England). 4. 25 mM dNTPs. 5. 50 mM MgCl2. 6. T.01E (10 mM Tris–HCl pH 7.6, 0.1 mM EDTA). Filtersterilize and store at room temperature. 7. 10 mM Tris–HCl pH 7.6. Filter-sterilize and store at room temperature.

2.2. Capillary Electrophoresis

1. CSCE polymer – 5.25% native genescan polymer (Applied Biosystems), 10% glycerol, 5% Hi-Di formamide – toxic (Applied Biosystems) and 1× 3100 running buffer with EDTA (Applied Biosystems). Stir well to mix. Store at 4°C; make fresh every 7 days. 2. Electrophoresis buffer – 1× 3100 running buffer with EDTA. Dilute from 10× stock and store at 4°C; make fresh every 7 days. 3. Genescan 500 Liz size standard (Applied Biosystems).

3. Methods 3.1. Primer and Multiplex Design

1. Design fragments between 200 and 500 bp in length. Include at least 50 bp of sequence either side of sequence of interest. If your sequence of interest is larger than this, split it into two (or more) fragments. If possible, avoid repeat tracts as these tend to give such a complex wild-type peak that detection of other variants is more difficult. (See Note 1). 2. It is preferable to use FAM, VIC, and NED fluorescent labels. It is possible to use PET-labelled fragments; however, there are two isomers giving a more complex peak pattern under mildly denaturing conditions and making mutation detection more complex, so if possible, avoid using this dye.

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3. Ideally, each fragment would resolve as a single wild-type peak, but in practice, some fragments have been found to resolve as a main peak and accompanying second peak in the wild-type state. As long as a consistent wild-type pattern is seen, this does not impede mutation detection. 4. Multiplex fragments according to size and color (up to seven fragments should be straightforward; up to 12 or more is possible, but it is more difficult to ensure even peak heights). Avoid multiplexing fragments of exactly the same size as bleed through between capillaries may interfere with heteroduplex analysis (see Note 1). 3.2. PCR Setup

1. Set up 25 ml PCR reactions as follows: 5 ml genomic DNA (25 ng/ml), 2.5 ml 10× PCR buffer, 1.75 ml MgCl2 (50 mM), 0.5 ml dNTPs (25 mM), 0.5 ml Immolase DNA polymerase (5 U/ml), 0.2 ml of forward and reverse primer for each fragment in the multiplex (5 pmol/ml), T.01E to 25 ml final volume (see Note 2). 2. Carry out a PCR with the following cycling conditions: 95°C 7 min, 95°C 30 s, 55°C 30 s, 72°C 2 min (30 cycles), 72°C 10 min, and 60°C 1 h (see Notes 3 and 4). 3. Heteroduplex formation Form heteroduplexes with the following incubations, either as a separate step or tagged at the end of the PCR reaction: 95°C 10 min, 95°C 1 min decreasing by 1.5°C every minute for 23 cycles, 60°C 30 min. NB: If the sequence of interest is X-linked or if there is any possibility of homozygosity for a sequence variant (e.g., if consanguinity is a possibility), then mix normal control PCR product with test PCR product so that heteroduplexes will form (see Note 5).

3.3. Dilution of PCR Products

Dilution of PCR products is necessary to ensure that the CCD camera is not saturated by fluorescent signal. Overloading will cause flat-topped or split peaks and may cause heteroduplexes to be missed. Minimum and maximum peak heights depend on the genetic analyzer used, but for the ABI 3100, they should ideally be between 200 and 5,000 relative fluorescent units. 1. Dilute PCR products 1:20 in 10 mM Tris–HCl. Mix 2 ml diluted PCR product with 0.3 ml Genescan 500 Liz 500 size standard (see Note 6), 0.2 ml 10× PCR buffer (see Note 7), and 17.5 ml 10 mM Tris–HCl. Pooling of PCR products from single PCR reactions can be carried out at this stage as an alternative to multiplex PCR.

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3.4. Capillary Electrophoresis (See Note 8)

1. If samples are being run for the first time, perform a spectral calibration with the CSCE polymer installed on the genetic analyzer and save this using a letter/number code not designated for other applications (e.g., E5). 2. Run the samples on the genetic analyzer using a 50 cm capillary array, which gives improved heteroduplex detection compared to a 36 cm array. Standard run module Genescan36_POP4 should be used with the following modifications: temperature 18°C, voltage 10 kV, run time 4,200 s, and data delay 500 s (see Notes 9 and 10).

3.5. Data Analysis

1. Products run on the ABI 3100 should be analyzed using Genotyper software (Applied Biosystems) (see Note 11). The size standard will not run true to size due to the mildly denaturing electrophoresis conditions and should only be used for relative peak location. Assign the first three digits of the scan number to each peak, rather than the expected size in base pairs, and save this as the size standard. 2. Some fragments, especially larger ones, have greater altered mobility than others under the mildly denaturing conditions. Ensure that the correct fragment is being analyzed when first setting up an assay by either running PCR products individually or by using mutation/polymorphism controls to ensure correct peak identification. 3. Import the data files and use visual inspection of peaks to identify heteroduplexes. Inspect one fragment at a time (see Note 12). Zoom in and out around each peak for maximum detection of variants. Ensure that the peaks fall between the minimum and maximum peak heights recommended (see Note 13). Ideally, a single peak should be seen for a wild-type fragment, whereas two homoduplex and two heteroduplex peaks should be seen for fragments containing a sequence variant. In practice, patterns range from the expected four peaks to subtle shoulders in peaks or wide peaks and experience is needed to identify variants accurately (see Note 14). 4. Some run-to-run variation for some fragments may occur, and it is, therefore, recommended to only compare samples from the same run. Comparing fragments from different electrophoretic runs may make the analysis more difficult. 5. Monitor the life of the capillary and polymer and the success of each injection using the size standard. The 490 and 500 bp peaks in the Genescan 500 Liz 500 size standard should be visible for each sample. If they are not visible, heteroduplexes may remain undetected. Reinject the sample or renew the array or polymer as necessary. 6. CSCE is not a direct mutation detection method, and fragments showing altered peak morphology should also be analyzed by

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DNA sequencing to determine the nature of variant detected. (See chapter on DNA sequencing).

4. Notes 1. When initially setting up an assay, optimize the multiplexes so that all fragments are of roughly equal strength after PCR. PCR products with the same fluorescent label should be separated by at least 50 bp to avoid confusion over which fragment heteroduplex peaks have been generated by. It may be necessary to alter primer concentrations of some fragments or switch fragments into different multiplexes to obtain an optimal combination. Care at this stage will ensure a more robust assay and easier analysis. 2. PCR reagents from any source can be used as long as the final PCR products are of sufficient (and roughly equal) strength and do not contain any nonspecific peaks, which may interfere with heteroduplex analysis. 3. The final incubation step for 1 h at 60°C ensures that the additional adenosine base has been added to all fragments so that they are all identical in length. 4. 30 cycles of PCR have been found to be optimal to maximize the detection of heteroduplexes. A further increase in cycle number was not found to improve detection. 5. In this case, set up test and normal control PCRs separately, and mix an equal volume of each (e.g. 10 ml each) before the heteroduplex step to ensure the formation of heteroduplexes. Run the normal control and test PCR products separately under nondenaturing conditions first to ensure that the fragments have amplified before mixing, to avoid the possibility of just analyzing the normal control if the test PCR has failed. 6. Use the appropriate size standard as recommended by the manufacturers, e.g., Genescan 500 Liz-500 for the ABI 5-dye system. 7. Further dilution of PCR products does not result in lower peak heights as it also reduces the overall salt concentration in the sample which acts to inhibit electrokinetic injection. Therefore, a reduction in salt concentration results in an increase in the amount of PCR product taken up during electrokinetic injection. Addition of extra salt to the reaction (in the form of PCR buffer) corrects for this. 8. Capillary electrophoresis as described here is specific to the ABI 3100; refer to the manufacturer’s guidelines for other genetic analyzers.

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9. Increased data delay avoids collection of primer flare as the smallest products are 200 bp. Note that the best separation of peaks was found to be produced at 18°C, but higher ambient temperatures may result in poorer separation and the higher temperature will also decrease the life of the polymer (7 days at room temperature). 10. The polymer described here was designed for use on the ABI 3100. The Conformation Analysis Polymer (Applied Biosystems) is designed for use with the ABI 3130 and 3730. Refer to the manufacturer’s instructions regarding suitable polymer as differing system pressures mean that specific polymers may not be suitable for all genetic analyzers. The polymer should be made weekly and the quality monitored with a known mutation to ensure correct separation. 11. Use the specific analysis software appropriate to the genetic analyzer. 12. In our experience, it was found optimal to analyse batches of eight samples, comparing these against each other and overlaying peaks where wider or shoulder peaks are suspected. 13. If samples are too weak, either reduce the amount of PCR buffer in the injection solution or increase the injection time and/or voltage on the genetic analyzer. Do the opposite for samples that are too strong. If this is a recurring problem, reoptimize the PCR multiplex. 14. Initially, it is better to err on the side of caution and sequence all subtle changes until familiar with the data. This will also depend on whether there is a requirement to detect all sequence variants. Polymorphisms will produce a classic pattern, but the same pattern may also be caused by different sequence variants on rare occasions. References 1. Hoskins, B.E., Thorn, A., Scambler, P.J., Beales, P.L. (2003). Evaluation of multiplex capillary heteroduplex analysis: a rapid and sensitive mutation screening technique. Hum Mutat 22: 151–157. 2. Esteban-Cardenosa, E., Duran, M., Infante, M., Velasco, E., Miner, C. (2004). Highthroughput mutation detection method to scan BRCA1 and BRCA2 based on heteroduplex analysis by capillary array electrophoresis. Clin Chem 50: 313–320. 3. Davies, H., Dicks, E., Stephens, P., Cox, C., Teague, J., Greenman, C., Bignall, G., O’Meara,

S., Edkins, S., Parker, A., Stevens, C., Menzies, A., Blow, M., Bottomley, B., Dronsfield, M., Futreal, P.A., Stratton, M.R., Wooster, R. (2006). High throughput DNA sequence variant detection by conformation sensitive capillary electrophoresis and automated peak comparison. Genomics 87: 427–432. 4. Chen, Y.L., Jong, Y.J., Ferrance, J., Hsien, J.S., Feng, C.H., Wu, M.T., Wu, S.M. (2008). Single nucleotide polymorphism detection in the hMSH2 gene using conformationsensitive CE. Electrophoresis 29: 634–640.

Chapter 2 Conformation Sensitive Gel Electrophoresis Marian Hill Abstract Conformation sensitive gel electrophoresis (CSGE) is a rapid screening method for the detection of DNA sequence variation, specifically single-base changes or small insertions and deletions. It has been widely used for mutation screening in genetic disorders and for the detection of single nucleotide polymorphisms (SNPs). CSGE is a simple manual method, based on heteroduplex analysis, and compares well in terms of sensitivity with other screening technologies. CSGE also lends itself to automation and such modi fications have been useful in increasing sample throughput and sensitivity. However, manual CSGE remains a low-cost, accessible, and effective approach for mutation screening, which can be carried out with minimal specialist equipment. This chapter describes manual CSGE, and outlines some of the uses, modifications, and limitations of this method. Key words: CSGE, Heteroduplex, Homoduplex, Mutation screening, Genetic analysis, CSCE, F-CSGE

1. Introduction Conformation sensitive gel electrophoresis (CSGE) was initially developed by Ganguly et al. (1, 2) as a screening method to minimise the amount of nucleotide sequencing required when investigating large genes for mutations. This method is used for the detection of single-base changes or small insertions and deletions within PCR products. It specifically detects heterozygous changes, although homozygous and hemizygous changes are readily detectable when samples are mixed with an equivalent “normal” control. CSGE was initially used to improve the analysis of multiple genes associated with collagen disorders (3), and has been shown to be a highly sensitive tool in the analysis of a large range of inherited genetic disorders (4–13). CSGE has also been widely


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used in the analysis of sequence variations in cancer susceptibility genes such as BRCA1 and BRCA2 (14, 15) and in the study of the MEN1 gene in endocrine neoplasia type 1 (16). CSGE has been compared with other screening methods in a number of studies. Markoff et al. (17) reported that CSGE was better than single-stranded conformation polymorphism (SSCP) for the analysis of mutations in BRCA1 and although these two methods were comparable in a study by Eng et al. (18), they concluded that denaturing high-pressure liquid chromatography (dHPLC) was more sensitive. However, under optimum conditions, manual CSGE has been shown to detect up to 100% of mutations (3) and has the advantage that it can be carried with a minimum of specialist equipment. The requirement for higher throughput testing, particularly for single nucleotide polymorphism (SNP) analysis, has facilitated a number of modifications to the basic CSGE method, to improve both speed and sensitivity (19–21). Higher throughput and sensitivity has also been achieved by the use of fluorescent labels and the automation of CSGE on genetic analysers. Fluorescent-CSGE (F-CSGE) offers improved resolution and reproducibility, and has been developed on both a gel and capillary format (14). Hashemi Soteh et al. (13) compared manual and fluorescent CSGE in the study of mutations in the VWF gene, concluding that F-CSGE was a more sensitive method, allowing higher throughput analysis. The transfer of CSGE screening to multicapillary genetic analysers has increased potential throughput significantly (22–24). Such approaches to mutation screening have been comprehensively evaluated by the UK National Genetics Reference Laboratory, Wessex (25) It is apparent that automated approaches allow higher throughput screening and are more reproducible and sensitive than the manual method, however, they do require specialised equipment and increased cost (14). In addition, the availability of a genetic analyser may make direct nucleotide sequencing a costeffective alternative.

2. Materials 2.1. Samples and Controls for CSGE Analysis

1. CSGE is carried out on PCR products, the design of which is critical to the success of this method (see Notes 1–4). The PCR product must be of good quality and sufficient concentration to be clearly visible on electrophoresis. 2. It is important to include suitable controls when screening, a previously sequenced equivalent PCR product with no sequence variation should always be included as a “normal”

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control, for comparison of band patterns. A previously sequenced “positive” control should also be included. This may be a sample with the same sequence variation when screening for a known change, or may be an unrelated variable. 2.2. Gel Preparation and Electrophoresis

Manual CSGE involves the use of a non-proprietary polyacrylamide gel for electrophoretic separation. These gels are typically manual sequencing format (e.g. 30 × 45 cm) and 1 mm thick. 1. 20× stock TTE buffer: 1.78 M Tris, 570 mM taurine, 4 mM EDTA, pH 9.0. 2. 40% Acrylamide solution. 3. BAP (1,4 bis (acrolyl) piperazine)*. 4. Ethylene glycol*. 5. Formamide*. 6. Ammonium persulphate (10%, freshly prepared)*. 7. TEMED (N,N,N,N¢ tetramethylethylenediamine)*. 8. DNA loading dye: e.g. 30% glycerol, 0.25% bromophenol blue, 0.25% xylene cyanol FF. 9. DNA stain (e.g. ethidium bromide (1 mg/ml) or Gelstar)*. Electrophoresis/ultra-pure grade reagents should be used and all solutions should be prepared in high quality distilled, deionised water. Note: *These chemicals are potentially harmful. Refer to Material Safety Data Sheets (MSDS) and use appropriate handling and disposal procedures.

3. Method CSGE is based on the ability to distinguish between homoduplex and heteroduplex DNA fragments by electrophoresis under partially denaturing conditions. DNA homoduplexes consist of doublestranded DNA fragments in which all the bases are paired correctly with their complementary base on the opposite stand. Heteroduplex DNA contains mismatched bases, and in PCR products that originate from a patient with a heterozygous mutation, both homoduplexes and heteroduplexes can form when double-stranded DNA is allowed to dissociate then reanneal with the complementary strand originating from a different allele (Fig. 1). The presence of mismatched bases induces subtle conformational changes in the heteroduplex compared with the homoduplex as the misaligned bases do not conform to the typical Watson– Crick base-pairing rules.

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Fig. 1. Illustration of heteroduplex and homoduplex generation. A PCR product from a patient who is heterozygous (A/C) at a specific nucleotide position will contain two species of double-stranded DNA. Heating to 98°C will dissociate the double-stranded DNA, incubation at 65°C allows the strands to reanneal. Heteroduplexes are formed when a strand from one allele reanneals with the complementary strand from the other allele, and will contain a mismatched base pair.

The CSGE protocol involves the generation of heteroduplexes and/or homoduplexes in PCR products through heating and slow reannealling, followed by gel electrophoresis on a large format polyacrylamide gel, cross-linked with BAP (1,4 bis (acrolyl) piperazine) – which greatly improves gel strength and increases conductivity (1). The gel also contains ethylene glycol and formamide, which act as mild denaturants. Under these conditions, heteroduplexes can be resolved from homoduplexes as they generally migrate more slowly through the gel matrix. Bands are visualised by staining with a DNA stain such as ethidium bromide. Multiple bands will be detectable in samples containing “heterozygous” changes, while a single-homoduplex band is generally visible in samples containing no sequence variation. Like most screening methods, CSGE gives limited information on the nature of the sequence variation, and further analysis, usually by nucleotide sequencing is essential for identification. 3.1. Sample Mixing and Heteroduplex Formation

1. In order to also detect homozygous changes (or e.g. hemizygous changes in X-linked disorders), the PCR product must be mixed with an equivalent “normal” control. Mix the test sample 1:1 with a previously sequenced male control sample for the detection of hemizygous changes in X-linked disorders,


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or a 2:1 mix (test:normal control) in autosomal disorders. Where a number of different fragments are to be screened, PCRs may be multiplexed, or products pooled, and then analysed simultaneously – providing the different PCR products can be easily differentiated. 2. For heteroduplex formation, incubate the PCR product mix (10 ml) in a thermocycler at 98°C for 5 min then 65°C for 30 min, followed by a slow cool down to room temperature. This should be done immediately before electrophoresis. 3.2. Gel Preparation

1. Ensure that gel electrophoresis plates are clean and grease-free. 2. Prepare a gel solution consisting of: 44 ml of 99:1 acrylamide:BAP (1,4 bis (acrolyl) piperazine) (10% final concentration) 17.5 ml of ethylene glycol (10% final concentration) 26 ml of formamide (15% final concentration) 4.4 ml of TTE buffer 81 ml distilled, deionised water 3. Initiate polymerisation by adding 1.75 ml of 0.1% ammonium persulphate and 100 ml of TEMED; pour the gel immediately. 4. Allow a minimum 1 h for polymerisation.

3.3. Electrophoresis and Staining

1. Add 2 ml loading dye to 10 ml PCR product mix and load samples onto the gel in a standard loading buffer. 2. Carry out electrophoresis, 0.5× TTE buffer, typically for 16 h at 400 V (1). 3. After electrophoresis, carefully remove one of the glass plates and place the gel (on the remaining plate) in an appropriate container. Pour ethidium bromide solution (1 mg/ml in 0.5× TTE) onto the gel, and allow up to 30 min to stain. 4. To transfer the gel to the UV imaging system, blot with Whatman filter paper, carefully wet the filter paper with water to release from the gel when in position. 5. Visualise the separated products under UV illumination.

3.4. Analysis

Homoduplexes are generally detected as a single band, one or more additional bands representing the Co-Migrating heteroduplexes may be seen if a mismatch is present. Comparison to a “normal” control is important to avoid false-positive results, as additional bands may also be seen due to secondary structure. A detectable and consistent CSGE band pattern should be noted in the positive controls used. As changes in band pattern may be subtle, experience is invaluable for the interpretation of results and optimum PCR product design (see Note 5).

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Fig. 2. Illustration of variation in band patterns seen in CSGE. Investigation of F11 gene, exon 9. Lane 1: normal control, lane 2: patient with a single-base deletion (g), lane 3: patient with t → c substitution.

Figure 2 illustrates the CSGE band patterns associated with two different mutations (1 bp deletion (−g) and a t → c substitution) in exon 9 of the F11 gene. Although the heteroduplex band/s is commonly seen above the homoduplex band, this type of pattern is not always seen, as illustrated in lane 3. The pattern of bands is highly variable depending on the type of mutation and the sequence context although insertions and deletions tend to produce the largest band separation as they effect a larger conformation change.

4. Notes 1. The exact nature of the mismatch, size of the PCR product, the location, and sequence surrounding the mismatch (sequence context) will all affect the sensitivity of CSGE. The reported detection rate of this method has ranged from 60% of BRCA1 mutations in a study co-ordinated by Eng et al. (18) to 100% in a number of studies, including those of Korkko et al. (3). 2. Optimal size for the PCR product is 200–500 bp, although sequence mismatches have been detected in products up to 800 bp in length. Size is limited by the inherent flexibility of DNA, which may mask any conformation change due to mismatch (26). Korkko et al. (3) suggested that PCR products should be limited to below 450 bp for optimum sensitivity, and demonstrated that a single-base polymorphism in the


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COL1A2 gene which could not be detected in a 755 bp product was clearly seen when PCR primers were redesigned to reduce the size to 276 bp. 3. The band pattern seen is dependent on the nature of the mismatch and the surrounding nucleotide sequence (sequence context). Ganguly et al. (27) analysed the ability of CSGE to distinguish specific mismatches within the same sequence context, and found the following order of sensitivity G:G = G:T = T:G > G:A = A:G = T:T > A:A > C:T > C:C = C:A = A:C = T:C. Initial studies with CSGE by Ganguly et al. (1) suggested that mismatches within high temperature melting domains may be particularly difficult to resolve. However, the same mutations were detectable in later studies when primers were redesigned further away (3). It is generally accepted that CSGE is more sensitive to mismatches within an AT-rich sequence context than a GC-rich region (27). 4. Mismatches which are close to the end of the PCR product are less easily detected. Ganguly et al. (1) initially failed to detect a mismatch that was located 51 bp from one end of the PCR fragment. This became detectable when the primers were redesigned to position the mismatch 81 bp from the end. It is therefore advisable to allow for 50–100 bp of additional sequence at either end of the region of interest when designing primers for CSGE. A tagged primer system can be used (e.g. add 5¢ M13 universal sequence tags to primers), which will facilitate sequencing reactions where multiple regions are being analysed (25). Figure 3 illustrates the effect of mismatch position on detection. Primers were redesigned to position a c → a substitution within exon 5 of the F11 gene at 174, 70, and 42 bp from the end of the PCR product. Size was maintained at 290–292 bp. The ability to detect the mismatch is lost as the position of the mismatch approaches the end of the PCR product. 5. CSGE has some limitations as it can only detect single-base changes and small insertions or deletions. However, this method has been successfully used as part of a screening protocol for disorders associated with a wide range of mutation types (28). In common with most screening methods, CSGE provides limited information on the nature of a previously unknown sequence variation or its significance, and may not distinguish between two closely positioned sequence variations. Although band patterns associated with specific mismatches within the same PCR product are reproducible, it is advisable to sequence to confirm the nature of the mismatch.

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Fig. 3. Illustration of effect of position on the ability of CSGE to detect mismatches. Investigation of F11 gene, exon 5. Primers were redesigned to position a c → a substitution at different points from the end of the PCR product. Size was maintained at 290– 292 bp. Lane 1: mismatch is 174 bp from the end, lane 2: mismatch is 70 bp from the end, and lane 3: mismatch is 42 bp from the end.

CSGE is particularly useful for genes that are not very polymorphic such as F8. The sequencing load may increase significantly where highly polymorphic genes such as BRCA1 are analysed but it remains a powerful tool as it greatly reduces the amount of sequencing required during investigation of such genes (2). References 1. Ganguly, A., Rock, M. J., and Prockop, D. J. (1993) Conformation-sensitive gel electrophoresis for rapid detection of single-base differences in double-stranded PCR products and DNA fragments: evidence for solventinduced bends in DNA heteroduplexes. [erratum appears in (1994) Proceedings of the National Academy of Sciences of the United States of America May 24;91(11), 5217]. Proceedings of the National Academy of Sciences of the United States of America 90, 10325–10329. 2. Ganguly, A., and Williams, C. (1997) Detection of mutations in multi-exon genes: comparison of conformation sensitive gel electrophoresis and sequencing strategies with respect to cost and time for finding mutations. Human Mutation 9, 339–343. 3. Korkko, J., Annunen, S., Pihlajamaa, T., Prockop, D. J., and Ala-Kokko, L. (1998) Conformation sensitive gel electrophoresis for simple and accurate detection of mutations: comparison with denaturing gradient gel

electrophoresis and nucleotide sequencing. Proceedings of the National Academy of Sciences of the United States of America 95, 1681–1685. 4. Zhou, J., Kherani, F., Bardakjian, T. M., Katowitz, J., Hughes, N., Schimmenti, L. A., Schneider, A., and Young, T. L. (2008) Identification of novel mutations and sequence variants in the SOX2 and CHX10 genes in patients with anophthalmia/microphthalmia. Molecular Vision 14, 583–592. 5. Blesa, J. R., Solano, A., Briones, P., PrietoRuiz, J. A., Hernandez-Yago, J., and Coria, F. (2007) Molecular genetics of a patient with Mohr–Tranebjaerg Syndrome due to a new mutation in the DDP1 gene. Neuromolecular Medicine 9, 285–291. 6. Zhang, X. L., Liu, M., Meng, X. H., Fu, W. L., Yin, Z. Q., Huang, J. F., and Zhang, X. (2006) Mutational analysis of the rhodopsin gene in Chinese ADRP families by conformation sensitive gel electrophoresis. Life Sciences 78, 1494–1498.

Conformation Sensitive Gel Electrophoresis 7. Blesa, J. R., Prieto-Ruiz, J. A., and HernandezYago, J. (2004) Conformation-sensitive gel electrophoresis as an ideal high-throughput strategy for accurate detection of sequence variations in DNA: screening hTomm and hTimm genes. Journal of Biomolecular Screening 9, 621–624. 8. Korkko, J., Kaitila, I., Lonnqvist, L., Peltonen, L., and Ala-Kokko, L. (2002) Sensitivity of conformation sensitive gel electrophoresis in detecting mutations in Marfan syndrome and related conditions. Journal of Medical Genetics 39, 34–41. 9. Finnila, S. H. I., and Majamaa, K. (2001) Phylogenetic analysis of mitochondrial DNA in patients with occipital stroke. Evaluation of mutations by using sequence of the entire coding region. Mutation Research 458, 31–39. 10. Hill, M., Deam, S., Gordon, B., and Dolan, G. (2005) Mutation analysis in 51 patients with haemophilia A: report of 10 novel mutations and correlations between genotype and clinical phenotype. Haemophilia 11, 133–141. 11. Santacroce, R., Acquila, M., Belvini, D., Castaldo, G., Garagiola, I., Giacomelli, S. H., Lombardi, A. M., Minuti, B., Riccardi, F., Salviato, R., Tagliabue, L., Grandone, E., Margaglione, M., and AICE-Genetics Study Group. (2008) Identification of 217 unreported mutations in the F8 gene in a group of 1,410 unselected Italian patients with hemophilia A. Journal of Human Genetics 53, 275–284. 12. Fard-Esfahani, P., Lari, G. R., Ravanbod, S., Mirkhani, F., Allahyari, M., Rassoulzadegan, M., and Ala, F. (2008) Seven novel point mutations in the F11 gene in Iranian FXIdeficient patients. Haemophilia 14, 91–95. 13. Hashemi Soteh, M., Peake, I. R., Marsden, L., Anson, J., Batlle, J., Meyer, D., Fressinaud, E., Mazurier, C., Goudemand, J., Eikenboom, J., Goodeve, A., and MCMDM-1VWD Group. (2007) Mutational analysis of the von Willebrand factor gene in type 1 von Willebrand disease using conformation sensitive gel electrophoresis: a comparison of fluorescent and manual techniques. Haematologica 92, 550–553. 14. Ganguly, T., Dhulipala, R., Godmilow, L., and Ganguly, A. (1998) High throughput fluorescence-based conformation-sensitive gel electrophoresis (F-CSGE) identifies six unique BRCA2 mutations and an overall low incidence of BRCA2 mutations in high-risk BRCA1-negative breast cancer families. Human Genetics 102, 549–556.

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15. Spitzer, E. A. M., Schmidt, F., Hauser, H., Buwitt, U., Lauter, F. R., Potschick, K., Krocker, J., Elling, D., and Grosse, R. (2000) Detection of BRCA1 and BRCA2 mutations in breast cancer families by a comprehensive two-stage screening procedure. International Journal of Cancer 85, 474–481. 16. Arancha, C. R.-L. S., Cascon, A., Osorio, A., Martinez_Delgado, B., Benitez, J., and Robledo, M. (2002) A rapid and easy method for multiplex endocrine neoplasia type 1 mutation detection using conformation-sensitive gel electrophoresis. Journal of Human Genetics 47, 190–195. 17. Markoff, A., Sormbroen, H., Bogdanova, N., Preisler-Adams, S., Ganev, V., Dworniczak, B., and Horst, J. (1998) Comparison of conformation-sensitive gel electrophoresis and single-strand conformation polymorphism analysis for detection of mutations in the BRCA1 gene using optimized conformation analysis protocols. European Journal of Human Genetics 6, 145–150. 18. Eng, C., Brody, L. C., Wagner, T. M., Devilee, P., Vijg, J., Szabo, C., Tavtigian, S. V., Nathanson, K. L., Ostrander, E., Frank, T. S., and Steering Committee of the Breast Cancer Information Core (BIC) Consortium. (2001) Interpreting epidemiological research: blinded comparison of methods used to estimate the prevalence of inherited mutations in BRCA1. Journal of Medical Genetics 38, 824–833. 19. Leung, Y. F., Tam, P. O. S., Tong, W. C., Baum, L., Choy, K. W., Lam, D. S. C., and Pang, C. P. (2000) High through-put conformation-sensitive gel electrophoresis for discovery of SNPs. Biotechniques 30, 334–340. 20. Fard-Esfahani, P., Khatami, S., Zeinali, C., Taghikhani, M., and Allahyari, M. (2005) A modified conformation sensitive gel electrophoresis (CSGE) method for rapid and accurate detection of low density lipoprotein (LDL) receptor gene mutations in Familial Hypercholesterolemia. Clinical Biochemistry 38, 579–583. 21. Herzog, J. S., Jancis, E. M., Liao, S., Somlo, G., and Weitzel, J. N. (2002) Restriction endonuclease fingerprinting enhanced conformation sensitive gel electrophoresis (REFCSGE) in the analysis of BRCA1 exon 11 mutations in a high-risk breast cancer cohort. Human Mutation 19, 656–663. 22. Velasco, E., Infante, M., Duran, M., EstebanCardenosa, E., Lastra, E., Garcia-Giron, C., and Miner, C. (2005) Rapid mutation detection in complex genes by heteroduplex analysis with capillary array electrophoresis. Electrophoresis 26, 2539–2552.

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23. Velasco, E., Infante, M., Duran, M., PerezCabornero, L., Sanz, D. J., EstebanCardenosa, E., and Miner, C. (2007) Heteroduplex analysis by capillary array electrophoresis for rapid mutation detection in large multiexon genes. Nature Protocols 2, 237–246. 24. Davies, H., Dicks, E., Stephens, P., Cox, C., Teague, J., Greenman, C., Bignell, G., O’Meara, S., Edkins, S., Parker, A., Stevens, C., Menzies, A., Blow, M., Bottomley, B., Dronsfield, M., Futreal, P. A., Stratton, M. R., and Wooster, R. (2006) High throughput DNA sequence variant detection by conformation sensitive capillary electrophoresis and automated peak comparison. Genomics 87, 427–432.

25. http://www.ngrl.org.uk/wessex/csce.htm (2007) Conformation sensitive capillary electrophoresis (NGRLW_CSCE_3.1). National Genetic Reference Laboratories 26. Calladine, C. R., Collis, C. M., Drew, H. R., and Mott, M. R. (1991) A study of electrophoretic mobility of DNA in agarose and polyacrylamide gels. Journal of Molecular Biology 221, 981–1005. 27. Ganguly, A. (2002) An update on conformation sensitive gel electrophoresis. Human Mutation 19, 334–342. 28. Ashton, E. J., Yau, S. C., Deans, Z. C., and Abbs, S. J. (2008) Simultaneous mutation scanning for gross deletions, duplications and point mutations in the DMD gene. European Journal of Human Genetics 16, 53–61.

Chapter 3 Denaturing HPLC for Mutation Screening Mike Mitchell and Jacqueline Cutler Abstract Denaturing High-Performance Liquid Chromatography (dHPLC) is probably the most versatile and one of the most widely used mutation screening technologies. It benefits from a combination of relative technical simplicity and a very high sensitivity (mutation detection rate), approaching 100%. DHPLC can reliably detect single-base mismatches in fragments between 150 and 500 bp, although detection in fragments up to 1,500 bp has been reported. The ability of dHPLC to detect both known and unknown mutations/SNPs, and its’ high sensitivity and specificity (reproducibility) has put this technology at the forefront of genetic analysis for a wide variety of diseases. Key words: dHPLC, Mutation screening, Mutation detection, SNP detection, Gene analysis, Heteroduplex, WAVE®, RP-HPLC

1. Introduction The success of the Human Genome Mapping Project (HGMP) has resulted in significant advances in the field of human genetics and disease. This has led to an increasing number of, and increasingly large and more complex, genes being screened for unknown and heterogeneous mutations. This, in turn, has necessitated the development of rapid, high-throughput mutation screening techniques. In less than 10 years since its conception in the late 1990s (1–4), dHPLC has emerged as one of the most popular and versatile technologies for the analysis of genetic variations. Utilising the principles of heteroduplex analysis and reverse-phase HPLC, dHPLC can detect a single base difference in amplicons greater than 1 kb in size. This user-friendly technology has been exploited for the genetic analysis of a large number of genes involved in a diverse spectrum of diseases (5–13); in fact, more than 350 human genes have been


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analyzed by dHPLC. DHPLC has been shown to be more sensitive than most other technically straightforward, high throughput methodologies, e.g. Single-Stranded Conformational Polymor phism analysis (SSCP), with potential detection rates of >95–100%, along with excellent and consistent specificity. In addition to mutation screening, dHPLC, or ion-pair reversed-phase high-performance liquid chromatography (IP-RP-HPLC), has also been utilised for single nucleotide polymorphism (SNP) analysis, fragment sizing, oligonucleotide purification, and RNA “clean-up” (14–17), amongst other applications. DHPLC has also been utilised for applications beyond the scope of other mutation screening methodologies or even routine sequencing, e.g. the analysis of somatic mutations and the detection of early cancers (18–22). 1.1. Principal of dHPLC

Denaturing HPLC is one of a number of mutation screening techniques which utilises heteroduplex formation. This involves the mixing of fragments of DNA (PCR amplicons) from two different sources, in a 1:1 ratio, heat denaturing the fragments and cooling slowly to allow double-stranded fragments to reform. This process allows not only the original strand pairs to re-anneal and form homoduplexes but also complementary strands from the wild-type and patient amplicons to form heteroduplexes. If there is sequence variation between the two fragments, the heteroduplex will contain a mismatch (Fig. 1). Using temperature to partially denature the double-stranded fragments, and to emphasise the mismatch, dHPLC utilises IP-RP-HPLC to detect differences between homo- and heteroduplexes. The double-stranded DNA binds to a hydrophobic column, the stationary phase, via an intermediary ion pairing agent, (usually triethylammonium acetate, TEAA). An increasing gradient of an organic solvent (usually acetonitrile, ACN), the mobile phase, is then passed over the column. This gradually removes the TEAA and results in the elution of the bound DNA from the column. Under non-denaturing conditions, this elution is size-dependent and sequence-independent. However, under partial denaturing conditions, achieved by elevated temperature,

Fig. 1. A cartoon showing the four possible duplex configurations when a sequence variant is present. Reproduced with permission from Transgenomic.

Denaturing HPLC for Mutation Screening

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Fig. 2. DNA is bound to the column via an interaction between the phosphate groups on the backbone of the DNA and the positive ions of the TEAA which interacts hydrophobically with the beads that pack the column. Under temperature-induced partially denaturing conditions, the DNA begins to lose is helical structure in a sequence-dependent manner and is eluted from the column. Reproduced with permission from Transgenomic.

elution is affected by the helical structure of the double-stranded DNA fragments (Fig. 2). Fragments containing mismatches (heteroduplexes) retain less helical structure and are consequently eluted from the column earlier than are homoduplexes (Fig. 3). The eluted DNA is normally detected by UV absorption, although fluorescent detection can be added if desired. The fragments pass through the detector, which measures the absorbance/ fluorescence over time. Proprietary software interprets the data as a chromatogram. If the paired samples have the same genotype for the particular amplicon, the four homoduplex DNA fragments will elute off the column at the same time producing a single peak on the chromatogram. If a mutation is present, two to four peaks will be visible (Fig. 4). A third operating mode employs fully denaturing conditions, i.e., temperatures above 75°C, and permits the sizing of singlestranded DNA and /or RNA. This can be used for quality control of primers, or for purification of fragments.

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Fig. 3. The effect of temperature on the separation of homo- and heteroduplexes. Below 53°C and above 59°C, there is no differentiation between homo and heteroduplexes. At 53°C, the separation of heteroduplexes begins, while at 55°C, the homoduplexes begin to denature. Optimal separation is achieved at 56°C.

2. Materials 2.1. The Equipment

Although it is theoretically possible to construct a dHPLC system “in-house,” the bespoke polymer chemistries required mean that the vast majority of dHPLC analysis is performed on one of the two commercially available systems, the WAVE® system from Transgenomic Inc. (Omaha, NE, USA) or the Helix system (Varian Inc., Walnut Creek, CA, USA), with the WAVE® being the market leader. Whilst being as generic as possible, the information in this chapter is based on the WAVE® HT system. The dHPLC system comprises several units controlled via a PC. A pump regulates the flow path, allowing reagent gradients to be established. The samples are introduced into the flow path from a chilled autosampler, move through the system to a temperature controlled oven housing the column, packed with either alkylated poly(styrene-divinylbenzene) particles (the WAVE® system) or 1,000 Å alkylated silica (the Helix system), and are eluted from


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Fig. 4. Complete resolution of homo- and heteroduplexes. The earlier eluting peak of the two homoduplexes will comprise the AT homoduplex as it is always slightly more denatured than the GC homoduplex. Reproduced with permission from Transgenomic.

the column and detected by either a UV or a fluorescent detection system. An optional addition to the system, the accelerator negates the requirement for a long equilibration period and a long column wash out. This both shortens the analysis time and extends the column life. Ferrous metals can cause rapid deterioration in the sensitivity of the system. It is, therefore, important that all parts of the flow path that come into contact with the sample are rust free. The materials of choice for these components are polyether ether ketone (PEEK), titanium, fluorocarbon, and sapphire. Whenever the column or in-line filter is removed from the system, the integrity of the flow path should be maintained by the insertion of a connector (PEEK union). 2.2. Reagents

Four separate buffers are required for dHPLC analysis, and the quality of their manufacture is critical for the sensitivity and reproducibility of the data produced.

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The buffers can be bought in premixed or made in-house. The benefits of premixed buffers are that they are manufactured under sterile conditions and are bottled under inert gas to prevent bacterial contamination. This results in reduced inter-batch variation, and a longer shelf life. Analytical chemistry precision must be used for the in-house manufacture of buffers. The quality of water used is critical: it must have a resistivity of 18.2 MW and a total organic content of less than 5 ppm and should be stored for no longer than 2 weeks before use. Buffers made with poor-quality water may damage the column, resulting in a significant reduction in its performance and life expectancy. The handling of glassware used in buffer manufacture is important. It is not necessary to regularly autoclave glassware, although it should be done at least annually. Post autoclaving, all glassware must be thoroughly rinsed with high-quality deionised water to remove any positively charged ions. ACN is highly flammable and is classified as slightly/moderately toxic. It must be stored appropriately, according to national and local regulations, and all buffers containing ACN should be made in a fume hood. All ACN waste must be disposed of as hazardous solvent waste and must not be allowed to enter the water supply. 1. Buffer A: Measure 50 ml TEAA in a volumetric flask. Transfer to a 1-l flask and wash out the 50-ml flask two to three times with water, adding this to the TEAA. Add 250 ml ACN to prevent bacterial growth. Make the final volume up to 1-l with water. Assign a 1-week shelf life 2. Buffer B: Measure 50 ml TEAA in a volumetric flask. Transfer to a 1-l flask and wash out the 50-ml flask two to three times with water, adding this to the TEAA in the 1-l flask. Measure 250 ml ACN. Transfer to the same 1-l volumetric flask and wash out the 250-ml flask twice with water, adding this to the 1-l flask. This begins an endothermic reaction: The flask will feel cold to the touch. Allow to equilibrate for at least 20 min, inverting the flask regularly until gaseous release has finished. Make the final volume up to 1 l with water. Assign a 1-week shelf life. 3. 8% Acetonitrile: Measure 80 ml ACN. Transfer to a 1-l volumetric flask and wash out the measuring cylinder two to three times with water, adding this to the 1-l flask. Make the final volume up to 1 l with water. Assign a 1-week shelf life. 4. 75% Acetonitrile: Measure 750 ml ACN and transfer to a 1-l volumetric flask. Wash out the measuring vessel with water, and add this to the 1-l flask. This begins an endothermic reaction. Allow the mix to


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equilibrate for at least 20 min, inverting the flask regularly until gaseous release has finished. Make the final volume up to 1 l with water. Assign a 3-week shelf life. 2.3. Software

Successful separation of homo- and heteroduplexes is based as much on the run temperature as on the sequence variation. The run temperature is calculated from an in silico reference sequence to reflect the point at which the amplicon has begun to denature, but where 75% still maintains a helical structure (Fig. 5). Less stable

Fig. 5. Melt profiles, generated by Navigator™ software, for determination of the optimal run temperature for partial denaturing analysis. (a) Shows a simple melt profile: a single run temperature should enable detection of all sequence variants in this 150 base pair (bp) fragment. (b) Shows a complex melt profile: sequence variants in the 275–420 bp region of this 800 bp amplicon will require a different run temperature than the rest of the fragment. Some complex melt profiles may require up to five temperatures to resolve a 75% helical fraction for the entire amplicon.

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heteroduplexes will “unravel” at a lower temperature than homoduplexes, and will denature from the point of mismatch outwards. Melt profiles can be calculated directly on the system software (e.g., Navigator™ on the WAVE® system) or by the use of freely available programs such as the dHPLC melt program available from The Stanford Genome Technology Center (http:// insertion.stanford.edu/melt.html), or MutationDiscovery.com (www.mutationdiscovery.com), a freely available resource developed and maintained by Transgenomic, Inc. (Omaha, USA). This site contains the genomic DNA sequences of more than 10,000 genes, the recorded sequence variations within those genes and PCR and dHPLC protocols for amplicons within some of those genes. It is a useful resource for those new to dHPLC and looking for a “quick start” for a gene in which analysis conditions have already been defined. Optimisation of the required melt profile is critical for achieving a high mutation detection rate, and it is worth investing time on this aspect of the protocol. A single temperature may be insufficient to achieve a 75% helical fragment across an entire amplicon (see Note 1). It may be beneficial to re-design primers to obtain better optimisation under certain conditions (see Notes 2–4). 2.4. Steps Required Prior to dHPLC Analysis 2.4.1. PCR Amplification

2.4.2. Heteroduplex Formation

DNA extracted by most routine techniques is suitable for dHPLC, although some post-extraction clean-up may be required (see Note 5). Samples are PCR amplified by standard techniques (see Notes 6 and 7) in a reaction volume of 50 ml. The WAVE® system requires a minimum injection volume of 5 ml, and amplicons often require analysis at multiple temperatures (multiple injections) so low-volume PCRs are not practical for dHPLC analysis. The addition of some PCR enhancers should be avoided completely (see Note 8), while others (see Note 9) should be used in low concentrations only. The efficiency of the amplification is determined by agarose gel electrophoresis. No post-PCR modification is required. For SNP analysis, a positive control as well as a confirmed normal DNA control should be included for each gene fragment amplified. For unknown mutation screening, this may not be possible or practical, as positive controls may be unavailable, and a “normal” control may contain polymorphic SNPs. Each sample is paired with another of approximately equal quantification, the amplification yield estimated from visualisation on an agarose gel. Patient/test samples are matched with either a known normal genotype or a second, unrelated, patient (see Note 10). The mixed pair are subjected to heating and controlled-rate cooling, to produce homo- and (if a sequence variant is present) heteroduplexes (Fig. 6).


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Fig. 6. A cartoon showing the formation of heteroduplexes by temperature-controlled hybridisation. The point of mismatch is shown as a bulge and represents the point of origin for heteroduplex denaturation. Reproduced with permission from Transgenomic.

Thirteen microlitres of each post-PCR DNA are combined into pairs of approximately equal band intensity in labeled 0.2-ml PCR tubes or a 96-well plate. Tubes are transferred to a thermal cycler and the samples heteroduplexed by heating to 95°C and then cooling to room temperature in a controlled manner at a rate of ~1.5°C per minute (see Note 11). A control sample should be included in this stage of the protocol (see Note 12). Alternatively, samples can be heated to 95°C and then allowed to cool slowly on the bench. However, this is a less-efficient heteroduplexing approach as the cooling rate is not controlled. NB/Faster protocols for heteroduplex formation may result in impaired or inefficient heteroduplexing. The heteroduplex formation is a vital part of the process and as worthy of optimisation as the PCR and downstream analysis stages. Following the formation of heteroduplexes, samples should be analysed immediately or stored at −20°C (see Note 13).

3. Methods 3.1. Analysis on the WAVE System

1. Heteroduplexed samples are transferred to the dHPLC autosampler. 96-well plates will fit directly into the chilled autosampler tray, while the lids should be cut off 0.2-ml tubes prior to their positioning (see Note 14). If performing highthroughput analysis, measures need to be in place to minimise sample evaporation, such as reducing the temperature of the autosampler or using foil or seals with 96-well plates (see Note 15).

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2. A run sheet is created according to specific software and local operation instructions, and the injection list ordered to operate sequentially from lowest to highest temperature. This is the most efficient protocol for the oven and offers the fastest run time. 3. Before starting, it is important to ensure that there are sufficient volumes of buffer A and B to complete the run (see Note 16). 4. The operator will be alerted to any problems as the analyser initialises, equilibrates or begins data processing. Errors can occur with any part of the system and may reflect component failure or problems with the flow path (see Note 17). 5. At the end of each run, the operator should ensure that there is sufficient buffer to prevent the column from running dry (see Note 18). 3.2. Data Analysis

It is not necessary to delay data analysis until after a run has been completed. Each injection is processed independently and is available for interpretation immediately. It is recommended that the chromatograms for the first injections are viewed as soon as possible to confirm successful processing, as not all problems will affect the sample flow through the system and so will not generate an error message (see Notes 19 and 20). Chromatograms for all unknown and control samples are overlaid for each amplicon specific method. All samples showing “wild-type” sequence should be resolved as a single peak, whereas samples varying from normal should display between two and four distinct peaks depending on the completeness of optimisation (Fig. 7).

Fig. 7. Overlaid chromatograms for a homoduplex (dotted line) and a heteroduplex (solid line). The homoduplex resolves as a single peak, while there are four distinct peaks for the heteroduplex.


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All samples that generate a chromatogram different from the normal trace should be investigated by sequence analysis. Changes in retention time do not accurately predict the presence of a sequence change, and the absence of a change in numbers of peaks should be interpreted with caution. All samples with the same sequence alteration should display the same peak pattern, making this system applicable to SNP detection (see Note 21). Where multiple temperature injections are required to resolve an amplicon, a sequence variant may only be detectable at one temperature. For larger fragments, the temperature of detection may enable the position of the sequence variant to be predicted, permitting the subsequent use of internal sequencing primers for identification. Navigator™ software permits the simultaneous analysis of data from one or more runs. Current versions of this software have a search facility for easy data retrieval, as runs can be searched on sample name and method name as well as by the run date (see Note 22). Data can be interpreted manually, or automatically via the “mutation calling” function of Navigator™. This function aligns all selected chromatograms, using the largest peak in each as a reference point. The software “normalises” the chromatograms, i.e., it aligns and reconfigures all traces to a standard 1 mV intensity with matched time points. The result is that when comparing a batch of results, similar traces are grouped together. This permits easy segregation of wild-type and mutation samples, again a useful function for SNP detection. This function should not, however, be used exclusively: It is important that all traces are viewed in an unmodified state, as miscalls can occur if there is excessive variation in intensity (peak height) between sample and normal control homoduplexes (see Note 23). If subsequent sequence analysis of an unknown sample does not identify a variant, despite a clear difference in peak profile, there may be a non-pathologic alteration in the “normal” partner of the heteroduplex. This can normally be visualised by the use of two normal controls heteroduplexed together, as well as with test/unknown samples. 3.3. Regular Maintenance Required to Maximise Column Life and Resolution

1. A 75% ACN wash should be run after every 100 injections to prevent build-up of residue in the column (see Note 24). The wash protocol, like all methods, will only run if linked to a well position. As no sample is injected into the flow path, it can be associated with any test well. 2. The in-line filter should be replaced every 1,000–1,500 injections. 3. A reverse hot wash should be performed if loss of resolution is noticed (see Note 25). This will clear any residual DNA from the column.

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4. No analyser, including a dHPLC system, will work optimally if left switched on but idle for long periods of time. If the system is not to be used for a week or more, the column, lines, and in-line filter should be flushed through with 75% ACN before the column is removed and the equipment switched off. After removal of the column, the flow path should be maintained by a PEEK union. Before switching on again, the column must be re-installed. The pump should not be switched on until the oven temperature reaches 50°C, to avoid pressure fluctuation, which can damage the column. It is important that the 75% ACN is washed from the system before any sample is run. The system should be flushed with a 1:1 ratio of buffer A and buffer B for 45–60 min at a flow rate of 0.9 ml/min. The syringe, needle, and injection port also need to be flushed with a 1:1 ratio of buffer A and buffer B before use. 5. Mutation standards are available to verify the oven sensitivity (a range of standards designed to give reproducible elution patterns at a preset elevated temperature) and buffer composition (standards designed to work under non-denaturing conditions). It is essential that these standards form part of the routine maintenance of the instrument; they should be run after every buffer change, routine maintenance event, and switching off/on of the analyser. 3.4. Notes

1. Commercial recommendations are that injection temperatures should increase by no less than 2°C. However, personal experience has shown that this may compromise detection rates. 2. The optimal amplicon size for mutation detection is 150–500 base pairs. If the detection rate is lower than expected, redesigning primers to give fragment sizes within this range should be considered. 3. Mutations in GC-rich regions (which have a higher melt profile) are often difficult to detect. It may be necessary to re-design primers so that these regions lie towards the ends of the amplicon. 4. Sequences uploaded for the determination of melt profiles must be trimmed to fit the exact length of the PCR amplicon (including the primers with any clamps used). Re-optimisation must follow any primer redesign. 5. DNA isolated using organic extraction procedures, such as phenol/chloroform extraction, may require back extraction with isoamyl alcohol/chloroform before a final ethanol precipitation and wash. Extraction kits using chaotropic salts and/or spin column purification will require ethanol precipitation and wash. Failure to follow these recommendations can significantly shorten the life of the column.


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Fig. 8. The normal trace (shown as a dotted line) shows a small secondary peak marked (double dagger), due to the use of a non-proofreading PCR enzyme. However, the unknown sample (shown as a solid line) is clearly distinguishable from normal with an increase in intensity of this peak and the presence of a distinct 3rd peak.

6. Taq DNA Polymerase – as with all PCR-based mutation analysis approaches, use of a proofreading enzyme is recommended. The use of non-proofreading PCR enzymes may result in misincorporation rates that are detectable by dHPLC. This is generally seen as a secondary peak eluting immediately before the primary product (see Fig. 8). Although not recommended for use with dHPLC systems, the data obtained from such PCRs is generally adequate. However, as the relative strength of this “misincorporation peak” increases, the validity of the data will decrease. 7. Well-optimised PCR amplification is essential as non-specific amplification will render downstream dHPLC analysis uninterpretable. 8. Use of any of the following components in PCRs should be avoided, as they may adversely affect the life of the column: ●●

Bovine Serum Albumin (BSA).

●●

Mineral oil (23).

●●

Formamide.

●●

Proteinase K.

●●

Unidentified “proprietary” ingredients (enhancers, stabilisers, or additives).

9. Use of any of the following components in excess of the stated final (PCR) reaction concentrations should be avoided: ●●

●●

High-molecular-weight stabilisers such as polyethylene glycol (PEG) – 1% max. Detergents including, but not limited to: Triton X100, NP40, Tween 20, and SDS/SLS – 1% max.

●●

Glycerol – 2% max.

●●

DMSO – 10% max.

●●

Betaine – 1.25–2.5 M max.

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10. Although self-heteroduplexing will detect heterozygous variants, the paired approach will also pick up homozygous and hemizygous nucleotide changes. Unequal pooling of samples for heteroduplexing will reduce the detection of homo- and hemizygous mutations. “Self-heteroduplexing” will still occur, enabling the detection of heterozygous variants. 11. Example of a suitable program on a PTC-200 Thermal Cycler: 1 = 95˚C for 5 min 2 = 94.5˚C for 20s –0.5 C˚ per cycle 3 = goto 2, 100 times END The run time for this program is approximately 48 min. In this example, the final temperature is ~40°C, significantly below the annealing temperature of any amplicon (>100 bp), and sufficient for all amplicons to have become, once again, double-stranded. 12. Failure of heteroduplexing will result in lack of mutation detection, but cannot be visualised any other way. An inhouse produced positive control should be included in every heteroduplex and dHPLC run to validate this aspect of the protocol. A sample previously demonstrated to have a detectable sequence variant is PCR-amplified together with a normal control. The two samples are mixed and stored frozen in 6–8 ml aliquots until required. The use of commercially available mutation standards will highlight many problems but will not detect heteroduplexing issues. 13. The stability of heteroduplexes, once formed, is not well understood, so it is preferable for heteroduplex formation to be performed immediately prior to analysis. 14. It may be necessary to recalibrate the needle height if switching between tubes and plates. This is done via the autosampler control panel. 15. The selection of seal type is critical, to prevent the injection of extraneous material into the flow path. 16. To calculate usage: flow rate for a rapid DNA method = 1.5 ml/ min total (A + B) Assuming an average ratio of 50:50 A:B, 0.75 ml/min of each will be required. Run time on the WAVE HT is approximately 4 min per injection; therefore, each injection requires approximately 3 ml of each buffer. 17. Pressure limit errors given by the instrument may indicate a leak in the flow path (inability to reach minimum required pressure) or a blockage in the system (maximum pressure exceeded). The latter can usually be rectified by performing a


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hot wash with ACN. If this doesn’t resolve the problem, the in-line filter should be changed. If further maintenance is required to release a blockage, it may be necessary to isolate different sections of the flow path in turns and monitor the pressure during a 50% A: 50% B run at 56°C. This is done by sequentially disconnecting the column, preheat coil, in-line filter, injection valve ports, and finally the static mixer. The outflow at each of these stages should be collected into a beaker or tissue. Once the region of blockage is located, the appropriate component or length of PEEK tubing can be replaced. 18. The idle flow rate is 0.05 ml/min at a 50:50 ratio of buffer A and B. 24 ml of each will be required to maintain the flow path overnight (5 pm–9 am) and 96 ml of each for an entire weekend (5 pm Friday–9 am Monday). 19. Lack of peaks on a chromatogram is normally suggestive of a buffer error. This could be that buffers A and B are switched, or more commonly that their entry into the system is impeded by air in the flow path due to inadequate priming of the system, blocked inlet filters, or injection port leak. However, as the most common fault is that one or other buffer has run out, it is essential that buffer levels are checked before every run. 20. Loss of resolution, seen as a broadening of chromatogram peaks, is usually related to an overextended column or to contamination of the column or the sample. It can also be related to the use of “prohibited” additions to the PCR buffer. 21. Peak patterns are highly reproducible, although not necessarily unique, i.e. a specific base change will repeatably produce the same peak pattern, but it is possible that a different base change might produce an indistinguishable peak pattern. 22. Although it should be possible to overlay chromatograms from different runs, minor mobility shifts (of less than 0.1 min) are acceptable and reflect slight inter-batch buffer variations. 23. Chromatograms in which the peaks register a signal intensity of less than 1 mV should be rejected, and those between 1 and 2 mV interpreted with caution. Reduced peak height may reflect a sampling error (see troubleshooting section) or may be due to poor PCR amplification. 24. For ease, we recommend that a 75% ACN wash is performed at the end of every run regardless of size, with additional washes being included for runs of greater than 100 injections. 25. To perform a reverse hot wash, switch the pump off, remove the column from its housing and replace in the reverse orientation. Raise the oven temperature to 75°C and flush the column

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with 75% ACN at a medium flow rate (1.2 ml/min) for 30 min. After this period, switch the pump off and replace the column in its correct orientation. NB: exercise caution, as the column will be hot to touch. Return the oven to 56°C and run 50% A/50% B at a flow rate of 0.9 ml/min for 45 min to equilibrate the column.

Acknowledgment Some Figures are reproduced with kind permission from Transgenomic Inc. (Omaha, USA). References 1. Oefner PJ. (1995) Comparative DNA sequence by denaturing high performance liquid chromatography (DHPLC). Am J Hum Genet; 57: A266. 2. Oefner PJ, Underhill PA. (1999) DNA mutation detection using denaturing high-performance liquid chromatography (DHPLC). In: Dracopoli NC, Haines J, Korf BR, Morton C, Seidman CE, Seidman JG, Moir DT, Smith DR, editors Current Protocols in Human Genetics, Wiley, New York; pp. 7101–71012. 3. Underhill PA, Jin L, Lin AA, Mehdi SQ, Jenkins T, Vollrath D, Davis RW, CavalliSforza LL, Oefner PJ. (1997) Detection of numerous Y chromosome biallelic polymorphisms by denaturing high-performance liquid chromatography. Genome Res; 7: 996–1005. 4. Kuklin A, Munson K, Gjerde D, Haefele R, Taylor P. (1997) Detection of singlenucleotide polymorphisms with the WAVE DNA fragment analysis system. Genet Test; 1: 201–6. 5. Loyant V, Jaffre A, Breton J, Baldi I, Vital A, Chapon F, Dutoit S, Lecluse Y, Loiseau H, Lebailly P, Gauduchon P. (2005) Screening of TP53 mutations by DHPLC and sequencing in brain tumours from patients with an occupational exposure to pesticides or organic solvents. Mutagenesis; 20: 365–73. 6. Montagna G, Di Biase A, Cappa M, Melone MA, Piantadosi C, Colabianchi D, Patrono C, Attori L, Cannelli N, Cotrufo R, Salvati S, Santorelli FM. (2005) Identification of seven novel mutations in ABCD1 by a DHPLCbased assay in Italian patients with X-linked adrenoleukodystrophy. Hum Mutat; 25: 222. 7. Oldenburg J, Ivaskevicius V, Rost S, Fregin A, White K, Holinski-Feder E, Muller CR, Weber

8.

9.

10.

11.

12.

13.

14.

BH. (2001) Evaluation of DHPLC in the analysis of hemophilia A. J Biochem Biophys Methods; 47: 39–51. Ravnik-Glavac M, Atkinson A, Glavac D, Dean M. (2002) DHPLC screening of cystic fibrosis gene mutations. Hum Mutat; 19: 374–83. Santer R, Rischewski J, Block G, Kinner M, Wendel U, Schaub J, Schneppenheim R. (2000) Molecular analysis in glycogen storage disease 1 non-A: DHPLC detection of the highly prevalent exon 8 mutations of the G6PT1 gene in German patients. Hum Mutat; 16: 177. Su YN, Lee CN, Hung CC, Chen CA, Cheng WF, Tsao PN, Yu CL, Hsieh FJ. (2003) Rapid detection of beta-globin gene (HBB) mutations coupling heteroduplex and primerextension analysis by DHPLC. Hum Mutat; 22: 326–36. Takashima H, Boerkoel CF, Lupski JR. (2001) Screening for mutations in a genetically heterogeneous disorder: DHPLC versus DNA sequence for mutation detection in multiple genes causing Charcot-Marie-Tooth neuropathy. Genet Med; 3: 335–42. Wagner T, Stoppa-Lyonnet D, Fleischmann E, Muhr D, Pages S, Sandberg T, Caux V, Moeslinger R, Langbauer G, Borg A, Oefner P. (1999) Denaturing high-performance liquid chromatography detects reliably BRCA1 and BRCA2 mutations. Genomics; 62: 369–76. Wuyts W, Radersma R, Storm K, Vits L. (2005) An optimized DHPLC protocol for molecular testing of the EXT1 and EXT2 genes in hereditary multiple osteochondromas. Clin Genet; 68: 542–7. Xu E, Lai M, Lv B, Xing X, Huang Q, Ma Y, Wang W. (2005) DHPLC analysis of the


15.

16.

17.

18.

19.

matrix metalloproteinase-1 promoter 1G/2G polymorphism that can be easily used to screen large population. J Biochem Biophys Methods; 63: 222–7. Wolford JK, Blunt D, Ballecer C, Prochazka M. (2000) High-throughput SNP detection by using DNA pooling and denaturing high performance liquid chromatography (DHPLC). Hum Genet; 107: 483–7. Azarani A, Hecker KH. (2001) RNA analysis by ion-pair reversed-phase high performance liquid chromatography. Nucleic Acids Res; 29: E7. Fang N, Lin L, Ren J, Wu D. (2004) Detection of C677T mutation in methylenetetrahydrofolate reductase gene by denaturing high performance liquid chromatography. Biomed Chromatogr; 18: 625–9. Keller G, Hartmann A, Mueller J, Hofler H. (2001) Denaturing high pressure liquid chromatography (DHPLC) for the analysis of somatic p53 mutations. Lab Invest; 81: 1735–7. Betsalel OT, van de Kamp JM, MartinezMunoz C, Rosenberg EH, de Brouwer AP, Pouwels PJ, van der Knaap MS, Mancini GM,

20.

21.

22.

23.

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Jakobs C, Hamel BC, Salomons GS. (2008) Detection of low-level somatic and germline mosaicism by denaturing high-performance liquid chromatography in a EURO-MRX family with SLC6A8 deficiency. Neurogenetics; 9: 183–90. Jones AC, Sampson JR, Cheadle JP. (2001) Low level mosaicism detectable by DHPLC but not by direct sequencing. Hum Mutat; 17: 233–4. Lu C, Xu HM, Ren Q, Ao Y, Wang ZN, Ao X, Jiang L, Luo Y, Zhang X. (2003) Somatic mutation analysis of p53 and ST7 tumor suppressor genes in gastric carcinoma by DHPLC. World J Gastroenterol; 9: 2662–5. Rugg EL, Magee GJ. (2005) Use of denaturing high-performance liquid chromatography in molecular medicine. In: Walker JM, Rapley R, editors. Medical Biomethods Handbook, Springer, Totowa, NJ; pp 315–325. Harvey J, Schollen E. (2004). Practice Guidelines for the Use of the WAVE System in Diagnostic Service. CMGS: http://cmgsweb.shared.hosting.zen.co.uk/BPGs/ pdfs%20current%20bpgs/DHPLC.pdf.

Chapter 4 In Situ Detection of Human Papillomavirus DNA After PCR-Amplification Gerard J. Nuovo Abstract Human papillomavirus (HPV) is an essential cofactor for cancer at many sites, including the genital tract, oral cavity, conjunctiva, and periungual region. The in situ detection of HPV allows us to determine the cellular targets of the virus. In situ-based coexpression analyses of HPV with putative target proteins provide tremendous insight into the molecular evolution of these viral-associated cancers. HPV DNA is present in high copy number in the precancerous lesions and is, thus, readily detected by in situ hybridization. However, viral integration, typical during oncogenesis, is associated with reduced copy number of the virus, necessitating in situ polymerase chain reaction amplification for accurate in situ detection of HPV. This chapter provides the protocols that can be used to detect HPV DNA in situ as well as to correlate viral DNA with coexpression of relevant protein targets. Key words: HPV, In situ hybridization, PCR, Cervix, Coexpression, Immunohistochemistry

1. Introduction Sensitive molecular biology techniques, such as Southern blot hybridization and the polymerase chain reaction (PCR) have greatly enhanced our understanding of infectious disease processes and oncogenesis. However, an important limitation of these techniques is that the DNA or RNA extraction prior to analysis precludes histologic correlation. This drawback is overcome by in situ hybridization. However, one cannot reliably detect low copy numbers in a given cell with in situ hybridization (1–3). Thus, a negative result with in situ hybridization does not disprove the presence of the target. It is now possible to do PCR in intact cell and tissue preparations and, thus, to combine the extreme sensitivity of PCR with the cell localizing ability of in situ


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hybridization (1–5). Further, by performing in situ hybridization (or PCR in situ hybridization) and then, on the same tissue section, doing immunohistochemistry, one can determine whether a specific human papillomavirus (HPV) molecule is coexpressed with a putative cellular target (6). This chapter will describe methods for in situ hybridization and PCR in situ hybridization, as well as providing the foundation for in situ-based coexpression analyses.

2. Materials Whether doing in situ hybridization, PCR in situ hybridization, or immunohistochemistry, the same starting principle applies. The tissue section will possibly fall off during the procedure unless the slide is precoated with silane. Silane confers a positive charge on the glass slide and strongly attaches to the negatively charged molecules in the tissue. In my experience, this increases the tissue adherence rate of any in situ-based method to over 98%, as compared to about 40% if the slide is not precoated. The silane-coated microscopic glass slides were obtained from Fisher Scientific (e.g., catalog no. 07-200-008). I continue to use autoclavable plastic coverslips from BelArt (catalog no. S1370-14) to prevent the evaporation of the overlying solution and to be able to perform three different reactions per slide. Of course, the latter presupposes that the histotechnologist place three serial sections per glass slide which is simple to do for most surgical biopsies. The other basic principle with in situ hybridization or PCR in situ hybridization is that one must use tissues fixed in 10% neutral buffered formalin. This fixative cross links DNA and RNA to proteins and causes very few nicks in the DNA sequences. Fixatives which contain heavy metals such as mercury or picric acid (e.g., Bouin’s solution) will not allow for successful PCR in situ hybridization or in situ hybridization (7, 8). 2.1. In Situ and PCR In Situ Hybridization Reagents

1. The thermocyler was obtained from Perkin-Elmer Corporation. For most experiments, the DNA Thermal Cycler (catalog no. N801-0150) was used, although equivalent results were obtained with the 480 and 9600 models. 2. All of the reagents for PCR are included in the GeneAmp kit from Perkin-Elmer (catalog no. N801-0055). As previously published, I recommend the hot start maneuver, which can be done by simply withholding the Taq polymerase until the thermal cycler reaches 60°C (1). 3. The primers and probes for the detection of HPV DNA can be readily obtained from the medical literature (9).

In Situ Detection of Human Papillomavirus DNA After PCR-Amplification

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4. The HPV probes and biotin detection system may be obtained from a variety of sources; we use the HPV in situ detection kits from Enzo Life Sciences (catalog no. ENZ-32874). The detection system employs a streptavidin–alkaline phosphatase conjugate and the chromagen blue tetrazolium, which, in the presence of 5-bromo-4-chloro-3-indolynitrolphosphate, yields a purple-blue precipitate at the site of hybridization, is from Enzo Life Sciences (catalog no. ENZ-32700). The counterstain, nuclear fast red, which stains nuclei pale pink and does not stain cytoplasm, is also part of the Enzo Life Sciences HPV in situ kit. 5. Pepsin (from DAKO – catalog no. S3002) was employed although equivalent results were obtained with proteinase K (ONCOR; catalog no. S4508). I only use pepsin for in situ hybridization or PCR in situ hybridization, whereas I only use proteinase K for immunohistochemistry. 6. The formamide (catalog no. S4117), dextran sulfate (catalog no. S4010), and 20× SSC (catalog no. S2600) were obtained from ONCOR. However, they can be obtained as one reagent from Enzo Life Sciences (catalog no. ENZ-33808). This socalled in situ hybridization buffer can be used to dilute the probe for either in situ hybridization or PCR in situ hybridization. 2.2. Immunohistochemical Reagents

1. There are many commercial sources for primary antibodies that can be used for immunohistochemical reactions. The three companies that I use most in this regard are Ventana Medical Systems, Enzo Life Sciences, and ABCAM. Special attention needs to be placed on the animal used to generate the primary antibody. Since I use a detection system based on rabbit or mouse primary antibodies, if I am forced to use a goat or rat primary antibody I would have to add another step to the procedure after the primary antibody that would contain a rabbit-based anti-goat or anti-rat secondary antibody. 2. I use an automated immunohistochemical system. It is the Benchmark system from Ventana Medical Systems. This system does every step from removing the paraffin, to adding the primary and secondary antibodies, to detecting the antigen, and adding the counterstain. Indeed, the Benchmark system is capable of coverslipping the slide, with an optional add-on. The Benchmark system uses two different detection systems: Ultrasensitive Universal Fast Red and Ultrasensitive Universal DAB. In each case, the detection systems are based only on mouse and rabbit primary antibodies. Thus, if one is using a goat or rat primary antibody, he or she must manually add the rabbit anti-goat or anti-rat 30 min after the primary antibody has been added to the glass slide.

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3. Methods 3.1. In Situ Hybridization 3.1.1. Solutions

1. Protease solution: Add 1.3 mg of pepsin per ml of RNase free water which contains 0.1 N HCl. The solution can be kept at −20°C and thawed when ready to use. 2. In situ hybridization probe cocktail (for HPV genomic probes, although one may substitute other probes): Add 25 ml of formamide, 15 ml of 25% dextran sulfate, 5 ml of 20× SSC, and 5 ml of the probe (stock solution of 1 mg/ml of biotin-labeled probe). It is easier to add the 5 ml of the labeled probe to 45 ml of the in situ hybridization buffer. 3. Wash solution: The wash solution contains 2.5% bovine serum albumin and 0.2× SSC (or 30 mM sodium chloride) and is heated to 45°C. 4. Detection solution: The detection solution contains 0.1 M Tris–HCl (pH 9–9.5) and 0.1 M NaCl. 5. Substrate solution: The substrate solution is the detection solution to which is added the chromagen. For every 200 ml of the substrate solution, add 500 ml of nitro blue tetrazolium and 500 ml of 5-bromo-4-chloro-3-indolynitrolphosphate.

3.1.2. Steps

1. Place two or three (depending on the tissue size) 4-mm paraffin embedded sections on a silane-coated glass slide. 2. Wash slides in xylene for 5 min; wash slides in 100% ethanol for 5 min, then air dry. 3. Digest in pepsin solution for 20 min at room temperature. 4. Inactivate protease by washing in sterile RNase free water for 3 min. 5. Wash slides in 100% ethanol for 1 min, then air dry. 6. Add 5–10 ml of the probe cocktail to a given tissue section. 7. Overlay with plastic coverslip cut slightly larger than tissue section. 8. Place slide on hot plate, 95–100°C, for 5 min. 9. Remove bubbles over tissue gently with a toothpick. 10. Place slides in humidity chamber at 37°C for 2–15 h. 11. Remove coverslips – hold down one end with fingernail and lift off coverslip with toothpick. 12. Place in wash solution for 10 min at 40–60°C (higher temperature to determine HPV type or if background is a pro blem; lower temperature to increase detection rate of novel HPV types). 13. Wipe off excess wash solution and put slides in a humidity chamber – do not let slides dry out.


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14. Add 50–100 ml of streptavidin–alkaline phosphatase conjugate per tissue section. 15. Incubate in humidity chamber for 20 min at 37°C. 16. Wash slides at room temperature for 3 min in a detection solution. 17. Place slides in substrate solution. 18. Incubate slides for 30 min to 2 h, checking results periodically under microscope. 19. Counterstain with nuclear fast red and coverslip; view under microscope. 3.2. PCR In Situ Hybridization 3.2.1. Solutions

1. Amplifying solution: Add 3 ml of the GeneAmp buffer, 5.6 ml of MgCl2 (25 mM stock solution), 4.6 ml of dNTPs (stock prepared as per the GeneAmp kit), 1 ml of primer 1 and primer 2 (stock solution 20 mM), and 1 ml of bovine serum albumin (stock solution 2% w/v); add RNase free water to 29 ml. For three reactions, remove 6 ml and place in a separate tube. 2. Protease, probe cocktail, wash, detection, and substrate solutions: Same as above for in situ hybridization.

3.2.2. Steps

Follow steps 1–5 as above. 6. Add 10 ml of amplifying solution per tissue section. If possible, have three tissue sections per glass slide (one tissue no PCR – negative control, one tissue PCR with Taq polymerase – test, and the final tissue PCR with no Taq – additional negative control). 7. Overlay solution with plastic coverslip, anchor with nail polish; place slide in aluminum “boat” directly on thermal cycler (Fig. 1). 8. Time delay file – 82°C for 7 min. 9. At the onset of this file, add 1.0 ml of Taq to tube on ice which contains 6 ml of the amplifying solution (this would be for 30 ml total and for three separate reactions). 10. At 55°C, lift one edge of the coverslip gently and add 2 ml of the Taq solution per tissue section, overlay with preheated mineral oil. 11. Switch to time delay file – 94°C for 3 min. 12. Link this time delay file to a cycling file of 55°C – 2 min and 94°C – 1 min for 30–40 cycles; at conclusion link to soak file of 4°C; remove and ethanol washes, air dry. 13. Proceed to step 6 above (part A, standard in situ hybridization).

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Fig.1. PCR in situ hybridization done on the block of the DNA Thermal Cycler. The glass slides may be placed in an aluminum foil “boats.” A plastic coverslip is placed over the amplifying solution. The coverslip may then be anchored with a drop of nail polish and then overlaid with mineral oil during cycling. The boats and slides conduct the heat well and the boats will retain the mineral oil during the PCR phase of the technique. One may need to add mineral oil during the cycling process in case there is a small leak of the oil from the aluminum chamber.

3.3. Immunohistochemical Assay 3.3.1. Solutions

3.3.2. Steps

1. Antibody solution: Dilute the primary antibody in the diluent (catalog no. 251-018) from Ventana Medical Systems. 2. Detection system: As per manufacturer’s recommendations, use the Ultrasensitive Universal Fast Red or DAB detection systems from Ventana Medical System (catalog nos. 760-500 and 760-501). Since the steps are as outlined by the manufacturer, I will focus on the keys variables for successful immunohistochemical analysis using the Benchmark system. 1. Determine the optimal pretreatment condition: Using either a Medline search or the information provided by the manufacturer, determine which tissue should contain a high amount of the protein of interest. At the same time, determine the cell type(s) that likely will contain the high amount of the protein


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of interest. Then, take two silane-coated glass slides that have this tissue of interest (positive control). Make certain that each slide contains at least two tissue sections. For one slide, do nothing to one of the tissue sections and for the other section digest in the Protease 1 solution (catalog no. 760-2018) of Ventana for 4 min, then wash in tap water, dip in 100% ethanol, and air dry. Then do immunohistochemical testing on the two slides using cell conditioning #1 for the untreated slide and no additional pretreatment for the slide already subjected to either protease digestion or no pretreatment. 2. Determine the optimal concentration of the primary antibody: For each slide listed in step 1, use a dilution of 1:100. For another set of slides treated exactly the same way as in step 1, use a dilution of 1:500. 3. Score the results: Score each slide for signal (0–3) and background (0–3). Whichever concentration/pretreatment yields a three signal, zero background score is the optimal condition for that antibody.

4. Notes 1. The most common problems encountered with in situ hybridization are background and the absence of a signal. Background may be defined as the presence of a hybridization signal with a specific probe in areas of the tissue where the target should not be present (i.e., normal endocervical cells or in basal cells with HPV). Of course, in some instances one may not be sure where the in situ signal should localize. A more strict definition of background would be a hybridization signal when the labeled plasmid vector is employed (the plasmid is the vehicle used to clone the probe of interest). Background is the result of nonspecific binding of the probe to nontarget molecules. Two simple and logical ways to deal with background are to decrease the concentration of the probe and/or to increase the stringency of the posthybridization wash. If background is a problem, first try increasing the temperature of the wash by 10° increments. If background remains a problem at 55°C, then decrease the concentration of the probe tenfold. Another potential problem with in situ hybridization is the absence of a hybridization signal. I recommend that one test a vulvar condyloma for HPV 6/11, especially if they do not have much experience with in situ hybridization. The rationale is that most vulvar lesions that show the classic features of condyloma (perinuclear halos towards the surface of the lesion with concomitant nuclear atypia) will be strongly positive for

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Fig. 2. The usefulness of a positive control for in situ hybridization. Vulvar condylomas typically contain very high copy numbers of either HPV 6 or 11. Thus, the viral DNA should be readily detectable by in situ hybridization. The presence of a hybridization signal with the probe in most of the cells with the perinuclear halos (arrow ) typical of productive HPV infection demonstrates that the various conditions for in situ hybridization, notably protease digestion, fixation, hybridization, and detection, are adequate.

HPVs 6 or 11 (Fig. 2). If a signal cannot be generated, then the most common problems are (a) insufficient protease digestion; (b) tissue fixed in either a heavy metal or picric acid; (c) denaturing temperature had not reach 95°C; and (d) defective component of the detection system – usually NBT/BCIP. 2. The probe size for standard in situ hybridization is from 80 to 120 bp in size. However, one may want to use much smaller (20–40 bp) probes called oligoprobes. Oligoprobes are more readily available than the larger probes which require a cloned sequence of DNA. One only needs to know the sequence of the target of interest, readily available in the literature, to generate an oligoprobe. Also, one is obliged to use an oligoprobe internal to the sequence being amplified in PCR in situ hybridization in order to assure themselves that the signal is indeed the PCR product. Because oligoprobes are much shorter than a “standard” probe, there is a substantial reduction in the number of base pair matches and thus the strength of the hybridized complex compared with the larger probes. The practical consequence is that the wash conditions must be carefully chosen so as to minimize background but not to lose the signal. In practical terms, I have seen the signal lost for a 20mer oligoprobe with a posthybridization wash of 30 mM salt at 45°C; under these conditions


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the signal for a larger homologous probe would remain intact. Hence, I use different probe cocktail and posthybridization wash conditions for oligoprobes. Specifically, for the probe cocktail with an oligoprobe I decrease the formamide concentration to 10% and the salt in the wash solution is increased to 300 mM (2× SSC). A recent development has revolutionized the field of in situ hybridization. These are the locked nucleic acid (LNA) probes. These are made with nucleotides where the two and four positions are locked in three dimensions via a methylene bridge. This bridge prevents the associated probe from moving once attached to the target DNA sequence. Since the probe cannot move on the target DNA sequence, it is much more difficult to denature the complex, and the melting temperature of the annealed hybrid goes up considerably. Indeed, if one takes a 20mer and includes six LNA-modified nucleotides, the melting temperature will be over 30°C higher (10, 11)! Hence, with an LNA probe, one can use a 20mer and a posthybridization wash of 50°C and still have a signal after removing background. For this reason, despite their expense ($300–$400/probe), I strongly recommend that one use LNA probes for their in situ hybridization or PCR in situ hybridization work if the full-length genomic probe is not available. 3. Note that the Taq polymerase is not added to the tissue section until the temperature reaches 55°C. In any PCR reaction, several pathways compete with target-specific DNA synthesis. These include mispriming, DNA repair, and primer oligomerization. If the hot start modification is not employed, these “unwanted” pathways can overwhelm target-specific DNA synthesis such that a large amount of DNA that is mostly nonspecific is synthesized! This is not surprising when one considers that there is far more nontarget and primer DNA in a reaction mixture relative to target DNA. It has been shown that under nonhot start conditions that the detection threshold for the target of interest with standard PCR may be greater than several thousand copies, not the 1–100 copies most articles quote (1, 5). However, nonspecific DNA synthesis is greatly curtailed by the hot start modification. The end result is that one can reliably detect one copy per tissue DNA extract with the hot start modification using PCR. For PCR in situ hybridization, it has been demonstrated that the hot start modification readily allows for the routine detection of one target per cell using a single primer pair (1–5). 4. To demonstrate the utility of PCR in situ hybridization for the analysis of HPV in tissue sections, let us examine two specific scenarios: (a) The equivocal penile or vulvar biopsy: About 40% of papillary lesions of the vulva or penis that are

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clinically suggestive of an HPV-induced condyloma (i.e., low-grade SIL) lack the diagnostic features of this lesion on histologic examination (12, 13). Terms such as equivocal for condyloma, suggestive of condyloma, or borderline condyloma may be used for such lesions. Although these terms reflect the difficulty inherent in making the histologic diagnosis, they may be confusing to the clinician and, perhaps more importantly, to the patient who may be understandably confused whether he or she has a sexually transmitted disease. About 10–20% of these equivocal penile and vulvar lesions are HPV positive as determined by either Southern blot hybridization or PCR (12, 13). However, the histologic markers of HPV detection in such tissues cannot be ascertained as the DNA extraction prior to molecular analysis precludes histologic correlation. This problem is eliminated with PCR in situ hybridization. Typically, a weak signal is evident with standard in situ hybridization for HPV 6/11. A much more intense signal is evident in many more cells if in situ hybridization is preceded by PCR. Such analyses have shown that about 10–20% of equivocal tissues contain HPV DNA as demonstrated by PCR in situ hybridization and that a focally thickened granular layer in conjunction with epithelial cervices and para- or hyperkeratosis is the marker for HPV detection in such equivocal cases (12, 13). (b) The cervical squamous cell or adenocarcinoma: Although most cervical carcinomas contain HPV DNA and RNA, the detection rate varies considerably with the detection method. While the rate of HPV detection is greater than 90% with Southern blot hybridization or PCR, which have detection thresholds of about 1 virus per 100–10,000 cells, respectively, it is often about 30% with in situ hybridization, for which there must be ten viruses per cell for a positive result. Even when viral nucleic acids are detected in cervical cancers by in situ hybridization, most cancer cells appear to be HPV negative. It is unclear whether these HPV-negative carcinoma cells contain low copy numbers of the virus or indeed contain no viral DNA or RNA. By using PCR in situ hybridization, we have shown that the vast majority of cervical cancers do contain HPV DNA and that most of the cancer cells in a given lesion are also positive for the virus (4). Further, we have modified the procedure to detect RNA by using a reverse transcriptase (RT) step (4). 5. New horizons for HPV analyses: In my opinion, in situ-based coexpression of HPV and related proteins will yield substantial information that will allow us to better understand how to stop HPV infection and how to prevent the infection, once present, from progressing to cervical cancer. P16 is the classic protein


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Fig. 3. Colocalization of a protein (p16) and HPV-associated RNA. MicroRNAs are small regulatory RNAs involved in host defenses against viruses, as well as in oncogenesis. P16 is a protein strongly associated with HPV infection. In this CIN 1 lesion, we did colocalization of p16 with a microRNA (miRNA 125b), that is apparently much downregulated by HPV proliferation. In this mixing experiment, the Nuance software showed that p16 and the microRNA that is being downregulated by HPV are in mutually exclusive cells.

marker of HPV infection. We did a series of experiments (Nuovo et al., in press) whereby we showed that HPV strongly downregulated microRNA 125b expression. Given that observation, it follows that microRNA 125b and p16 should be present in distinct cell populations in CIN 1 lesions. We were able to demonstrate that after doing coexpression analyses of p16 and microRNA 125b in the same tissue section of a CIN 1 lesion (Fig. 3).

5. Pitfalls 1. If tissue morphology is poor, the most likely explanation is over protease digestion. Decrease the protease digestion time accordingly. 2. Always include an HPV positive tissue with each experiment. Vulvar or penile biopsies that show the granular cell changes (perinuclear halos and nuclear atypia) of a condyloma invariably are HPV positive, usually for HPVs 6 or 11 (Fig. 2). 3. Note that the optimal concentrations of the Mg2+ and Taq polymerase are greater than those for standard PCR. This may reflect difficulty in entry of these reagents to the site of

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DNA amplification and, in part, sequestration of the Mg2+ by cellular components. Consistent with this hypothesis is our observation that one may use tenfold less Taq polymerase with the addition of 1 mg/ml of bovine serum albumin (BSA) to the amplifying solution in PCR in situ hybridization; BSA can block absorption of the enzyme to the glass slide or plastic coverslip (1). 4. Add mineral oil as needed to prevent evaporation of the amplifying solution during the PCR step of PCR in situ hybridization (Fig. 1). References 1. Nuovo, G.J. (1992) PCR in situ hybridization. Protocols and applications. Raven Press, New York, NY. 2. Nuovo, G.J. (2002) Diagnosis of human papillomavirus using in situ hybridization and in situ polymerase chain reaction. Methods Mol Biol 179:113–136. 3. Bagasra, O., and Harris, T. (2006) Latest developments in in situ PCR. Methods Mol Biol 334:221–240. 4. Morrison, C., Catania, F., Wakely, P., and Nuovo, G.J. (2001) Highly keratinizing squamous cell cancer of the cervix: a rare, locally aggressive tumor not associated with human papillomavirus or squamous intraepithelial lesions. Am J Surg Pathol 25: 1310–1315. 5. Nuovo, G.J., Gallery, F., MacConnell, P., Becker, J., and Bloch, W. (1991) An improved technique for the detection of DNA by in situ hybridization after PCR-amplification. Am J Pathol 139:1239–1244. 6. Nuovo, G.J., Nana-Sinkam, P., Elton, T., Croce, C., Volinia, S., and Schmittgen, T.S. (2009) A methodology for the combined in situ analyses of the precursor and mature forms of microRNAs and correlation with their putative targets. Nat Protoc 4:107–115. 7. Greer, C.E., Peterson, S.L., Kiviat, N.B., and Manos, M.M. (1991) PCR amplification from paraffin-embedded tissues: effects of fixative and fixative times. Am J Clin Pathol 95: 117–124.

8. Nuovo, G.J., and Silverstein, S.J. (1988) Comparison of formalin, buffered formalin, and Bouin’s fixation on the detection of human papillomavirus DNA extracted from genital lesions. Lab Invest 59:720–724. 9. Zaravinos, A., Mammas, I.N., Sourvinos, G., and Spandidos, D.A. (2009) Molecular detection methods of human papillomavirus (HPV). Int J Biol Markers 24(4): 215–222. 10. Nuovo, G.J. (2008) In situ detection of precursor and mature microRNAs in paraffin embedded, formalin fixed tissues and cell preparations. Methods 44(1):39–46. 11. Godlewski, J., Nowicki, M.O., Bronisz, A., Nuovo, G., Palatini, J., De Lay, M., Van Brocklyn, J., Ostrowski, M.C., Chiocca, E.A., and Lawler, S.E. (2010) MicroRNA-451 regulates LKB1/AMPK signaling and allows adaptation to metabolic stress in glioma cells. Mol Cell 37(5):620–632. 12. Nuovo, G.J., Hochman, H., Eliezri, Y.D., Comite, S., Lastarria, D., and Silvers, D. (1990) Human papillomavirus DNA in penile lesions histologically negative for condylomata: analysis by in situ hybridization and the polymerase chain reaction. Am J Surg Pathol 14:829–836. 13. Nuovo, G.J., Becker, J., MacConnell, P., Margiotta, M., Comite, S., and Hochman, H. (1992) Histological distribution of PCRamplified HPV 6 and 11 DNA in penile lesions. Am J Surg Pathol 16:269–275.

Chapter 5 LATE-PCR and Allied Technologies: Real-Time Detection Strategies for Rapid, Reliable Diagnosis from Single Cells* Kenneth E. Pierce and Lawrence J. Wangh Abstract Accurate detection of gene sequences in single cells is the ultimate challenge of PCR sensitivity. Unfortunately, commonly used conventional and real-time PCR techniques are often too unreliable at that level to provide the accuracy needed for clinical diagnosis. Here we provide details of Linear-AfterThe-Exponential-PCR (LATE-PCR), a method similar to asymmetric PCR in the use of primers at different concentrations, but with novel design criteria to insure high efficiency and specificity. LATEPCR increases the signal strength and allele discrimination capability of oligonucleotide probes such as molecular beacons and reduces variability among replicate samples. The analysis of real-time kinetics of LATE-PCR signals provides a means for improving the accuracy of single-cell genetic diagnosis. Key words: Asymmetric PCR, Cell lysis, Fluorescent probes, Molecular beacons, PCR inhibitors, Primer and probe design, Primer melting temperature, Proteinase K, Quantitative PCR, Real-time PCR, Sample preparation for PCR

1. Introduction The polymerase chain reaction (PCR) provides a method for identifying alleles of specific genes or the mRNA transcribed from those genes. Through the 1980s and most of the 1990s, the products of PCR amplification were characterized using postamplification methods such as restriction enzyme treatment followed by electrophoresis through agarose or polyacrylamide gels. These and other postamplification detection strategies are time-consuming and increase the risk of contaminating subsequent assays. This is particularly problematic in the case of single-cell

*This chapter is revised from an earlier version published in “Methods Mol Med” (2007) vol. 132 pp. 65–85 Bimal D.M. Theophilus and Ralph Rapley (eds.), PCR Mutation Detection Protocols, Methods in Molecular Biology, vol. 688, DOI 10.1007/978-1-60761-947-5_5, © Springer Science+Business Media, LLC 2011

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samples, since a single product molecule inadvertently introduced into a sample tube can generate a false-positive result and result in a misdiagnosis. Real-time PCR using fluorescent probes (1–3) allows the kinetics of the amplification process to be observed and analyzed. Moreover, the fact that real-time PCR is carried out in closed tubes greatly reduces the risk of laboratory contamination, saves time, and is amenable to automation. Real-time assays using TaqMan™ probes have become popular for many applications, primarily due to the “assays on demand” program from Applied BioSystems for primer and probe design. However, the TaqMan™ assay requires digestion of the probe by the 5′ exonuclease activity of Taq polymerase, a process that requires probes with a relatively high melting temperature (Tm). This, in turn, makes it more difficult to distinguish allelic variants and can also reduce amplification efficiency. Molecular beacons and several other types of commercially available probes have greater allele discriminating capacities than TaqMan™ probes, but have design constraints of their own. Regardless of which type of probe is used to monitor a symmetric real-time amplification, hybridization of the probe to its target must compete with the reannealing of the complementary amplicon strands. By the end of the reaction, amplicon strand reannealing predominates and the probe only detects a fraction of the total number of amplicons produced (Fig. 1a). In order to circumvent this problem, we investigated the use of asymmetric PCR. Asymmetric PCR uses two primers of unequal concentration to first amplify both DNA strands exponentially until the limiting primer is depleted and then shifts to linear amplification

Symmetric PCR

LATE-PCR

Fig. 1. Schematic comparison of symmetric PCR and LATE-PCR for amplicon detection using molecular beacons. Near the completion of symmetric PCR (left panel) the complementary strands of the amplicon (black and gray lines with arrows representing the 3′ ends) reach high concentrations and reanneal. Molecular beacon molecules unable to hybridize with those targets remain in the nonfluorescent, hairpin configuration. LATE-PCR (right panel) generates an excess of the amplicon strand that is the target of the molecular beacon. Molecular beacons readily hybridize to those strands and emit fluorescence, generating a much greater total fluorescent signal from the LATE-PCR sample.

LATE-PCR and Allied Technologies

Fluorescence

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Fig. 2. Real-time PCR results for the detection of the DF508 allele in single, heterozygous lymphoblasts using molecular beacons. Symmetric PCR (a) replicate samples exhibit wide ranges of CT values (the point at which fluorescence reaches the dashed threshold line) and low final fluorescence. LATE-PCR (b) replicate samples have relatively low variation in CT values and much higher final fluorescence.

of one strand driven by the excess primer. The strand produced by the extension of the excess primer during the linear phase is freely accessible for hybridization to the probe (Fig. 1b). However, traditional asymmetric PCR that makes use of primers designed for symmetric amplifications (4) is inefficient, highly variable, and tends to generate high levels of nonspecific product. Those undesirable characteristics can be overcome if primers are designed for use at unequal concentrations. The resulting amplification strategy, termed Linear-After-The-Exponential-PCR (LATE-PCR), is efficient and specific (5, 6). A comparison of symmetric PCR and LATE-PCR for the detection of the DF508 allele of the cystic fibrosis gene (CFTR) in single cells is shown in Fig. 2. LATE-PCR also makes it possible to use lower temperature detection, since the probe does not need to compete with hybridization and extension of the limiting primer during the early, exponential phase of the reaction. Hybridization of probe and target is unimpeded once the limiting primer is depleted and can be done either by lowering the annealing step at that point or by introducing a low-temperature detection step between the extension and melting steps. Probes with lower melting temperatures are easier to design, more allele discriminating, and have lower background fluorescence. Moreover, because the probe dissociates from its target strand well below the extension temperature of the reaction, sufficient probe can be added to the reaction to measure all product strands without inhibiting the amplification reaction (5). We have used these features of LATE-PCR for constructing single-cell assays for several alleles of cystic fibrosis ((7) and unpublished), Tay–Sachs disease (5), B-thalassemia (8), and p53 (9). Here we provide practical information for the design and use of LATE-PCR assays.

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2. Materials 1. Cells with desired genotypes for positive controls. 2. Microscope of choice for cell analysis and transfer. 3. PCR enclosure hoods (e.g., Labconco Purifier™). 4. Low attachment culture dishes (e.g., Corning six well, cat. no. 3471). 5. Narrow bore cell transfer pipets and micromanipulators. 6. Mechanical pipetters and aerosol-resistant pipet tips. 7. Calcium-free, magnesium-free phosphate-buffered saline (PBS). 8. Nonacetylate bovine serum albumen (BSA) or polyvinylpyrrolidone (PVP) (optional). 9. Lysis solution containing 100 mg/ml proteinase K, 5 mM sodium dodecyl sulfate (SDS), 10 mM Tris –HCl, pH 8.3. 10. PCR primer design software. 11. Thermal Cycler with fluorescence detection capability (e.g., ABI PRISM® 7000 or 7700, Bio-Rad iCycler, Stratagene Mx3000P™ or Mx4000®, or Cepheid SmartCycler®). 12. Optical sample tubes appropriate to the thermal cycler. 13. Racks for placing sample tubes on ice (e.g., ABI MicroAmp® Bases). 14. Standard Thermal Cycler or heating blocks (with heated cover) for lysis reaction (optional). 15. PCR reagents: (a) Taq polymerase with hot-start capacity [either with antiTaq antibodies, e.g., Platinum Taq (Invitrogen) or modified enzyme, e.g., AmpliTaq Gold (ABI)]. (b) Buffers containing Tris and potassium chloride (usually supplied with commercial Taq polymerases). (c) Magnesium chloride stock solution at 25 or 50 mM. (d) Custom oligonucleotide primers and probes. (e) Deoxynucleotide triphosphates (dNTPs), PCR grade. (f) Water, molecular biology grade. (g) SYBR Green I (Molecular Probes) (optional).

3. Methods Obtaining reproducible results from samples of single cells requires: (1) sample preparation that avoids inhibitors of PCR and removes chromosomal proteins from the DNA; (2) the use


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of primers and probes that maximize amplification efficiency, specificity, and signal strength; (3) analysis of real-time signal kinetics from tested samples and controls with known genotypes. 3.1. Preparing and Lysing the Cell

The choice of methods for isolating single cells will vary considerably depending on cell type and available equipment. For instance, cells in suspension can be isolated individually by hand-controlled micromanipulation or by fluorescence-activated cell sorting. Alternatively, fixed or embedded cells can be isolated using laser capture microscopy, although the required equipment is expensive and not widely available. This chapter provides only general information on this topic with the intension of pointing out potential pitfalls that can affect cell lysis, genomic DNA preparation, and subsequent PCR.

3.1.1. Isolating and Washing the Cell

When cell isolation is carried out manually, cells should first be diluted to a density that facilitates picking up individual cells using either a hand held pipette or a pipet in a micromanipulator. Adherent cells should be dissociated by repeated pipetting, preferably in a calcium-free, magnesium-free media or PBS. Petri dishes or microtiter plates with low-adhesion surfaces can reduce the chances for cell damage or loss. Solution additives such as BSA or PVP can also be used for this purpose, but any additive should be carefully evaluated for its effect on cell lysis and amplification (see Note 1). Several components present in culture media or used in cell isolation techniques can inhibit PCR and must be removed by transferring the cell through PBS or culture media that lacks the inhibitors (see Note 2). One or two rinses may be sufficient if the transferred volume can be kept to a minimum (e.g., overall volume dilution of 1:100 or greater per step). Transfers should be practiced before attempting to manipulate valuable, limitedsource cells. First, aspirate a small amount of the wash solution into the transfer pipet, then aspirate the cell into the tip of the pipet. Carefully expel the contents of the pipet into the wash while examining under the microscope. As soon as the cell exits the pipet, remove the pipet from the wash dish, expel the remaining solution into a separate container, and rinse the pipet in unused wash solution. Repeat this procedure as necessary to reduce the concentration of potential PCR inhibitors. All washes should be brief.

3.1.2. Preparing Lysis Solution and Transferring the Cell

Using real-time detection of multicopy genes, we demonstrated that a properly buffered solution containing proteinase K and SDS provides the greatest number of targets for amplification (8). This lysis reagent can be prepared in advance and stored up to at least 1 year at −20°C in a constant temperature freezer (i.e., not frost free). Other tested lysis methods resulted in more variable recovery and/or delayed detection, presumably due to either

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DNA damage or incomplete removal of chromosomal proteins from the DNA. A delay in PCR signals can also indicate inefficient amplification due to the presence of PCR inhibitors. Shortly before preparing the cells, an aliquot of the lysis reagent is thawed on ice and 10 ml pipetted to each PCR sample tube (see Note 3). It is extremely important to use procedures that minimize the likelihood of contamination when preparing or working with lysis solution and PCR reagents (see Note 4). Sample tubes should be kept on ice until cells have been transferred, as proteinase K is self digesting at rates that are temperature dependent. Any unused solution should be discarded, as repeated freezing and thawing may reduce enzyme activity. The final cell transfer is done directly into lysis solution in a PCR tube (see Note 5), keeping the volume of the transferred wash solution to a minimum. Depending on the type of sample tube, it may not be possible to observe the cell during this transfer. Careful observation of the fluid height in a fine-bore pipet is usually sufficient to insure the transfer of the cell and avoid adding excessive volume of wash solution. The sample tube should be centrifuged briefly (a few seconds) to insure all liquid is at the bottom of the tube and returned to ice until the completion of all cell transfers. 3.1.3. Lysis Incubation

The lysis incubation should be carried out in a temperature- controlled block or thermal cycler separate from that used for amplification (see Note 6). Incubate samples at 50°C for 30 min, then 95°C for 15 min. It may be possible to shorten the 50°C incubation depending on cell type. The high-temperature incubation is required to completely inactivate proteinase K. A heated cover must be in place over the samples to prevent condensation. If condensation is present on the cap or sides of the tubes following this incubation, subsequent amplification efficiency may be reduced. Reaction tubes can be frozen at −20°C (constant temperature freezer) or placed on ice for immediate use.

3.2. LATE-PCR

There are three main criteria for LATE-PCR design. First, the concentration-adjusted melting temperature of the limiting primer ( Tm L ) at the start of the reaction must be at least as high as that of the excess primer. This is achieved by either making the limiting primer longer or higher in percent guanine and cytosine (G + C) relative to the excess primer. Second, the concentration-adjusted temperature of the excess primer ( Tm X ) must be reasonably close to the melting temperature of the double-stranded amplicon ( Tm A ) in order for that primer to successfully compete with the accumulating single-stranded product for hybridization to the target strand. Third, if real-time detection is utilized, the concentration of the limiting primer should be chosen such that the limiting primer is depleted approximately when the probe signal reaches the detection threshold, i.e., at the C T value of the reaction.

LATE-PCR and Allied Technologies 3.2.1. Designing Limiting and Excess Primers

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Primers originally designed for symmetric PCR can be modified to fit LATE-PCR criteria (usually by lengthening the primer chosen as limiting), or primers can be newly selected according to those criteria. In either case, primer design software should be used to evaluate internal stability characteristics and 3¢ dimer formation in the same manner as would be done for symmetric PCR primers. Computer software can also be helpful in selecting primers. Preferred primer software provides input for primer concentration and should calculate Tm according to nearest neighbor methods using accurate thermodynamic values (11–13). Do not rely on primer Tm calculations based on the earlier estimates of nearest neighbor thermodynamic values by Breslauer (14). When selecting new primers for LATE-PCR, it is useful to scan the sequences neighboring the site to be probed (e.g., mutation or polymorphism) for a region with relatively high GC content. The initial choice of a limiting primer can be made from that region. It generally does not matter which DNA strand is chosen for the sequence of the limiting primer, as long as the hybridization probe is later chosen from a sequence on the same strand. An initial evaluation of the region to be amplified also can provide an estimate of Tm A , which will be needed to determine the required Tm X value. The concentration of limiting primers should be about 50 nM (1.25 pmol per 25 ml reaction) when used in combination with molecular beacons labeled with FAM or TET. At that concentration, limiting primer length of approximately 24–32 nucleotides is needed to achieve Tm L in the vicinity of 65°C (see Note 7). Excess primer concentration is usually 1 or 2 mM. Optimal amplification efficiency and specificity are achieved with Tm X about 5° below Tm L , when the primer concentration ratio is in this 20:1 to 40:1 range (6). Primer Tm calculations are made using the nearest neighbor formula (15): Tm =

∆H + 12.5 log [M ] − 273.15. ∆ S + R ln (C / 2)

The thermodynamic values DH and DS are calculated according to Allawi and SantaLucia (11). R is the universal gas constant and C is the initial concentration of the primer. The salt correction is that of SantaLucia et al. (16) using [M] as the total molar concentration of the monovalent cations, sodium and potassium in the PCR buffer. The Tm calculations can be made using the MELTING program available on the Internet site http://mobyle.pasteur.fr/ cgi-bin/portal.py. Another consideration in designing primers for LATE-PCR is Tm A . That value depends primarily on amplicon length and GC content. Short amplicons (around 100 nucleotides) are preferred for gene expression analysis or diagnosis of a specific genetic allele.

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When multiple alleles are tested or sequencing information is desired, longer regions can be successfully amplified. We have been able to amplify a 611 nucleotide segment of the p53 gene using LATE-PCR criteria and use samples directly for sequencing (9). Even for short amplicons, Tm A does not vary significantly with concentration, as the helix growth steps dominate the helix initiation step, producing a pseudo first-order equilibrium for which no concentration effect is observed (14). Therefore, good estimates of Tm A are obtained using a “%GC” formula (17):

Tm A = 81.5 + 16.6 log

[M] + 0.41(%G + %C) − 500 / length. 1 + 0.7[M]

The formulas do not include a factor for magnesium concentration, which can raise the actual Tm several degrees, but still provide valuable comparisons for designing amplification reactions. As mentioned above, Tm X must be reasonably close to Tm A in order for that primer to successfully compete with the accumulating single-stranded product for hybridization to the target strand. We have observed the strongest real-time detection signals when Tm A − Tm X is about 10–15°C, as calculated using these formulas (6). Signal strength was lower as that value increased and was unacceptably low when it exceeded 20°C. Therefore, primers must have higher Tm for amplicons that are long or GC rich. 3.2.2. Probe Design

We describe the design of molecular beacons for LATE-PCR, although it should be recognized that many other types of oligonucleotide probes can be used with this amplification technique (see Note 8). Molecular beacons are fluorescently labeled oligonucleotides that assume a stem-loop structure in the absence of homologous target, bringing a fluorophore on the 5¢ end of the molecule into close proximity of a quenching moiety (e.g., DABCYL) on the 3′ end (Fig. 1) (1). The molecular beacon is able to hybridize with a DNA strand (such as a PCR product) with sequence homologous to its loop. In that configuration, the fluorophore emits its fluorescent signal when illuminated at parti cular wavelengths. Thus, increasing PCR product in the presence of the homologous molecular beacon generates corresponding increases in fluorescent signal (Fig. 2). Multiple targets can be monitored in the same reaction by labeling different molecular beacon sequences with different fluorophores. LATE-PCR makes it possible to use molecular beacons with shortened loop sequences and greater allele discrimination. The sequence of the molecular beacon loop (or any other probe that fluoresces upon hybridization) must be chosen from the same DNA strand as the limiting primer. If the probe is used for distinguishing a single-nucleotide polymorphism (SNP), that site should be in the center third of the loop. The Tm of the beacon


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loop sequence to its target ( Tm P ) should be at least 5°, and preferably about 10° below Tm L . The lower Tm P value insures that the probe will not interfere with extension of the limiting primer. Thus, amplification efficiency during the exponential phase of LATE-PCR remains high even in the presence of high concentrations of probe. This contrasts to the situation in symmetric PCR, where Tm P must be greater than the Tm of the primers. Good estimates of Tm P can be obtained using the same nearest neighbor formula used for determining Tm L and Tm X, even though variations in the molecular beacon stem do affect the empirically determined Tm of the beacon-target hybrid. Stem designs are similar to conventional design, typically 5–6 bp, predominantly G and C. Estimations of the stem Tm are made using the intramolecular hybridization program mfold (18), available online at http://www.bioinfo.rpi.edu/applications/mfold/. That program is also useful in identifying sequences that can form stable nonhairpin structures. Unlike conventional designs for molecular beacons in which Tm P and stem Tm are both typically 7–10°C above the annealing temperature, we prefer to increase the stem Tm 5–10°C above Tm P in order to assure lower background fluorescence at the annealing temperature. Molecular beacons should be tested using synthetic target oligonucleotides prior to use in LATE-PCR. The complementary oligonucleotide target should include at least three nucleotides beyond each end of the molecular beacon loop, using the sequence of the target gene, so that possible interaction between the stem and target is included in the empirically determined Tm (see Note 9). A melting analysis of molecular beacon in the absence of target, with complementary target, and with mismatched target (in the case of SNP analysis), is carried out to determine the best temperature for allele-specific detection (Fig. 3). Molecular beacon is used at the concentration that will be present during LATEPCR, typically 0.5–1 mM, and the concentration of targets should be about 0.5 mM, the estimated final concentration of singlestranded product following LATE-PCR. Sodium, potassium, and magnesium concentrations should be the same as those used for amplification. Additional details for molecular beacon design, synthesis, and testing are available in Marras et al. (19) and on the internet site http://www.molecular-beacons.org. 3.2.3. Components of LATE-PCR

With the exception of the primer and probe concentrations, other components used in LATE-PCR samples are similar to those used in symmetric reactions for single cells. The use of a “hot-start” method to prevent mispriming prior to the first denaturation step is required. Several commercially available Taq polymerases are modified so that they become active only after the initial hightemperature incubation. We prefer to use Taq polymerase with antibodies, as the required denaturation step is usually shorter.

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Relative Fluorescence

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Taq polymerases from different commercial sources are supplied with buffers containing sodium Tris (or other buffer) and potassium chloride. Begin testing using the recommended buffer solution, keeping in mind that varying the concentration of the monovalent cations will affect primer and probe Tm. The dNTPs (specifically dATP, dCTP, dGTP, and dTTP) should be PCR grade and included at about 0.2 mM each. Higher concentrations may be needed for multiplex reactions and lower concentrations are useful if the single-stranded product will be used directly for sequencing. Remember that dNTPs chelate magnesium ions and thereby affect the free magnesium concentration in the sample. Therefore, changes to dNTP concentration may affect reaction efficiency and specificity. We generally use a magnesium concentration of 3 mM. That concentration works well with most Taq polymerase enzymes and molecular beacon probes. 3.2.4. LATE-PCR Cycling Parameters 3.2.4.1. Initial Cycling Steps and Duration

An initial denaturation step of 95°C for 2 min is followed by 25–35 initial cycles with steps for primer annealing, primer extension, and product denaturation. Fluorescence detection is not needed during these cycles. The annealing step should be no more than 10–15 s. Longer incubations promote nonspecific amplification. The extension step is usually carried out at 72°C, at which Taq polymerase has maximal activity. If amplicon size is only 100–200 nucleotides, 15 s is more than sufficient to complete primer extension. A denaturation step of 5 s at 95°C should separate the DNA strands of most amplicons, enabling hybridization with primers during the subsequent annealing step.


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Testing replicate samples containing low, equal concentrations of genomic DNA (e.g., 10 or 100 genome equivalents) at three or four different annealing temperatures is usually sufficient to identify optimal conditions (see Note 10). That optimum is usually close to the calculated Tm X value in samples containing 3 mM magnesium and 50 mM potassium. Amplification efficiency at different annealing temperatures is evaluated by comparing mean CT values of replicate samples. Lower CT values (earlier detection) indicate higher efficiency. Reaction specificity can be evaluated by analyzing products using gel electrophoresis. Alternatively, the DNA-binding dye SYBR Green I can be substituted for the hybridization probe and product melting analysis can be done following amplification (20) (see Notes 11 and 12). One of the difficulties of traditional asymmetric PCR is lowamplification efficiency, which for real-time reactions causes delays in detection and the inability to obtain quantitative information. Another problem encountered using traditional asymmetric PCR is the high level of nonspecific amplification, which can reduce the yield of specific product and the resulting signals from hybridization probes. By designing primers for which Tm L is higher than Tm X , LATE-PCR makes it possible to use annealing temperatures that are low enough to insure high amplification efficiency by the limiting primer, yet high enough to minimize mispriming by the excess primer. 3.2.4.2. Fluorescence Detection During Linear Amplification

Molecular beacon signals for single copy targets usually reach detection threshold around cycle 40–45, depending on the detection equipment and the specific molecular beacon. Detection threshold will be reached about 3.2 cycles earlier for each tenfold increase in the initial target concentration. Fluorescence detection should be included during cycling starting about ten cycles before reaching threshold and those initial values are then used to determine a fluorescence baseline for subsequent readings (see below). Detection can either be carried out during the annealing step of a standard thermal cycle or, preferably, during a step added after extension, since most of the amplicon strand detected by the probe remains single stranded during the linear phase of LATEPCR. The temperature at which detection is done is chosen based on the tests with the probe and synthetic targets (see Subheading 3.2.2 and Fig. 3). That temperature should be low enough to provide strong signal from the complementary targets, but high enough to avoid signal from mismatched targets. Dropping the temperature a few degrees below the optimal annealing temperature at this point of the reaction usually does not present a problem in terms of nonspecific amplification. However, large drops in temperature should be avoided, as mispriming by amplicon strands may produce a phenomenon we refer to as “product evolution” (see Note 12).

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The CT value of the reaction should be reached at or slightly before the limiting primer is depleted. Under these circumstances, the observed CT value will reflect the number of copies of the target sequence present in the sample at the start of the reaction, as in the case of symmetric real-time PCR. It may be necessary to test higher concentrations of the limiting primer (e.g., 100 or 200 nM), if the CT values are higher than anticipated or the subsequent rate of fluorescence increase is low. Conversely, nonlinear fluorescence increase may reflect limiting primer concentrations that are too high. Remember that altering the concentration of the primers will change their Tm. 3.2.5. Preparing and Running Diagnostic Assays on Lysed Cells

Large volumes of solutions containing all PCR reagents except Taq polymerase can be prepared and stored frozen in aliquots sufficient for concurrently tested samples, including positive and negative controls. Using the same mixture provides the highest reproducibility between assays run on different days. Taq polymerase should be added to the thawed aliquot just before use. The reagent solution should be thoroughly mixed before and after being added to individual samples containing lysed cells, but it is advisable not to vortex in order to avoid shearing genomic DNA, which increases mispriming. A final volume of 25 ml containing 1.25 U of Taq Polymerase is used for monoplex reactions, while concentrations up to 2.5 U are recommended for multiplex reactions. Samples should be kept on ice to insure minimal polymerase activity during preparation. Even in the presence of antibodies, some mispriming may occur if samples are kept at room temperature for long periods prior to PCR. The thermal cycler is programmed with the optimal cycling parameters with the detection step included about ten cycles before the anticipated CT values. In most cases with single copy genes, this will mean that the detection step will be added after the first 30 cycles. We typically run a total of 60 cycles. Specific requirements for selecting sample wells, programs, and detection wavelengths will vary with different thermal cyclers.

3.3. Analysis of Real-Time Signals

In order to properly display and analyze real-time signals, a fluorescence baseline is set using readings in the cycles before amplicon detection. Using a baseline normalizes background variations and gradual increases in fluorescence unrelated to the amplicon. The baseline readings can include any or all of the cycles before fluorescence increase. At least five cycles are usually necessary. Baselines are determined separately for each fluorophore used. A threshold of five or ten standard deviations above baseline detection values is typically suggested by thermal cycler manufacturers. We have found that choosing a threshold with a specific fluorescent value often provides better reproducibility between assays run at different times using the same PCR reagent mixture.

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One of the advantages of real-time PCR is the ability to identify samples with atypical signal kinetics. Assay accuracy can be increased by excluding such samples from diagnosis (21, 22). LATE-PCR increases final fluorescence intensity and reduces sample-to-sample variation, thereby improving the kinetic analysis. In order to establish limits for diagnosis, cells with known genotypes should be tested at the same time or at least using the same PCR mixture as unknown samples. Each genotype should be represented by about ten or more samples. Mean values for CT, fluorescence increase (slope), and final fluorescence are determined for samples with the same genotype. Individual sample values are evaluated using the extreme studentized deviate method to identify outliers (23). Any positive control sample that does not yield the expected signals, or yields a value that is a statistical outlier is not used for establishing diagnostic limits. We typically set those limits at three standard deviations from the means. Figure 4, illustrates this method for final fluorescence values obtained from human lymphoblasts homozygous or heterozygous for the DF508 mutation in the cystic fibrosis gene (CFTR) or homozygous normal at that location. An initial assay was run to establish diagnostic limits, including the limits for final fluorescence indicated by the dashed lines for homozygous normal cells (box 1), homozygous mutant cells (box 2), or heterozygous cells (box 3). Since LATE-PCR yields a narrower range of final 180

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Fig. 4. Scatter plot of LATE-PCR final fluorescence values in replicate samples of individual lymphoblasts homozygous normal for CFTR (filled circles), homozygous for the DF508 mutation (filled squares), or heterozygous for the DF508 mutation (open triangles). The boxes labeled 1, 2, and 3 indicate diagnostic limits for those genotypes that were established by previously tested samples. The accuracy of the assay is improved by excluding from diagnosis all samples outside these limits. See text for details.

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uorescence values compared to other real-time methods, the fl size of these boxes is relatively small and gives a useful first step for data analysis. Individual data points shown include all results from a second assay using the same PCR reagent mixture, simulating “unknown” sample testing. The large majority of data points fall within the boxes established for the corresponding genotypes. A few samples did not give the expected results. Two samples with values outside the boxes might have been misdiagnosed, but are excluded based solely on quantitative analysis of final fluorescence. First, one heterozygous cell yielded an extremely low fluorescence value for the DF508 allele (open diamond near the upper right corner of box 1). Such a preferential amplification result would have almost certainly been misdiagnosed as homozygous normal using conventional PCR and electrophoresis, but the molecular beacon provides the sensitivity to detect the mutant allele. Second, a single homozygous mutant sample generated a low-level signal for the normal allele, presumably due to contamination (solid square to the left of box 3). That signal, however, was outside the limits for final fluorescence and well outside the limits for CT value (latter not shown on Fig. 4.) and therefore would not be misdiagnosed as heterozygous. Evaluating CT values and rates of fluorescence increase (slope) provides additional means for reducing misdiagnosis. Figure 5, plots those values for the samples that yielded only signals from the normal allele, i.e., those indicated by the data points in box 1 of Fig. 4, including two heterozygous cell samples that failed to 10

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CT Value Fig. 5. Scatter plot of LATE-PCR CT values and rates of fluorescence increase for samples in box 1 of Fig. 4 (i.e., those generating only normal allele signal). Assay accuracy can be increased using the diagnostic limits indicated by the broken lines. A similar analysis can be done for samples generating only DF508 allele signals (box 2 of Fig. 4) and for samples generating both signals (box 3 of Fig. 4). See text for additional details.


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generate a DF508 allele signal. The diagnostic limits for CT values and rates of fluorescence increase were established using data from the prior assay, as described in the preceding paragraph. One of the heterozygous cell samples generated a CT value above those limits for diagnosis as homozygous normal. The other heterozygous cell sample gave values within the limits, although the slope was higher than that from any of the homozygous cells. In the setting of preimplantation genetic diagnosis (PGD) where both parents carry the same mutant allele, failure to identify allele dropout (ADO) of the DF508 allele has no phenotypic consequence, but becomes extremely important as the assay is extended to multiple mutation sites within a gene (see Subheading 3.4). Analysis of the real-time signals using the three standard deviation limits reduces misdiagnosis of ADO by about half for both symmetric PCR and LATE-PCR (7, 22) and unpublished data). 3.4. Assays for Detecting Compound Heterozygotes

Genotyping multiple mutation or SNP sites within a gene requires either a single amplicon that encompasses both sites or coamplification of the two regions using separate pairs of limiting and excess primers. If possible, the single amplicon approach is preferred, since it is simpler to design and optimize, and provides a means to detect ADO in PGD cases. (Absence of the normal allele signal at either site is an indication of ADO.) In contrast to symmetric real-time assays where signal strength declines rapidly as amplicon size increases, LATE-PCR can generate strong signals with amplicons several hundred nucleotides long. Although we have not tested the limits in this area, strong signals have been obtained for a 650 nucleotide amplicon. Limits may depend more on Tm A values for reasons discussed in Subheading 3.2, and thus be longer when the %GC of the amplicon is low and relatively short when %GC is high. In cases where the distance between the mutation sites is too great for single amplicon design, the individual sites can be coamplified using LATE-PCR design criteria for each. Primers need to be evaluated for possible 3′ dimer formation, as would be the case for any multiplex PCR, paying particular attention to possible interaction between the two excess primers. Optimizing PCR reagent concentrations and the annealing temperature enables coamplification of both targets to similar levels. Moderate increases the concentrations of one pair of primers can be used to equalize amplification efficiencies. We have successfully coamplified CFTR exon 10 and exon 11 sequences using LATE-PCR (unpublished data). ADO has greater consequences when multiplex amplication is used in PGD to identify compound heterozygotes, because a failure to amplify either of the mutant alleles can lead to misdiagnosis and transfer of an affected embryo. Several PGD centers have implemented tests that include amplification of polymorphic

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sites found near the tested gene (24). This approach, however, only detects loss of the entire region of the gene (e.g., due to aneuploidy) or poor accessibility of the DNA in that entire region due to inadequate cell lysis. Those situations are also detected by the absence of one of the normal allele sequences when testing two sites within the gene. When the target gene copies are both present, amplification from different sites within or neighboring the gene are independent events and therefore coamplification of neighboring polymorphic sites has limited value. In contrast, the analysis of real-time kinetics provides a means of identifying samples that exhibit atypical amplification including ADO. The strong signals and reliability of LATE-PCR offer the best opportunity to increase the diagnostic accuracy of single-cell PCR.

4. Notes 1. PVP at 0.01 mg/ml in the final cell wash step in a cystic fibrosis assay does not delay detection (21). Preliminary tests did show that detection was delayed slightly if higher concentrations of that solution were added to the lysis solution. Much larger and more variable detection delay was found with polyvinyl alcohol (PVA), possibly due to interference with fluorescence detection. 2. We have observed that calcium can inhibit amplification. Serum and other culture additives such as hemoglobin, immunoglobulin, and heparin also interfere with amplification (25, 26). Adding bovine serum album (nonacetylated, nuclease-free) can improve PCR efficiency in the presence of some of these inhibitors (26). 3. The volume of the lysis solution can be adjusted for the specific application and final PCR volume. Volumes below 10 ml can be used if the volume of transferred PBS is less than about 10% of that volume. Higher volumes are only limited by the volume of subsequently added PCR reagents and the final PCR volume. 4. A cell inadvertently introduced at any step can provide a DNA template for amplification. Contamination control measures should include dedicated pipetters for solution preparation, aerosol-resistant pipet tips, lab coats, disposable caps, masks, high-cuff gloves, and containment hoods, all in rooms separate from the PCR amplification area. While UV treatment offers some protection from contaminating DNA, its effectiveness is limited. Treating surfaces with 10% bleach (1% sodium hypochlorite) is more effective for eliminating contaminating cells and DNA. The work area should be a “DNA-free zone”


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that resembles a “sterile field” in an operating room. Only clean gloves should touch items within that zone (sample tubes, pipetters, pipet tip boxes, reagent vials, etc.) and gloves that come in contact with any area outside the zone should be changed immediately. These procedures should be used when handling sample tubes at any step prior to PCR. 5. We have found that proteinase K-based lysis is not as effective in PCR tubes with glass-like properties, such as those for the Cepheid Smart Cycler, presumably due to adhesive properties of those tubes. Doing the lysis step in a standard sample tube, then adding PCR reagents and transferring to the required PCR tube is an alternative, but losing material during that transfer increases the possibility of ADO. Preliminary results indicate that alkaline lysis without dithiothreitol (DTT) can produce acceptable results in the Smart Cycler tubes. DTT was present in the initial protocol for reducing protamines in sperm (27), but is unnecessary for most cell types and residual DTT reduces PCR efficiency (10). 6. Contamination of a thermal cycler block with PCR product is nearly impossible to avoid, even when great care is taken not to open tubes following PCR. If the same thermal cycle is used in a subsequent assay for the lysis incubation, it is possible to introduce those product molecules into samples when the tubes are opened to add PCR reagents. If the same block must be used for both lysis and PCR, it must be decontaminated between each assay. The block should be flooded with 10% bleach, and then rinsed thoroughly with water. 7. If an adequate limiting primer cannot be designed using the actual DNA sequence of the target gene, Tm L can be increased by substituting one or two guanine bases for adenine near the 5′ end of the primer. Hybridization of that primer with the initial target will have low affinity G-T pairing, but not destabilizing mismatches, and subsequent hybridization with complementary amplicon strands will provide high amplification efficiency during the exponential phase of LATE-PCR. Since the annealing temperature during the initial cycles cannot be lowered without risking mispriming by the excess primer, this option has obvious limits, particularly with a low initial target number, and therefore the Tm of the limiting primer with the initial target sequence should not be more that 5°C below Tm X . 8. We have used double-stranded displacement probes (28, 29) for some allele-specific assays. Those probes are easy to design and are relatively inexpensive, since each oligonucleotide is modified with a single fluorophore or quencher, not both. Extensive purification is not necessary, greatly increasing the manufacturing yield relative to dual-labeled probes such as molecular beacons. More recently, we have designed

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short-stem” molecular beacons that form a hairpin of only “ 2 or 3 bp and use a Black Hole Quencher® instead of DABCYL. The Black Hole Quencher® provides a probe with low background fluorescence over a wide range of temperatures, yet generates a strong fluorescent signal upon hybridization. Short-stem beacons are mismatch tolerant and multiple variants can be distinguished through melting analysis. In general, the LATE-PCR benefits of increased signal strength can be accrued using any probe that signals upon hybridization. Although TaqMan™ probes could be designed to work with LATE-PCR amplification, the need for hydrolysis requires that those probes have high melting temperatures and hybridize with the extension products of the limiting primer, rather than the accumulating single-stranded product. Therefore, benefits with TaqMan™ probes are limited. 9. When designing the molecular beacon stem, it is worthwhile to check for complementarities with nucleotides in the target sequence. It is usually possible to modify the stem slightly to minimize hybridization between the stem and target. Alternatively, those hybridizations can be allowed, but should be taken into account when predicting Tm P . 10. Alternatively, annealing temperature can be held constant and magnesium concentration is varied to identify optimal annealing conditions. We have observed that increasing the magnesium concentration from 3.0 to 3.5 mM has a similar effect to lowering the annealing temperature 2°. Large changes in magnesium concentration, however, may affect Taq activity and change hybridization characteristics of molecular beacons and other probes. Also, note that these tests can be done on genomic DNA rather than single cells, since the limiting primer becomes depleted once it makes sufficient product to reach the detection threshold, regardless of the initial target concentration. Using 600 pg of DNA (equivalent to about 100 genomes) will lower the CT value about eight cycles compared to single cells, but does not change the subsequent linear signal kinetics (6). 11. SYBR Green I binds to double-stranded DNA regardless of nucleotide sequence. Fluorescence therefore plateaus after the limiting primer is exhausted. Following PCR cycling, fluorescence is monitored as temperature is gradually increased. As PCR products denature, a large fluorescence drop is observed. Multiple fluorescence drops, usually evaluated as “melting peaks” on plots of temperature versus the rate of fluorescence decrease, indicate the presence of nonspecific product. Specific reactions should have a single melting peak about 3–6° above the calculated value, depending on the magnesium concentration.


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12. If LATE-PCR is continued for many linear cycles, a second rise in SYBR Green fluorescence may be observed. This corresponds to a phenomenon we call “product evolution,” which involves the single strands priming on one another with a resulting increase in product size and melting temperature. Product evolution usually can be avoided by limiting the number of linear cycles and minimizing the temperature drop needed for probe detection. In rare cases, it may be necessary to modify the 5′ end of the limiting primer, thereby changing the 3′ end of the amplicon single strands, in order to avoid this type of mispriming.

Acknowledgments Aquiles Sanchez, John Rice, Christina Hartshorn, Kevin Soares, and Jesse Salk have made contributions to the development and testing of LATE-PCR. This work was funded by Brandeis University. References 1. Tyagi, S. and Kramer, F.R. (1996) Molecular beacons: probes that fluoresce upon hybridization. Nat Biotechnol 14, 303–308. 2. Heid, C.A., Stevens, J., Livak, K.J., and Williams, P.M. (1996) Real time quantitative PCR. Genome Res 6, 986–994. 3. Kostrikis, L.G., Tyagi, S., Mhlanga, M.M., Ho, D.D., and Kramer, F.R. (1998) Spectral genotyping of human alleles. Science 279, 1228–1229. 4. Gyllensten, U.B. and Erlich, H.A. (1988) Generation of single-stranded DNA by the polymerase chain reaction and its application to direct sequencing of the HLA-DQA locus. Proc Natl Acad Sci USA 85, 7652–7656. 5. Sanchez, J.A., Pierce, K.E., Rice, J.E., and Wangh, L.J. (2004) Linear-after-theexponential (LATE)-PCR: an advanced method of asymmetric PCR and its uses in quantitative real-time analysis. Proc Natl Acad Sci USA 101, 1933–1938. 6. Pierce, K.E., Rice, J.E., Sanchez, J.A., and Wangh, L.J. (2005) Linear-after-theexponential (LATE)-PCR: primer design criteria for high yields of specific single-stranded DNA and improved real-time detection. Proc Natl Acad Sci USA 102, 8609–8614. 7. Pierce, K.E., Rice, J.E., Sanchez, J.A., and Wangh, L.J. (2003) Detection of cystic fibrosis alleles from single cells using molecular beacons

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and a novel method of asymmetric real-time PCR. Mol Hum Reprod 9, 815–820. Salk, J.J., Sanchez, J.A., Pierce, K.E., Rice, J.E., Soares, K.C., and Wangh, L.J. (2006) Direct amplification of single-stranded DNA for pyrosequencing using linear-after-the-exponential (LATE)-PCR. Anal Biochem 353, 124–132. Rice, J.E., Sanchez, J.A., Pierce, K.E., Reis, A.H. Jr., Osborn, A., and Wangh, L.J (2007) Monoplex/multiplex linear-after-theexponential-PCR assays combined with PrimeSafe and Dilute-‘N’-Go sequencing. Nat Protoc 2, 2429–2438. Pierce, K.E., Rice, J.E., Sanchez, J.A., and Wangh, L.J. (2002) QuantiLyse™: reliable DNA amplification from single cells. Biotechniques 32, 1106–1111. Allawi, H.T. and SantaLucia, J. (1997) Thermodynamics and NMR of internal G.T mismatches in DNA. Biochemistry 36, 10581–10594. SantaLucia, J. (1998) A unified view of polymer, dumbbell, and oligonucleotide DNA nearest-neighbor thermodynamics. Proc Natl Acad Sci USA 95, 1460–1465. Owczarzy, R., Vallone, P.M., Gallo, F.J., Paner, T.M., Lane, M.J., and Benight, A.S. (1998) Predicting sequence-dependent melting stability of short duplex DNA oligomers. Biopolymers 44, 217–239.

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14. Breslauer, K.J. (1986) Methods for obtaining thermodynamic data on oligonucleotide transitions. In Hinz, H. (ed.) Thermodynamic Data for Biochemistry and Biotechnology, Springer-Verlag, New York, pp. 402–427. 15. Le Novère, N. (2001) MELTING, computing the melting temperature of nucleic acid duplex. Bioinformatics 17, 1226–1227. 16. SantaLucia, J., Allawi, H.T., and Seneviratne, P.A. (1996) Improved nearest-neighbor parameters for predicting DNA duplex stability. Biochemistry 35, 3555–3562. 17. Wetmur, J.G. (1991) DNA probes: applications of the principles of nucleic acid hybridization. Crit Rev Biochem Mol Biol 26, 227–259. 18. Zuker, M. (2003) Mfold web server for nucleic acid folding and hybridization prediction. Nucleic Acids Res 31, 3406–3415. 19. Marras, S.A.E., Kramer, F.R., and Tyagi, S. (2003) Genotyping single nucleotide polymorphisms with molecular beacons. In Kwok, P.Y. (ed.) Single Nucleotide Polymorphisms: Methods and Protocols, Humana Press, Totowa, NJ, Vol. 212, pp. 111–128. 20. Ririe, K.M., Rasmussen, R.P., and Wittwer, C.T. (1997) Product differentiation by analysis of DNA melting curves during the polymerase chain reaction. Anal Biochem 245, 154–160. 21. Pierce, K.E., Rice, J.E., Sanchez, J.A., Brenner, C., and Wangh, L.J. (2000) Realtime PCR using molecular beacons for accurate detection of the Y chromosome in single human blastomeres. Mol Hum Reprod 6, 1155–1164.

22. Rice, J.E., Sanchez, J.A., Pierce, K.E., and Wangh, L.J. (2002) Real-time PCR with molecular beacons provides a highly accurate assay for Tay–Sachs alleles in single cells. Prenat Diagn 22, 1130–1134. 23. Rosner, B. (1995) Fundamentals of Biostatistics. Wadsworth Publishing Company, Belmont, CA, pp. 277–282. 24. Rechitsky, S., Verlinsky, O., Amet, T., Rechitsky, M., Kouliev, T., Strom, C., and Verlinsky, Y. (2001) Reliability of preimplantation diagnosis for single gene disorders. Mol Cell Endocrinol 183, S65–S68. 25. Al-Soud, W.A., Jonsson, L.J., and Radstrom, P. (2000) Identification and characterization of immunoglobulin G in blood as a major inhibitor of diagnostic PCR. J Clin Microbiol 38, 345–350. 26. Al-Soud, W.A. and Radstrom, P. (2001) Purification and characterization of PCRinhibitory components in blood cells. J Clin Microbiol 39, 485–493. 27. Cui, X.F., Li, H.H., Goradia, T.M., Lange, K., Kazazian, H.H. Jr., Galas, D., and Arnheim, N. (1989) Single-sperm typing: determination of genetic distance between the G gamma-globin and parathyroid hormone loci by using the polymerase chain reaction and allele-specific oligomers. Proc Natl Acad Sci USA 86, 9389–9393. 28. Li, Q., Luan, G., Guo, Q., and Liang, J. (2002) A new class of homogeneous nucleic acid probes based on specific displacement hybridization. Nucleic Acids Res 30, e5. 29. Cheng, J., Zhang, Y., and Li, Q. (2004) Realtime PCR genotyping using displacing probes. Nucleic Acids Res 32, e61.

Chapter 6 Long-PCR Amplification of Human Genomic DNA Stephen Keeney Abstract Standard polymerase chain reaction (PCR) protocols amplify relatively small fragments precluding the use of this approach when examining gross rearrangements of DNA. By using combinations of DNA polymerases, which feature either good polymerase activity or error-correction abilities, it is now possible to extend the length of DNA fragment that can be amplified. These “long-PCR” protocols have allowed the development of more rapid and convenient ways to analyse large-scale rearrangements of DNA and in many cases has superseded alternative approaches such as Southern blotting. The protocol described in this chapter illustrates some of the key points to be considered when developing a long PCR protocol. Key words: Polymerase chain reaction, Long PCR, DNA polymerase, Primer design, F8 gene

1. Introduction The development of polymerase chain reaction (PCR) methodology (1) allowed a revolution in molecular genetic analysis to take place. The original protocol utilised a thermostable DNA polymerase derived from the thermophile Thermus aquaticus. This DNA polymerase (Taq) is still used in the majority of PCR applications as it has a high processivity (good 5′-3′ polymerase activity allowing a higher speed of incorporation of bases into a growing DNA chain), but it does have limitations. Taq has a relatively high error rate when incorporating new bases, estimated at somewhere between 1 × 10−4 and 9 × 10−5 errors per base pair, dependent on the assay used. This lack of replication fidelity results in disassociation of the polymerase from the elongating DNA strand, imposing a limit on the length of DNA fragment that can be amplified (amplicon). The size of amplicon obtainable when using native Taq varies depending on the nature and source Bimal D.M. Theophilus and Ralph Rapley (eds.), PCR Mutation Detection Protocols, Methods in Molecular Biology, vol. 688, DOI 10.1007/978-1-60761-947-5_6, © Springer Science+Business Media, LLC 2011

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of the template, but for human genomic DNA the upper limit for reliable amplification in a diagnostic context tends to be between 2 and 3 kb and certainly no more than 4–5 kb. An alternative class of thermostable DNA polymerases, which are capable of error correction (good 3′-5′ exonuclease or “proofreading” abilities) exist. This feature allows for higher fidelity during replication; however, proofreading enzymes tend to have a poor processivity resulting in low amplicon yields. Combining DNA polymerases with these two complementary features allows the processivity of Taq to be paired with the error-correction abilities of a proofreading enzyme such as Pfu, resulting in the amplification of much larger fragments (2). This approach allowed the development of “long-PCR” (L-PCR) protocols where amplicons much larger than those achieved using standard Taq polymerase can be generated. Amplification of fragments greater than 20 kb has been demonstrated from human genomic DNA and in excess of 40 kb from phage lambda DNA (3). L-PCR based protocols have subsequently been applied to the detection of gross rearrangements of DNA, which were previously the domain of Southern blotting based protocols including: detection of partial gene deletions and gene rearrangements, detection of chromosomal translocations, and amplification through long stretches of GC rich or repetitive sequence elements. Since the first description of L-PCR a variety of thermostable DNA polymerase combinations have been developed by various manufacturers (see Table 1). These enzyme mixes often come in kit form and may have several buffers designed to optimise amplification over different size ranges. In routine diagnostic conditions, several factors influence the upper limit of amplification product size (see Notes 1–7). A minimum amplicon size of 12–15 kb should be routinely achievable, and given the appropriate conditions, in excess of 20 kb fragments will amplify. The L-PCR protocol detailed below for detection of the FVIII gene (F8) intron 22 inversion

Table 1 Examples of commercial long-PCR kits Product name

Supplier

Demonstrated amplicon size (kb)a

LongAmp™ Taq PCR Kit

New England Biolabs

30

LA PCR Amplification Kit, Ver. 2.1

TaKaRa Bio

27

LongRange PCR Kit

Qiagen

27

Expand Long Range dNTPack

Roche

25.6

Extensor Hi-Fidelity PCR Master Mixb ABgene

23

GeneAmp® XL PCR Kit

19.6

Applied Biosystems

Information from supplier relating to amplification from human genomic DNA This product is used in the protocol listed below

a

b

DNA Amplification by Long PCR

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mutation (4, 5) generates analysed results within 24 h, compared to 1 week for the original published protocol based on Southern blotting (6).

2. Materials 1. Source of DNA to be amplified: The template DNA must be of good quality. Degraded or sheared DNA will not permit amplification of larger fragments. For human genomic DNA, this equates to a band which runs on a gel with a size estimate of >50 kb. The DNA extraction protocol utilised must produce DNA of a high purity (Optical Density 260/280 nm ratio as near to 1.8 as is possible). For general considerations relating to DNA extraction, see Note 1. 2. Oligonucleotide primers: (a) INT22 P 5′-GCCCTGCCTGTCCATTACACTGATGA CATTATGCTGAC-3′ (38 mer): Make a 4 mM stock. (b) INT22 Q 5′-GGCCCTACAACCATTCTGCCTTTCAC TTTCAGTGCAATA-3′ (39 mer): Make a 4 mM stock. (c) INT22A 5′-CACAAGGGGGAAGAGTGTGAGGGTGT GGGATAAGAA-3′ (36 mer): Make a 2 mM stock. (d) INT22B 5′-CCCCAAACTATAACCAGCACCTTGA ACTTCCCCTCTCATA-3′ (40 mer): Make a 2 mM stock. Store stocks at −80°C. Working stocks can be made by combining equal volumes of individual primers, which can be stored at −20°C. For primer design considerations, see Note 2. 3. Extensor Hi-Fidelity PCR Master Mix (ABgene) contains Extensor PCR enzyme Mix, dNTPs, Extensor reaction buffer 2, and MgCl2. Additional MgCl2 or dNTPs are not required for this specific protocol (see Note 3). Store at −20°C. 4. DMSO (Dimethyl Sulphoxide): PCR-grade DMSO (Sigma). Store at −20°C (see Note 4). 5. Sterile water: This should be high-quality water. 10 mL vials designed for parenteral use are ideal. Store at room temperature. 6. Reaction vessels: Minimising thermal lag times in L-PCR protocols is critical. Use thin-walled plasticware for reactions. 7. Thermal Cycler: Machines should have fast ramp rates and the ability to increase the hold time per extension cycle. A heated lid is desirable to avoid mineral oil overlay (see Note 5). 8. Agarose Gel Electrophoresis (a) Molecular Biology-grade agarose powder. (b) 10× Tris borate EDTA buffer (TBE): 108 g Tris, 55 g boric acid, 40 mL 0.5 M EDTA, pH 8.0. Make up to 1 L

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with distilled water. For use, dilute to 0.5× concentration (50 mL/950 mL of water). Store at room temperature. (c) Ethidium bromide: 10 mg/mL in sterile water (this is considered to be a mutagen so observe appropriate safety precautions). Store at room temperature.

3. Methods Modifications for amplification of the F8 intron 22 gene inversion include the addition of DMSO to facilitate read through of a high GC content region upstream of F8. Use of the ABgene Extensor PCR kit simplifies the reaction set-up procedure from that presented in the original publication (4). This specific protocol uses two sets of L-PCR primers which detect the rearrangement of an inverted DNA segment by overlapping PCR (see Fig. 1). The amplification products range from 10 to 12 kb which is well within the range of the enzyme mix utilised. 3.1. Long-Range PCR Set-Up

1. Allow each reagent to thaw fully; mix and spin down before pipetting. 2. Combine equal volumes of the 2 mM stock A and B primers, mix and spin down. Repeat for the 4 mM stock P and Q primers. 3. Prepare the L-PCR master mix on ice (see Table 2). Reactions must be set up in thin-walled PCR tubes. For the above scheme make a master mix of all ingredients (excluding each DNA sample) times the number of samples to be amplified,

Fig. 1. L-PCR amplification of the F8 intron 22 inversion mutation. The lower 10 kb band (Int22 A+B primer pair product) and the 12 kb upper band (Int22 P+Q product) represent sequences that have not undergone rearrangement. The middle 11 kb band consists of two new specificities (Int22 PB and AQ) created when the DNA inversion is present. Carrier females for the inversion indicate every possible L-PCR product specificity since they have two copies of F8 (one normal and one mutated). Males have one copy of the gene and will only show either the inverted or normal pattern, depending on the presence or absence of the inversion mutation.


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Table 2 L-PCR master mix for amplification of the F8 intron 22 inversion Component

1× Reaction volume (mL)

10× Reaction master mix (mL)

ABgene Extensor Mix

12.5

125

PQ primer mix

2.0

20

AB primer mix

3.0

30

DMSO

1.9

19

H2O

4.6

46

DNA (100–200 ng/mL)

1.0

–

Final volume

25

24 per reaction

including controls. This minimises pipetting losses of reagents and should decrease any tube-to-tube variability between reactions. The above scheme assumes the addition of 1.0 mL of template DNA to each reaction. If different amounts of template are used then adjust the volume of water in the master mix accordingly. Ensure that reactions are prepared and kept on ice and that they are transferred immediately to the PCR step (see Note 6). Always include a “noDNA” control (blank reaction consisting of master mix with genomic DNA substituted by an equal volume of sterile water). 3.2. Subcycling Long PCR

1. Prior to reaction set-up, preheat the thermal cycler block to 94°C and leave on standby. 2. Working quickly, transfer the L-PCRs in their thin-walled tubes directly from ice to the thermal cycler, close the heated lid and activate the following programme (see Note 7): Initial denaturation of 94°C for 2 min. Then, ten cycles consisting of: 94°C for 12 s. Four subcycles per main cycle consisting of: 60°C for 120 s. 65°C for 120 s. Remaining 16 cycles: 94°C for 12 s. Four subcycles per main cycle consisting of: 60°C for 120 s plus an additional 3 s added per cycle. 65°C for 120 s plus an additional 3 s added per cycle.

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3.3. Agarose Gel Electrophoresis

1. Prepare a 0.7% agarose gel (see Note 8): 0.7 g agarose added to 100 mL of 0.5× TBE. Mix and heat in a microwave until fully dissolved, cool to hand hot (60°C). Take care when heating and handling hot agarose gel solutions/as they may superheat and boil over when mixed. Add 2 mL of 10 mg/mL ethidium bromide (0.5 mg/mL final concentration). Use a 1-mm thick 16-well gel comb. Pour gel and allow to set. 2. The PCR product does not need any loading dye as this is contained in the ABgene Extensor mix. Load 5 mL of each PCR product into individual wells and run the gel at 130 V for 3–4 h minimum (see Note 9). 3. Visualise bands on the gel using a transilluminator and photograph.

3.4. Interpretation of Banding Pattern

An upper 12 kb band and a lower 10 kb band only: The inversion is not present. A middle 11 kb band and a lower 10 kb band: The inversion is present in an affected male. Bands at 10, 11, and 12 kb indicate a female carrier of the inversion (see Fig. 1).

4. Notes 1. The chosen DNA extraction protocol should minimise the risk of shearing of DNA. If phenol-based extraction methods are used avoid vortex mixing and ensure that supernatant transfer steps are performed using sterile disposable pastettes or wide-bore tips. Phenol must be fresh and acidic conditions should be avoided. Alternative DNA extraction methods, such as those based on ammonium acetate precipitation, work well for this particular protocol. 2. PCR primers should be longer (typically 24–40 bp depending on sequence composition) than those typically used in standard PCR and should have a higher melting temperature (Tm). A Tm between 63 and 68°C is considered optimal, allowing for higher reaction specificity. The Tm difference between primers used should be as close as possible and certainly within 2°C. Close Tm tolerances across primer sets are best achieved using primer design software. A readily available tool suitable for this purpose is Primer3, which allows very precise control of primer parameters (7). 3. As a general rule the magnesium and dNTP concentrations are higher when performing L-PCR. When using kits with optimised component concentrations (see Table 1) this issue is largely taken care of. This particular protocol uses the ABgene Extensor kit which allows for a very streamlined


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reaction set-up since most reaction components are included in the mix. However, different sizes of amplicon will require careful selection of enzyme mix and additional reaction component concentrations, particularly when amplifying very large fragments. Too much enzyme in a reaction will tend to produce a smear of products across a broad size range. If a smear is present titrate the polymerase used in the reaction. 4. Some applications may require additional components to be added to the reaction mix to ensure efficient amplification. In particular, GC-rich templates may require substitution of a proportion of guanine dNTPs with the base analogue deaza G and/or inclusion of agents that facilitate strand separation such as DMSO or betaine. Some commercially available kits include buffers containing proprietary reagents that fulfil this function. 5. For some applications, the additional inclusion of a mineral oil overlay may be advised to ensure that the reaction volume remains constant. This is particularly important when setting up smaller volume reactions. 6. The 3¢-5¢ exonuclease activity of high-fidelity DNA polymerases may result in degradation of single-stranded oligonucleotide primer DNA in the reaction. Keep reactions on ice, and after addition of the reaction component containing the DNA polymerase mix, proceed immediately to the thermal cycling step. Some kits may require a two-stage master mix set-up, ensuring that the polymerase mix is kept separate from PCR primers until the last possible moment. 7. When optimising protocols cycling conditions are critical. Denaturation times need to be kept to a minimum (12 s or less is recommended) to avoid degradation of newly synthesised product. Extension times are long and need to be extended for each additional cycle. The subcycling temperature shuttle element of this protocol aids/read through GC-rich sequence elements (5). When optimising alternative protocols use of a subcycling approach may also prove useful. This particular protocol has a reduced number of cycles to avoid overamplification and alternative protocols may need additional cycles. 8. The amplicons produced in this example can be analysed on standard low-percentage agarose gels. However, larger fragments may require pulsed-field gel electrophoresis (PFGE). Ensure that the agarose has dissolved fully before pouring the gel. 9. An appropriate DNA size marker should be used to verify the size of amplicons for the particular protocol.

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Acknowledgments The author would like to thank Anne Goodeve, Marian Hill, and David Stirling for their advice during the development of this specific protocol and Dörte Wren for technical assistance. References 1. Saiki R. K., Gelfand D. H., Stoffel S., Scharf S. J., Higuchi R., Horn G. T., Mullis K. B., and Erlich H. A. (1988) Primer-directed enzymatic amplification of DNA with a thermostable DNA polymerase. Science 239, 487–91. 2. Barnes W. M. (1994) PCR amplification of up to 35-kb DNA with high fidelity and high yield from lambda bacteriophage templates. Proc Natl Acad Sci USA 91, 2216–20. 3. Cheng S., Fockler C., Barnes W. M., and Higuchi R. (1994) Effective amplification of long targets from cloned inserts and human genomic DNA. Proc Natl Acad Sci USA 91, 5695–99. 4. Liu Q., Nozari G., and Sommer S. S. (1998) Single-tube polymerase chain reaction for

rapid diagnosis of the inversion hotspot of mutation in hemophilia A. Blood 92, 1458–9. Erratum in Blood (1999) 93, 2141. 5. Liu Q. and Sommer S. S. (1998) SubcyclingPCR for multiplex long distance amplification of regions with high and low GC content: application to the inversion hotspot in the FVIII gene. Biotechniques 25, 1022–28. 6. Lakich D., Kazazian H. H. Jr, Antonarakis S. E., and Gitschier J. (1993) Inversions disrupting the factor VIII gene are a common cause of severe haemophilia A. Nat Genet 5, 236–41. 7. Primer 3 URL: http://primer3.sourceforge. net/webif.php.

Chapter 7 Human Papilloma Virus Strain Detection Utilising Custom-Designed Oligonucleotide Microarrays Duncan Ayers, Mark Platt, Farzad Javad, and Philip J.R. Day Abstract Within the past 15 years, the utilisation of microarray technology for the detection of specific pathogen strains has increased rapidly. Presently, it is possible to simply purchase a pre-manufactured “off the shelf ” oligonucleotide microarray bearing a wide variety of known signature DNA sequences previously identified in the organism being studied. Consequently, a hybridisation analysis may be used to pinpoint which strain/s is present in any given clinical sample. However, there exists a problem if the study necessitates the identification of novel sequences which are not represented in commercially available microarray chips. Ideally, such investigations require an in situ oligonucleotide microarray platform with the capacity to synthesise microarrays bearing probe sequences designed solely by the researcher. This chapter will focus on the employment of the Combimatrix® B3 CustomArray™ for the synthesis of reusable, bespoke microarrays for the purpose of discerning multiple Human Papilloma Virus strains. Key words: Microarray, Custom design, Combimatrix, HPV, Papilloma, Pathogen screening, Oligonucleotide

1. Introduction Over the best part of the last 15 years, the quest for the identification of pathogens at the molecular level has grown to rely heavily on the involvement of microarray technology (1). The plausible reasons for the utilisation of microarray platforms for such studies include the simplicity of the experimental procedure and the ever increasing throughput levels available through advances in microarray designs (2). These advances brought about enhanced sensitivity, reproducibility, and reliability of the data obtained (3), compared with the early days of microarray studies. Human papilloma viruses (HPVs) are small DNA viruses which have been established as causative factors in a wide spectrum Bimal D.M. Theophilus and Ralph Rapley (eds.), PCR Mutation Detection Protocols, Methods in Molecular Biology, vol. 688, DOI 10.1007/978-1-60761-947-5_7, © Springer Science+Business Media, LLC 2011

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of female genital conditions, including cervical cancer (4). The entire HPV class consists of over 120 viruses, with only 30 being known to induce female genital tract clinical conditions, the most notorious condition being high-grade cervical dysplasia and squamous cervical cancer which is induced by HPV16 and HPV18 infection (4). Due to the vast quantity of HPV strains present, efficient clinical HPV screening techniques at the genomic level may prove vital for the clinician as such methods would discern between HPV strains which are high/low risk for cervical cancer pathogenesis (5). Presently, the investigator may be able to simply purchase a pre-manufactured “off the shelf” oligonucleotide array bearing unique probes for known target pathogen strains (6, 7). However, there exists a problem if the investigator wishes to identify novel pathogen strain sequences which are not represented in commercially available microarray chips. Ideally, such investigations require an in situ oligonucleotide microarray platform with the capacity to synthesise microarrays bearing unique probe sequences, designed solely by the researcher. The experimental data collected from initial pathogen strain detection microarray designs would be used to identify any informative probes. Consequently, any such future microarray designs will carry only strain-specific informative probes, while minimising the risk of cross-hybridisation effects. Obviously, such a microarray platform bearing this level of versatility in oligonucleotide design must still adhere to high microarray manufacturing standards and data reproducibility. Furthermore, the method used for designing and manufacturing such bespoke oligonucleotide microarrays in the investigator’s laboratory must be rapid and user-friendly. Here, we present an example using the Custom array B3 synthesiser technology (Combimatrix, Seattle, USA) that allows laboratories to rapidly produce custom arrays that are capable of performing multiple hybridisations. The core of the technology of the CustomArray™ is a semiconductor chip (see Fig. 1) bearing up to 95,000 electrodes that can be individually activated digitally, enabling synthesis of oligonucleotides of varying lengths (up to a maximum of 40 bases) using standard phosphoramidite chemistry (8). The incorporated technology enables the microarrays not only to be analysed using standard fluorescence assays, discussed in detail here, but also uniquely via electrochemical detection (9). The essential steps necessary to carry out such a study involve the initial design of the microarray, followed by actual synthesis of the oligonucleotide probes on the microarray electrodes. The microarray is then suitable for hybridisation with any fluorescently labelled (ideally Cy3 or Cy5) DNA-binding sample, according to the study designed by the investigator. Finally, the hybridised microarray is then imaged and the resultant data may be collected and analysed via appropriate bioinformatics tools.

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Fig. 1. Diagrammatic representation of a 12K CustomArray™ microarray chip (left ) and the Combimatrix B3 Synthesiser (right ). The semiconductor surface consists of 12,544 individual electrodes, each capable of allowing the synthesis of user-defined oligonucleotide sequences (Copyright Combimatrix Corporation).

This chapter will describe in detail the practical steps necessary to simultaneously synthesise four bespoke microarray designs bearing probes complementary to 20 different HPV strains using the Combimatrix CustomArray™ B3 DNA synthesiser. Once synthesised, the microarrays are hybridised with a fluorescently labelled template oligonucleotide having a signature sequence for HPV18, followed by microarray imaging and data extraction steps.

2. Materials 2.1. Microarray Design and Protocols

1. All Combimatrix software is free to download via the website http://www.bioapps.combimatrix.com. Download the Layout Designer software package (40 MB, system requires 512 MB of RAM). Together with the Layout Designer software you will also need a factory layout for the type of chip you are planning to design and synthesise; these can be obtained directly from Combimatrix. 2. Combimatrix protocols can be found at http://www. combimatrix.com/support_docs.htm. B3 synthesiser software comes with the purchase of a Combimatrix custom array synthesiser. Best results can be achieved by following the official protocol closely; what follows is an overview of these procedures with modifications that this laboratory has found to compliment existing Standard Operating Procedures.

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2.2. Microarray Synthesis

1. 10 g of each of dimethoxytrityl (DMT)-dA(bz), DMT-dC(bz), DMT-dG(ib), and DMT-dT amidites. The amidites must be stored at 2–5°C (see Note 1). 2. 4 L Acetonitrile (ACN). 3. 450 mL TCA Deblock, ABI (see Note 1). 4. 2 L 0.25 M ETT Activator (see Note 1). 5. 2 L Cap A (see Note 1). 6. 2 L Cap B (see Note 1). 7. 0.02 M 2 L Oxidiser (see Note 1). 8. 500 g Tetraethylammonium p-toluenesulfonate. 9. 500 g 99% Hydroquinone, 99%, 500 g. 10. 4 L Reagent grade methanol. 11. 99% Redistilled 2,6-lutidine. 12. 25 g Benzoquinone. 13. 1 kg Molecular Sieve UOP Type 3 Å. 14. B3 synthesiser (Combimatrix, Seattle, USA). 15. Blank microarray chips (Stretton Scientific, UK – distributor of Combimatrix products for UK. Contact Combimatrix directly for your local distributor). 16. Pure shield 99.99% purity argon gas (BOC industrial, UK).

2.3. Microarray Deprotection

1. 100 mL Ethylenediamene (EDA). EDA is corrosive and should be utilised ideally in a fume cabinet. 2. 4 L reagent grade ethanol, SDA formula 3A. 3. 1 Ga reagent grade alcohol. 4. Deprotection manifold (Combimatrix, Seattle, USA).

2.4. Quality Control and Microarray Imaging

1. Random 9mer Cy5 or Cy3-modified 3′ (Tebu-Bio, Cambs., UK). 2. Dibasic sodium phosphate. 3. Monobasic sodium phosphate. 4. 100% glycerol. 5. Tween 20. 6. Microarray Lifter Slip™ cover slip (supplied with microarray slide). 7. Quality Control manifold (Combimatrix, Seattle, USA). 8. A fluorescent microarray scanner, such as the GenePix® 4000B scanner (Molecular Devices, Sunnyvale, CA, USA). 9. Microarray Imager package (68 MB, system requires 512 MB of RAM), which is free to download via the website http:// www.bioapps.combimatrix.com.


2.5. Microarray Stripping

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1. 12K or 90K microarray stripping kit (Combimatrix, Seattle, USA). The stripping solution included in this kit contains ethanolamine which could induce severe burns and is very corrosive. Consequently, this solution must always be handled in a fume cabinet, whilst wearing appropriate personal protective clothing. 2. Nuclease-free water. 3. A fluorescent microarray scanner, such as the GenePix® 4000B scanner (Molecular Devices, Sunnyvale, CA, USA). 4. 10× Phosphate-buffered saline (PBS), pH 7.4 (Ambion, Warrington, UK), diluted to 1× PBS. 5. 95% Ethanol (EtOH) solution (Sigma-Aldrich, Dorset, UK).

2.6. Microarray Hybridisation with Labelled DNA Sample

1. DNA sample with appropriate fluorescent dye (ideally Cy3 or Cy5). 2. PCR sealing tape. 3. Nuclease-free water. 4. 2× Hybridisation solution stock. This is prepared by adding 6 mL 20× SSPE, 100 mL of 10% Tween 20, 560 mL of 0.5 M EDTA, and 3.34 mL of nuclease-free water. The total volume of the solution should be 10 mL Filter sterilise and store at room temperature for up to 6 months. 5. Pre-hybridisation solution (pre-hyb) for 90K microarray. Each 120 mL pre-hyb solution should be prepared fresh and consist of 60 mL of 2× hybridisation solution stock, 41 mL nuclease-free water, 12 mL of 50× Denhardt’s solution, 1 mL of salmon sperm DNA 10 mg/mL (this is to be heated to 95°C for 5 min then cooled on ice for at least 1 min prior to adding to pre-hyb solution), and 6 mL of 1% sodium dodecyl sulphate (SDS). 6. Hybridisation (hyb) solution for DNA targets using a 90K microarray. Each 120 mL volume should consist of 60 mL 2× hybridisation solution stock, 1 mL of salmon sperm DNA 10 mg/mL (heat to 95°C for 5 min then cool on ice for at least 1 min prior to adding to hyb solution), and 5 mL of 1% SDS, up to 4 mg of the labelled DNA target in up to 24 mL volume. Add nuclease-free water to 120 mL. 7. Post-hybridisation wash solutions. The following solutions should be prepared as 10 mL aliquots which are to be filter sterilised and can be then stored at room temperature for up to 6 months. 6× SSPET wash: consists of 3 mL 20× SSPE, 50 mL 10% Tween 20, 6.95 mL nuclease-free water. 3× SSPET wash: consists of 1.5 mL 20× SSPE, 50 mL 10% Tween 20, 8.45 mL nuclease-free water. 0.5× SSPET wash: consists of 250 mL 20× SSPE, 50 mL 10% Tween 20, 9.7 mL nuclease-free water.

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PBST wash: consists of 2 mL 10× PBS, 100 mL 10% Tween 20, 7.9 mL nuclease-free water. PBS wash: consists of 2 mL 10× PBS pH 7.4, 8 mL nuclease-free water.

3. Methods The study described in this chapter is a preliminary step in the design of an effective and high-throughput microarray diagnostic test for discerning between varying HPV strains present in any clinical sample. This group has designed a specific tiling microarray (see Fig. 2) bearing nonamer probes complementary to unique nine consecutive base sequences. The nonamer target sequences consist of fragments from a 65 bp strain-specific DNA sequence for each of the 20 HPV most pathogenic strains (see Table 1). AATAGACTGACAAATATCCAATGGTACCTCACATTTAGTATCTTGCAATGTACTAAAGTCCATGGCACCATAT

a

AATAGACTG ATAGACTGA TAGACTGAC AGACTGACA

Actual target HPV strain DNA sequence (from NCBI)

Tiling approach to probe design

c

F

3’

F

TTTTTTTTTTTGTCAGTCT

Microarray surface

TTTTTTTTTTGTCAGTCTA

5’

F

F

TTTTTTTTTTTCAGTCTAT

Fluorescently tagged template oligonucleotide (mimicking HPV18 DNA) TTTTTTTTTTCAGTCTATT

TTTTTTTTTTTGTCAGTCT

3’

TTTTTTTTTTGTCAGTCTA

b

TTTTTTTTTTTCAGTCTAT

5’

TTTTTTTTTTCAGTCTATT

Reverse complement and T spacer addition

Microarray surface

Fig. 2. Steps involved in the design of a tiling microarray for the purpose of HPV strain detection. (a) The actual DNA sequence for the target HPV strain (from NCBI) is segmented into nonamer fragment sequences, varying by a single base pair shift along the entire target sequence. (b) The reverse complement sequences for the nonamers are used as the actual microarray probes for synthesis, together with a multiple thymidine tail which allows for more effective hybridisation due to the probe being raised away from the microarray surface, thus it is more freely accessible. (c) This group used a fluorescently tagged [Kreatech, Netherlands] template oligonucleotide (TO) [Metabion, Germany] having a 73 base pair sequence which mimics a unique signature sequence for HPV18. This TO is hybridised with the tiling microarray and the probes which are complementary to HPV18 are expected to bind avidly to the TO, with resultant fluorescence demonstrated after microarray scanning and data analysis.


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Table 1 List of target HPV strains utilised for designing the HPV detection tiling microarray, together with the NCBI accession numbers from which the actual DNA sequences were derived Human papilloma virus strain

NCBI accession no.

Tiling microarray probes

HPV18

NC_001357

1–65

HPV26

NC_001583

66–130

HPV28

U31783

131–195

HPV29

U31784

196–260

HPV3

X74462

261–325

HPV30

X74474

326–390

HPV45

X74479

391–455

HPV51

M62877

456–520

HPV53

NC_001593

521–585

HPV54

NC_001676

586–650

HPV56

X74483

651–715

HPV59

X77858

716–780

HPV66

U31794

781–845

HPV68A

DQ080079

846–910

HPV69

AB027020

911–975

HPV70

U21941

976–1,039

HPV77

Y15175

1,040–1,104

HPV82

AB0207021

1,105–1,169

HPV94

AJ620211

1,170–1,234

HPV97

DQ080080

1,235–1,299

Template oligonucleotide used to mimic HPVI8 DNA AATAGACTGACAAATATCCAATGGTACCTCACATTTAGTATCTTGCAATGTACTAAAGTCC ATGGCACCATAT The third column represents the probe annotation scheme for use during post-hybridisation data analysis (e.g. probes 1–65 are all uniquely complementary to nonamer regions of the HPV18 strain-specific DNA sequence). The HPV18 strain-specific template sequence utilised in the design of the HPV18 microarray probes and the fluorescent HPV18 mimicking template oligonucleotide is listed in the bottom of the table.

Synthesis of the designated oligonucleotide probe sequences occurs in each of the individual electrodes present on the semiconductor surface of the microarray, unless the electrode was deliberately designated not for oligonucleotide synthesis (null spots).

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Phosphoramidite chemistry is involved in the manufacturing of the user-defined probe sequence in the individual electrode’s “virtual flask” (see Figs. 3 and 4). 1. All probes sequences should be no longer than 40 bases in length (nine bases in this study) to ensure successful synthesis, although probes of up to 45 bases in length have been made by this group without adverse effects. For shorter probes (less than 15 bases in length), a multi “T” spacer can be added to the 3′-end. The addition of such a spacer extends the sequence away from the chip surface and can enhance the signal (see Note 2).

3.1. Microarray Design

Deprotection a

b

c

OCH3 BASE1

O H3CO

P O

O O

CN

P

O

O

NC O

CN

BASE2

DMTO

O

P

N(iPr)2

tetrazole

BASE2

DMTO

O

O O

O

O

O

MEMBRANE

MEMBRANE

NC

O O

O O

HO H+

O

BASE2

DMTO

BASE1

P

BASE1

O

O

O

P O

2. I2, pyridine, H2O, THF O

O

P

CN

MEMBRANE

BASE1

O

1. Ac2O, lutidine

O O

NC

O O P

O O

O

CN

MEMBRANE

BASE = protected A or T or G or C

Fig. 3. The four main steps involved in each nucleotide addition initiate with detritylation of the dimethoxytrityl (DMT) blocking group (a) on the 5′-hydroxyl group, in order to allow access to the additional base. This step is kick-started by electrochemically inducing an acidic environment (b) in the individual electrodes which are designated for oligonucleotide elongation by a specific base. Consequently, the synthesiser allows for the introduction of the following phosphoramidite and activating reagent, which leads to coupling to the deblocked 5′-hydroxyl group of the previous phosphoramidite (c). An acetylation capping step is also required to inhibit any uncoupled nucleotides from coupling in the following nucleotide addition cycle. Finally, the newly formed phosphite group is acid labile and thus needs to be stabilised by inducing an oxidation reaction from phosphite to a phosphate group. This cycle is repeated until the desired oligonucleotides are fully synthesised on the microarray electrodes, as initially designed by the investigator. The final phase involves deprotection of the newly synthesised oligonucleotides, which involves removal of the blocking groups, cleavage from the support, and substitution of the 5′-phosphate group with a hydroxyl group (Copyright Combimatrix Corporation).


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Fig. 4. The semiconductor electrode surface (a) possesses purposely designed “virtual flasks” (b) enclosing each of the individual electrodes. Each “flask” consists of a semipermeable matrix (c) which confers spatial restriction on the newly synthesised oligonucleotide (Copyright Combimatrix Corporation).

Fig. 5. Screen shot from Layout Designer software package. The area on the left-hand side of the screen displays the details of the user-designed oligonucleotide sequences, while the right-hand side illustrates a map of the entire electrode surface. The latter also illustrates details of where each individual oligonucleotide has been assigned for synthesis on the electrode surface (Copyright Combimatrix Corporation).

2. Design all target probes using standard annotation, 5′–3′. Place all probes in a spreadsheet into column 1, give each probe an individual name into a second column (N.B. probe names are not limited by any character or symbol). Please note that the probe sequences are synthesised onto the microarray surface from the 3′-end of the designated probe sequence. Save this file as a tab delimited text file, test.txt. 3. Open Layout Designer package (see Fig. 5); the procedure for designing a 12K chip and 90K chip is identical; for the purpose of this chapter a 90K chip shall be synthesised. Open the

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correct factory layout for the chip type planning on being synthesised, from the menu select advanced>open factory layout>90k factory layout.xml. In the window there appears a list of factory layout probes on the left, and on the right the position of the factory probes is displayed. For 90K chips, there exist a total of 94,928 spots for synthesis; 4,398 are taken up by the factory layout probes and null spots. For more complex chip designs such as the 4 × 2K chips, the factory layout and null spots must be left untouched as they show the user where the “O” ring for the hybridisation cap will come into contact with the chips surface, and therefore any sequences placed onto these spots will be excluded from the experiments. However, for basic chip designs with both the 12K and 90K chips, it is not essential to keep the factory layout probes. 4. Import the probe list file>import probe list. The user will be prompted to “select a file containing a two column, tab delimited, probe list with sequence first followed by unique probe name…”, click “ok” and select test.txt. If duplicate probes have been listed within test.txt, a window will open warning the user about the probes that have been identified as replicates and how many replicates are present. Any alterations that are needed to be done to remove these replicates must be done in a spreadsheet and the above steps repeated. If the choice is made to continue with replicates click “ok” and close the box listing the sequences. 5. After importing the target probes, these will be listed in the left-hand window below the factory probes. Highlight the target probes you wish to populate the array with. Next select the number of replicates you wish of the probes, i.e. if you wish to have an array with 90,530 individual sequences leave the value at 0, for 6,000 duplicate sequences enter 2, and so on. In this chapter, we shall use a probe list of 3,900 sequences where 1,300 probes were synthesised in triplicates on a randomised region of the microarray. From the menu, select Probe table>set requested replicates enter the value required, 3, and select “ok” (see Note 3). 6. Next select from the menu microarray>select all. The user will notice the entire array has been highlighted. 7. Select microarray>deselect assigned, spots that have been assigned with a sequence or factory layout have now been deselected. 8. From the menu, select microarray>populate array, you are then presented with several options, >random will be demonstrated here. The array will have now been randomly filled with triplicate spots of the selected sequences (see Note 4). 9. Save the file file>save as>standard>“name”.


3.2. Microarray Synthesis 3.2.1. Preparation of Electrochemical Deblocking Agent

3.2.2. Preparation of Phosphoramidite Solutions and Accessory Reagents

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1. Mix the following chemicals together, ideally in a large amber glass container and within the confines of a fume cupboard (see Note 5); 30 g tetraethylammonium p-toluenesulfonate, 220 g hydroquinone, 2.16 g benzoquinone, 400 mL methanol, 1,400 mL ACN. 2. Having ensured that all chemicals have dissolved, add 1.18 mL of 2,6-lutidine to the container (see Note 6). 1. Remove all amidite chemicals from the fridge an hour before intended use, thus allowing acclimatisation to room temperature. 2. Place ~2 g of molecular sieves into each amidite bottle. 3. Add 200 mL of ACN to each amidite bottle, replace lid, and shake until all the chemical has been dissolved.

3.2.3. Array Chip Loading and Final Procedures for Synthesis Run

1. Connect new chemicals to the B3 synthesiser, open the argon cylinder. 2. If this is the first synthesis run, open the Combimatrix B3 synthesiser software. When in-between runs, the storage procedure for a B3 synthesiser is to fill all reaction chambers and lines with ACN, in such cases you will also need to flush all chambers with argon when prompted by the software. 3. The B3 synthesiser is capable of manufacturing up to eight microarrays during a synthesis run with individual chip designs, and waste is minimised when all eight chambers are set to prepare new chips. 4. Remove four blank 90K microarray chips from their pack aging, open the first four synthesis chambers and remove the white ceramic tile. Place the blank chips into the chambers. 5. Each chip has a unique serial number, make a note of the chip number and synthesis chamber. Place chips into the chambers with the array facing away from the user, then close the chambers. 6. From the menu select action>load chip design file. 7. For chambers with blank chips inside, use the browse feature to find the chip design file saved earlier. If multiple versions of chip design files are being used, make sure to note which chip/chamber corresponds to the correct file. Select “ok”. 8. Initiate the microarray synthesis and leave to run overnight, for four chips containing 30mers the average synthesis time is around 20 h. 9. After the synthesis is complete, remove the chips, insert the white ceramic tiles back into the chambers, and fill the machine with ACN (action>service protocol>fill machine with ACN).

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3.3. Microarray Deprotection

1. The Combimatrix protocol requires the chambers of the manifold to be emptied by flushing with argon gas. For laboratories without an argon cylinder next to a fume cabinet, it is possible to simply use a syringe and flush the chambers with air. All deprotection should be carried out in a fume cabinet. 2. Prepare a solution of EDA/EtOH, 50:50, ~15 mL in total volume. 3. Remove the white ceramic slides from four of the chambers in the deprotection manifold, and place the 90K slides with the array facing the user, in their place. 4. Fill the manifold with the EDA/EtOH mixture, empty, and refill with fresh EDA/EtOH, making sure no air bubbles are trapped over the arrays. 5. Place the manifold into an oven at 65°C for 1 h. Add ~20 mL molecular grade water into the oven together with the manifold, for later use. 6. Remove the manifold and water from the oven and empty the EDA/EtOH from the chambers. 7. Fill manifold with reagent alcohol and then empty immediately. 8. Wash manifold with molecular grade water and then empty immediately. 9. Wash manifold twice with reagent alcohol; after the second wash dry for 60 s either using argon gas or by passing air over the chips. 10. Refill manifold with heated water and place back into the oven at 65°C for 10 min. 11. After 10 min, remove the manifold from the oven, empty the chambers, and allow the chips to dry, then they can be stored in a dessicator for up to 4 months (see Note 7). (The same procedure is followed for 12K arrays.)

3.4. Performing a Quality Control Check

1. Prepare the wash solution at pH 8.0 by dissolving 4.223 g sodium phosphate monobasic and 64.482 g of sodium phosphate dibasic into 1 L of water (see Note 8). 2. Prepare Quality Control (QC) pre-hybridisation solution (pre-hyb) by adding 0.05% Tween 20 to 50 mL of the wash solution (see Note 8). 3. The imaging solution is prepared by adding 10% glycerol to 10 mL of the wash solution (see Note 8). 4. Combine the dye-modified 9mer with 10 mL of pre-hyb solution to give a hybridisation solution (hyb) with a final oligonucleotide concentration of 100 nM. 5. Ensure that the chips are hydrated before performing QC. If the chips were stored after deprotection, place into nucleasefree water at 65°C for 10 min (see Note 9).


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6. Insert the chips into the QC manifold, ensuring the arrays are facing the user. 7. Fill chambers with pre-hyb solution. 8. After 10 min, remove the pre-hyb solution. 9. Fill QC chamber with hyb solution and leave for 60 min at room temperature. To ensure there is no photo bleaching of the dye, cover the manifold chambers with foil or place the manifold away from direct light. 10. Remove hyb solution and wash twice with the wash solution, leaving the second wash solution in the manifold. At this point, do not let the chips dry. 11. Empty the chambers of the second wash solution one at a time. As a chamber empties remove the chip and prepare for imaging. (The same procedure is followed for 12K arrays.) 3.4.1. Imaging

1. Remove the chip from the manifold and place onto a flat surface. 2. Apply 50 mL of imaging solution onto the microarray electrode surface. 3. Spray the Lifter Slip™ with argon compressed gas to remove any debris. 4. Use a razor blade to identify which side of the Lifter Slip™ is to be placed down on to the array. This can be done by dragging the blade over the white edge of each lifter slip. One side should have a raised feature, thus the side that has the raised white line feature should be placed down onto the surface, in order to create a 100 mm gap between the cover slip and electrode surface (see Fig. 6.).

Fig. 6. Illustration of correct orientation for applying cover slip over the CustomArray™ 12K microarray (Copyright Combimatrix Corporation).

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5. Avoid introduction of any air bubbles that may be trapped under the cover slip with the aid of the razor blade, by lowering gently the cover slip onto the electrode surface from one end only (see Fig. 6). 6. Wipe away any excess liquid and place into the array scanner. 7. Combimatrix recommends the GenePix® scanner series, and hence this protocol will use the GenePix® 4000B scanner, with the optimum scanning condition for this machine being: Cy3 PMT gain of 300 for 532 nm scans, Cy5 PMT gain of 400 for 635 nm scans, 100% Power, Pixel size of 5 mm, one line to average, focus position of 100 mm. The most important setting is that of focus position. As Combimatrix microarray chips are imaged wet, the focus position must be adjusted to account for both the solution and cover slip; typically a position of 100 mm is used for 12K chips. However with the 90K format an adjustment will have to be made (see Note 10). 8. Save the image file. (The same procedure is followed for 12K arrays.) 3.4.2. QC Image Analysis: Use of Microarray Imager

The purpose of imaging the chips is to ensure the quality of the synthesis, which can affect electrode (spot) morphology and quality. By performing a QC stage, bad spots can be identified before hybridising to targets: 1. Open Microarray Imager (see Fig. 7), select file>new analysis and select the chip design file that corresponds to the image and its chip number. 2. Images from the scanner are usually saved such that the preview and multiple laser scans are compressed into one file. To separate these scans, choose advanced>convert axon image file>“select image of chip” from the main menu. Microarray Imager will now create a new folder with the same name of the individual image. Placed into this folder are the separate, preview, and multiple laser scans. 3. Open the correct image: file>open image>“select appropriate file”. 4. Select analysis>new template; a grid will appear. To overlay the grid over the image click on advanced>find microarray. The grid should align itself with the array. If the grid fails to find the array, the user has the option of manually aligning the template. Edit>align template, zoom into the top left-hand corner of the grid using image>zoom in and double click on the feature 1, 1. Click onto feature 1, 1 and while holding down the LEFT mouse button move the grid until the first circle, 1, 1 covers the spot on the upper left-hand side of the array. Scroll across the image until the upper right-hand side of the image and grid can be seen. Click once onto the feature 272, 1 for the 90K chip (56, 1 for 12K chips) and while holding the left


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Fig. 7. Screen shot from the Microarray Imager software package. The area on the left-hand side of the screen contains details of the user-designed oligonucleotides (as presented in the layout Designer software package). The right-hand side of the screen represents the actual microarray image following hybridisation. Applying the template onto the image allows the user to collect fluorescence data and also to map each individual electrode to the oligonucleotide sequence which it was instructed to synthesise (Copyright Combimatrix Corporation).

mouse button, move the circle until it covers the spot on the upper right side of the array. Scroll down the image to view the lower right-hand side of the array and grid. Using the RIGHT mouse button click and hold the feature 272, 349 (or 56, 224 for 12K) that drag the cursor to where the bottom right spot can be seen. N.B. the grid itself will not move in the same direction as the cursor. Repeat the last two steps until the grid is aligned with the array. 5. Inspect the array; any spots with comet tails, donuts, or halos should be highlighted. Edit>select spots, use the cursor to select deformed spots, edit>exclude, each excluded spots should appear red. By performing this stage, the spreadsheet that is visible on the left-hand side of the microarray imager will store the defective spots. 6. Click on Analysis>extract data. 7. Click on Advanced>export data with file names. This creates a .txt file capable of being imported into any spreadsheet. This spreadsheet can then be used as a reference during the hybridisation. Any defective spots highlighted in the QC can then be removed from analysis later.

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3.5. Microarray Stripping

1. Wash the stripping plugs and cap with 70% EtOH and dry gently with a tissue. 2. Place a new gasket into the stripping cap O-ring. 3. Insert the hybridised microarray chip into a solution containing 1× PBS in order to allow the Lifter Slip™ to slide off the electrode surface (see Note 11). 4. Collect the microarray chip and wipe the back surface of the chip. 5. Place microarray in the appropriate stripping clamp (see Fig. 8), with the electrode surface facing upwards, then place the stripping cap onto the surface. 6. Close and secure the stripping clamp over the microarray and stripping cap. 7. Transfer the stripping clamp to a fume cabinet, and rinse the stripping chamber with 500 mL stripping solution. Replace with a second 500 mL aliquot of stripping solution. 8. Secure stripping chamber ports with the screw on plugs. Air bubbles might be seen inside the stripping chamber, however, this is normal since the chamber volume is large and the electrode surface will still be kept wet. 9. Place stripping clamp in a 65°C incubator and leave stationary inside for 60 min. 10. Remove clamp from incubator, leave to cool for a few minutes, then unscrew the plugs and remove the stripping solution from the chamber.

Fig. 8. Illustration of assembly of a CustomArray™ 12K chip in the 12K stripping clamp. Place the microarray face up in the designated area (a) electrode and cover the electrode surface with the stripping cap (b). Lock the cap onto the microarray and secure by turning the knob appropriately (c). Once stripping solution has been inserted through the portals, screw plugs can be placed on (d) (Copyright Combimatrix Corporation).


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11. Wash the chamber by inserting 500 mL of 95% EtOH once and immediately removing it afterwards. 12. Repeat step 11 with nuclease-free water. 13. Repeat step 11. 14. Open the stripping clamp and place the microarray in a tube containing 1× PBS. Incubate at 65°C for 20 min in order to rehydrate the electrode surface. Take care to wash the stripping cap and plugs with EtOH after use. 15. Remove the microarray from the incubator, collect it from the PBS solution, and gently dry the back surface of the slide. 16. Proceed with steps 1 and 2 of Subheading 3.4 (described above) for microarray imaging and data collection, respectively. 3.6. Microarray Hybridisation with Labelled DNA Sample

1. Attach hybridisation chamber over microarray electrode surface (see Fig. 9). 2. Fill microarray chamber with 120 mL of nuclease-free water and incubate at 65°C for 10 min in a rotisserie hybridisation oven (see Note 12). 3. Fill hybridisation chamber with 120 mL pre-hyb solution (see Note 13). Cover the port apertures with PCR plate sealing tape and incubate at 50°C for 30 min in the rotisserie hybridisation oven. 4. Replace the pre-hyb solution with approximately 115 mL hybridisation solution, using the same method described in step 3 above, and incubate at 50°C for 16 h in the rotisserie hybridisation oven.

3.6.1. Microarray Post-hybridisation Washing

1. Prior to washing, place the 6× SSPET solution in the hybridisation oven at 50°C for 10 min.

Fig. 9. Illustration of assembly of a CustomArray™ 12K (a) and 90K (b) microarray chips with appropriate hybridisation cap (Copyright Combimatrix Corporation).

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Probes which are specific for HPV18 DNA sequence

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Fig. 10. Fluorescence data analysis from one of the four replicate microarray chips used, following hybridisation of a labelled 73 bp template oligonucleotide (mimicking a strain-specific HPV18 DNA sequence) onto a HPV detection tiling microarray. Probes 1–65 are complementary to the HPV18 signature sequence. Ideally, all 65 probes should have demonstrated high-fluorescence readings as a reflection of binding avidity. This was not the case due to possible template oligonucleotide secondary structure effects which might have prevented effective target accessibility. In addition, the fluorescent label used for this study was designed to attach to all guanidine bases (i.e. no end labelling properties) and might have contributed to hybridisation hindrance. Probes designed to recognise other HPV strains which were found to have cross-hybridisation properties were identified and will thus be removed from utilisation in future HPV detection microarray designs.

2. Fill the hybridisation chamber with 6× SSPET (see Note 14). Incubate in rotisserie at 50°C for 5 min. 3. Replace the 6× SSPET solution in the hybridisation chamber with 3× SSPET (see Note 15) and incubate at room temperature for 1 min. 4. Repeat step 3 using 0.5× SSPET, PBST, and PBS solutions sequentially. 5. Repeat a final wash with PBS solution, as described in step 3 above. Leave the solution in the hybridisation chamber until ready for the microarray imaging step. Cover the port apertures with PCR plate sealing tape. 6. Proceed with steps 1 and 2 of Subheading 3.4 (described above) for microarray imaging and data collection, respectively (see Fig. 10).

4. Notes 1. Once opened, all reagents purchased for the B3 have a limited shelf life hence care must be taken to ensure reagents are changed regularly. Recommended shelf life (in days) of reagents are as follows: Amidites (7 days), Activator (14 days), Cap A (14 days), Cap B (14 days), Oxidiser (14 days), Echem


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deblock (3 days), and Deblock T (60 days). Heavy use of the B3 will see the chemicals used well within the shelf life. However, with the exception of the Echem Deblock, all have been used up to 7 days past the recommended shelf life by this group, with successful synthesis. 2. For probe sequences shorter than 25 bases, it is useful to insert a ten-base thymidine spacer sequence at the 3′-end (e.g. 5′-…NNTTTTTTTTTT-3′) to allow enhanced physical access for hybridisation. The choice of thymidine as a spacer proves to have additional advantage since the degree of crosshybridisation onto the spacer region is minimised, due to weaker binding properties. 3. Any large areas of unassigned electrodes should be avoided. Thus, if any remainder electrodes are present on the micro array design window, these must be assigned with additional redundant probe sequences which will not be utilised in the study. 4. Not all the probes have to be present with the same number of replicates. Simply repeat the above procedure for different sets of probes, making sure you select the appropriate number of spots on the array each time. Select microarray>select random>“enter appropriate amount of spots”, or use the cursor and highlight the amount of spots required. 5. Due to the 72 h shelf life of the resultant solution, it is best to prepare a fresh stock on the day when actual microarray synthesis will be performed. 6. After completing the hydroquinone addition step, the user’s gloves have to be changed as prophylaxis against contamination by hydroquinone powder. 7. If the chips are required for experiments immediately, place the chips straight into water to keep the surface hydrated; allow to cool to room temperature before use. 8. The solution should be filter sterilised prior to use. 9. It is good practice to perform a laser scan of the microarrays prior to any hybridisation procedure, in order to obtain the background fluorescence level from the electrodes. The data collected should be used for post-hybridisation background subtraction (see Subheadings 3.4.1 and 3.4.2 for details). 10. For the 90K arrays, the focus depth must be adjusted for each individual chip. To do this, the recommended procedure is to use the 532 nm laser and while performing a high-resolution scan adjust the focus depth until the repeat electronic feature on the slide of the array comes into sharp focus. (N.B. Other scanners are compatible with the technology as long as they have the ability to change the scanning depth.)

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11. Following the first hybridisation of the microarray chip, make sure the chip is kept wet in 1× PBS or imaging solution. If the electrode surface dried out it cannot be reutilised. Once wet, the microarrays can be stored for up to 2 weeks – place in a refrigerator in order to minimise bacterial contamination of the PBS/imaging solution. 12. Ensure that no air bubbles are present in the hybridisation chamber. It is important to lay the microarray chip flat on a hard surface prior to filling the chamber with water. You will also find that the chamber might need slightly less than 100 mL volume for it to be filled completely. Ensure the chip is filled by letting a small drop of water exit from the upper port aperture. Consequently, wipe the excess droplet with fibre-free tissue and seal both portals by applying an adequately sized PCR plate sealing tape (any brand is acceptable although Abgene® PCR sealing tape can be removed more efficiently following the hybridisation procedure). 13. It is useful to introduce an adequately sized air bubble into the hybridisation chamber in order to aid homogenous distribution of the solution across the microarray surface. At this point, the microarray chip should be positioned and secured in the rotisserie oven in such a way as to allow the movement of air bubble in a vertical manner across the microarray surface during the hybridisation period (see Fig. 6). 14. Rinse the hybridisation chamber with 120 mL of the solution for the 90K microarray (100 mL for a 12K microarray), and replace with a fresh 120 mL aliquot. Introduce an air bubble into the chamber in order to improve the incubation efficacy. 15. Rinse the hybridisation chamber with 120 mL of the solution for a 90K microarray (100 mL for a 12K microarray), and replace with a fresh 120 mL aliquot.

Acknowledgments The authors would like to thank Drs. Brooke P. Anderson, Mike Lodes, and Jeremy Dumsday from Combimatrix Corporation for their assistance in the realisation of this chapter. References 1. Barken, K. B., Haagensen, J. A., and TolkerNielsen, T. (2007) Advances in nucleic acidbased diagnostics of bacterial infections. Clin Chim Acta 384, 1–11.

2. Virtanen, C., and Takahashi, M. (2008) Muscling in on microarrays. Appl Physiol Nutr Metab 33, 124–9.

Human Papilloma Virus Strain Detection 3. Bellazzi, R., and Zupan, B. (2007) Towards knowledge-based gene expression data mining. J Biomed Inform 40, 787–802. 4. Ault, K. A. (2008) Human papillomavirus vaccines: an update for gynecologists. Clin Obstet Gynecol 51, 527–32. 5. Castle, P. E. (2008) The potential utility of HPV genotyping in screening and clinical management. J Natl Compr Canc Netw 6, 83–95. 6. Liu, R. H., Lodes, M. J., Nguyen, T., Siuda, T., Slota, M., Fuji, H. S., and McShea, A. (2006) Validation of a fully integrated microfluidic array device for influenza A subtype identification and sequencing. Anal Chem 78, 4184–93. 7. Lodes, M. J., Suciu, D., Wilmoth, J. L., Ross, M., Munro, S., Dix, K., Bernards, K., Stöver, A. G., Quintana, M., Iihoshi, N., Lyon, W. J.,

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Danley, D. L., and McShea, A. (2007) Identification of upper respiratory tract pathogens using electrochemical detection on an oligonucleotide microarray. PLoS One 2, e924. 8. Liu, R. H., Nguyen, T., Schwarzkopf, K., Fuji, H. S., Petrova, A., Siuda, T., Peyvan, K., Bizak, M., Danley, D., and McShea, A. (2006) Fully integrated miniature device for automated gene expression DNA microarray processing. Anal Chem 78, 1980–6. 9. Ghindilis, A. L., Smith, M. W., Schwarzkopf, K. R., Roth, K. M., Peyvan, K., Munro, S. B., Lodes, M. J., Stöver, A. G, Bernards, K., Dill, K., and McShea, A. (2007) CombiMatrix oligonucleotide arrays: genotyping and gene expression assays employing electrochemical detection. Biosens Bioelectron 22, 1853–60.

Chapter 8 Multiplex Ligation-Dependent Probe Amplification (MLPA®) for the Detection of Copy Number Variation in Genomic Sequences Petra G.C. Eijk - Van Os and Jan P. Schouten Abstract Multiplex Ligation-dependent Probe Amplification (MLPA®) is a high-throughput method developed to determine the copy number of up to 50 genomic DNA sequences in a single multiplex PCR-based reaction. MLPA is easy to perform, requires only 20 ng of sample DNA and can distinguish sequences differing in only a single nucleotide. The MLPA reaction results in a mixture of amplification fragments ranging between 100 and 500 nt in length which can be separated and quantified by capillary electrophoresis. The equipment necessary for MLPA is identical to that for performing standard sequencing reactions: a thermocycler and a fluorescent capillary electrophoresis system. Comparison of the peak pattern obtained on a DNA sample to that of a reference sample indicates which sequences show aberrant copy numbers. Fundamental for the MLPA technique is that it is not the sample DNA that is amplified during the PCR reaction, but MLPA probes that hybridise to the sample DNA. Each MLPA probe consists of two probe oligonucleotides, which should hybridise adjacent to the target DNA for a successful ligation. Only ligated probes can be exponentially amplified by PCR. In contrast to standard multiplex PCR, only one pair of PCR primers is used for the MLPA PCR reaction, resulting in a more robust system. This way, the relative number of fragments present after the PCR reaction depends on the relative amount of the target sequence present in a DNA sample. Key words: Multiplex ligation-dependent probe amplification, Copy number detection, Multiplex PCR

1. Introduction Multiplex Ligation-dependent Probe Amplification (MLPA®) is a multiplex PCR method used primarily to detect small copy number changes of up to 50 different genomic DNA sequences in one simple reaction. Using MLPA, sequences differing in only one


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nucleotide can be distinguished (1). The technique is easy to use and can be performed in most laboratories, as it only requires a thermocycler and capillary sequencing equipment. Up to 96 samples can be handled simultaneously, and results can be obtained within 20 h. SALSA MLPA kits for more than 300 different applications are commercially available. For each application, the same MLPA reagents and reaction conditions are used; only the MLPA probes differ per application. At present, a major application of MLPA is the detection of aberrant copy numbers of genomic DNA sequences causing hereditary diseases. For most hereditary diseases, complete or partial gene deletions and duplications account for less than 10% of all disease-causing mutations and are thus a less frequent causative factor than point mutations. However, for many other diseases, this can be 10–30% (2–8) or even higher (9, 10). Heterozygous deletions and duplications of one or more exons are usually not detected by sequencing or other techniques used to detect point mutations, as one normal gene copy is still present. Other applications of MLPA include the detection and detailed characterisation of microdeletion syndromes, subtelomeric deletions and duplications, and the analysis of complex genomic regions in which the presence of related (pseudo) genes complicate analysis. The basis of the MLPA technique is that, instead of amplifying target sequences, the MLPA probes that are bound to their target sequences are amplified. Each MLPA probe consist of two probe oligonucleotides, which should hybridise adjacent to the target DNA for a successful ligation. Only ligated probes can be exponentially amplified by PCR. Whereas conventional multiplex PCR requires one primer pair for each fragment to be amplified, all fragments in MLPA reactions are amplified with the use of only one PCR primer pair. Using a single primer pair for all target sequences eliminates problems with unequal amplification efficiency caused by multiple primer pairs in an ordinary multiplex PCR. The MLPA procedure can be divided into seven interrelated steps: (1) Experimental set-up, (2) Sample treatment, (3) MLPA reaction, (4) Fragment separation, (5) Raw data evaluation & Peak pattern evaluation, (6) Data analysis, and (7) Result interpretation. In case, earlier steps are not performed properly, later steps may fail. More detailed information about the MLPA procedure can be found on http://www.mlpa.com. 1.1. Experimental Set-Up

Reference DNA samples of healthy individuals are in general essential to generate reliable MLPA results. As slight differences between different experiments may affect the MLPA peak pattern, it is necessary to include the reference samples in each MLPA experiment. Furthermore, inclusion of a no-DNA control reaction and positive control samples is recommended.

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A no-DNA control reaction should be included in each experiment, as it will reveal contamination of dH2O, MLPA reagents, electrophoresis reagents, or capillaries. The four Q-fragments at lengths of 64–70–76–82 nt that are included in all SALSA MLPA probemixes (MRC-Holland) will be prominent in the no-DNA control reaction. This is because the Q-control fragments, in contrast to the MLPA probes, do not require ligation for exponential amplification during the PCR reaction. Positive control samples are samples with a known deletion, duplication, or point mutation and can be used to check whether the MLPA procedure, including data analysis, has been performed well. Inclusion of positive control samples in each MLPA experiment is preferred, but not essential. 1.2. Sample Treatment

Although the MLPA reaction is robust and reproducible, the treatment of DNA (storage and extraction method) can influence the MLPA results. MLPA does not require a special method or kit for DNA extraction. However, it is strongly recommended to use reference and positive control samples that have been extracted by the same method and that are derived from the same type of tissue as the test samples, to minimise non-biological differences and variation. As the effect of contaminants is generally consistent and reproducible, this effect is corrected when the samples have undergone identical treatment. In case, pure DNA samples without any contaminants are used, MLPA results are independent of the DNA concentration used within the range of 20–500 ng. However, as many DNA samples do contain contaminants such as salt or ethanol, it is recommended to use a similar DNA concentration for each MLPA reaction to minimise variation caused by these contaminants.

1.3. MLPA Reaction

The MLPA protocol consists of three parts: (a) DNA denaturation and hybridisation of MLPA probes, (b) Ligation reaction, and (c) PCR reaction. See Fig. 1. for the MLPA protocol in a nutshell. During part (a), the sample DNA is denatured by heating to 98°C and incubated overnight at 60°C with a mixture of MLPA probes and salt. Every MLPA probe consists of two 1.

DNA denaturation: heat 5 minutes at 98°C.

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Hybridization: add SALSA probemix and MLPA buffer. Incubate 1 minute at 95°C, hybridize for 16 hr at 60°C.

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Ligation: add ligase mix and incubate 15 minutes at 54°C. Heat inactivate the ligase for 5 minutes at 98°C.

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PCR: add primers, dNTPs and polymerase. Start PCR.

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Capillary electrophoresis: export fragment lengths and peak areas to analysis software or spreadsheet. Analyze results.

Fig. 1. The MLPA protocol in a nutshell.

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Fig. 2. Outline of the MLPA technique. 1. The sample DNA is denatured and incubated with a mixture of MLPA probes. Each probe consists of two oligonucleotides: the Left Probe Oligonucleotide (LPO) and the 5′ phosphorylated Right Probe Oligonucleotide (RPO) which hybridize to directly adjacent target sequences. 2. Probe oligonucleotides hybridised to their adjacent targets are ligated. 3. Each probe has a ligation product of a unique length which is amplified exponentially by PCR, using a single primer pair which recognises the PCR primer sequences included in the probe oligonucleotides. 4. The resulting peak pattern of the sample is analysed by comparing it to that of the reference samples.

separate oligonucleotides, each containing one of the PCR primer sequences. The two probe oligonucleotides hybridise to immediately adjacent target sequences (Fig. 2 – step 1). The concentration of probes present in an MLPA reaction and the duration of the hybridisation reaction are sufficient to completely cover all DNA target sites with probes. In a typical MLPA reaction, approximately 500,000,000 copies of each probe oligonucleotide are present, while there are only 20,000 copies of most target sequences in a 60 ng human DNA sample.


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As the hybridisation reaction reaches completion, small changes in temperature, duration of the hybridisation reaction, or probe concentration will not influence the results obtained. Only when both probe oligonucleotides are hybridised to their adjacent target, they can be enzymatically ligated during part (b) the ligation reaction (Fig. 2 – step 2). Note that the ligation of the two probe oligonucleotides can be prohibited by a single nucleotide difference at the ligation site, making it possible to detect known point mutations or to distinguish closely related sequences. Ligation is more than 90% complete after 2 min, but it is recommended to perform the ligation reaction for 15 min. The number of ligated probes depends directly on the number of target sequences in the sample. When no target sequence is present, the corresponding probe will not be ligated and hence no probe signal will be detected. Only ligated probes will contain both PCR primer sequences and will be exponentially amplified during part (c) the PCR reaction (Fig. 2 – step 3). The removal of unbound probe oligonucleotides, containing only one primer sequence, is therefore not necessary. In the MLPA PCR reaction, a single primer pair is used, of which only the forward primer is fluorescently labelled. The PCR conditions are identical for different applications, as the same PCR primer pair is used for all SALSA MLPA kits produced by MRC-Holland. Since the PCR reaction goes to completion and stops when the majority of the PCR primers have been consumed, only minor differences are observed between 32, 35 (recommended), or 40 PCR cycles. 1.4. Fragment Separation

Each MLPA probe is designed in such a way that its amplification product has a unique length in an MLPA probemix. The probe length increases in a stepwise fashion by 6–9 nt, with a total size range between 100 and 500 nt. The PCR amplification products are mixed with molecular weight markers and formamide, denatured by a brief heating step and then separated by capillary electrophoresis (Fig. 2 – step 4). Optimal capillary fragment separation settings differ between instruments. For a proper fragment separation, the initial instrument settings for fragment separation should be optimised.

1.5. Raw Data and Peak Pattern Evaluation

Raw data from the sequencer should be evaluated prior to analysis, and based on this evaluation, it may be necessary to run the fragments again. For example, a rerun with a lower injection voltage or a shorter injection time can be required when one or more peaks exceed the maximum fluorescence intensity of the sequencer. In case, the raw data evaluation is positive, you may proceed with peak pattern evaluation. Peak pattern evaluation is performed to locate possible failures of the MLPA reaction. This is done by visual examination of

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the electropherograms of the raw, size-called data. For example, the peak pattern of the MLPA probes and the size standard will both be affected by the use of deteriorated gel, old capillaries, or low-quality formamide. Only data passing the raw data and peak pattern evaluation should be used for the actual MLPA data analysis. 1.6. Data Analysis

The MLPA data analysis is facilitated by the aforementioned fact that it is not the target sequences that are amplified and quantified, but the ligated MLPA probes. One of the advantages of this approach is that the amplification products always have the same length, regardless of possible sequence variants in the probe target sequence in the sample DNA. In order to compensate for differences between different capillaries and pipetting errors, MLPA analysis starts with intra-sample normalisation. In other words, only relative peak areas are used, never absolute peak areas. Intra-sample normalisation is commonly done by using reference probes, which are included in most SALSA MLPA probemixes. Reference probes detect chromosomal locations that are more different than the target-specific probes, and which are assumed to have the same (usually diploid) copy number in all analysed samples. After the intra-sample normalisation, the probe ratios of a sample should always be compared to the corresponding probe ratios in the reference samples (inter-sample normalisation). The resulting ratio reflects the relative copy number of the probe’s target sequence in the analysed sample. A heterozygous deletion of an autosomal sequence, for example, is identified by a relative ratio of approximately 0.5: one copy per cell instead of two. Such deletion can usually be seen during visual examination of the electropherogram (Fig. 3).

1.7. Result Interpretation

Usually, probe ratios below 0.7 or above 1.3 are regarded as indicative of a heterozygous deletion (copy number change from two to one allele) or duplication (copy number change from two to three alleles), respectively. To determine whether the produced ratios are reliable, sufficient knowledge about the MLPA technique and the studied application is essential. Arranging probes according to chromosomal location facilitates interpretation of the results and may reveal more subtle changes such as those observed in mosaic cases. It is important to realise that, next to a true copy number change, a reduced probe signal can also be due to a change in the sequence detected by that probe. Sequence changes immediately adjacent to the ligation site (SNPs, point mutations) can influence probe signals by preventing ligation of the two probe oligonucleotides. Sequence changes at further distance, even as far as 15–20 nt from the ligation site, can also result in a reduced probe


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Fig. 3. Detection of copy number changes by MLPA with SALSA MLPA kit P245 Microdeletion syndromes. MLPA easily allows for the distinction between normal reference sample (b) and a sample (a) carrying heterozygous deletion of the 15q Prader Willi/Angelman region. The four probes for this region show an approximately 50% reduced signal. The other probes are on other chromosomes and remain unchanged.

signal when the mismatch between sample DNA and probe oligonucleotide destabilises binding of the probe. Copy number changes detected by only a single probe will, therefore, always require confirmation by other methods. Sequencing is often used to determine whether a mutation or polymorphism in the probe binding site is present. Long-Range PCR and qPCR are often used to confirm (single) exon deletions detected by MLPA. In addition, it is important to realise that not all deletions and duplications detected by MLPA will be pathogenic. A partial gene duplication within the gene may disrupt that copy of the gene resulting in a disease, while a duplication of the complete gene at another chromosomal location might not be pathogenic. Users of MLPA should always verify the latest scientific literature when interpreting their findings.

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1.8. Synthetic MLPA Probes for Research

Although performing an MLPA reaction is easy, the development of new assays is complex and time-consuming. SALSA MLPA kits for many different applications are commercially available from MRC-Holland, the Netherlands (http://www.mlpa.com). MRCHolland produces MLPA probes that are derived from M13phages which makes it possible to generate longer MLPA probes as compared to chemically synthesised oligonucleotides. This allows combining up to 50 probes in a single probemix. It is, however, possible for each laboratory to design smaller sets of chemically synthesised MLPA probes for research purposes. A detailed protocol on how to design synthetic probes is available on http://www.mlpa.com.

1.9. RT-MLPA and MS-MLPA

Several variations of MLPA have been developed. RT-MLPA (reverse transcriptase MLPA) can be used for mRNA profiling (11). As RT-MLPA probes cannot detect mRNAs directly, the RT-MLPA procedure starts with a reverse transcriptase reaction to synthesise complementary DNA (cDNA) from the mRNA template. After this, RT-MLPA is continued as ordinary MLPA. Another variation is methylation-specific MLPA (MS-MLPA). Next to the detection of copy number variation, MS-MLPA can be used for the identification of methylation changes (12) and has been proven to be a very useful technique for the detection of imprinting diseases (13–15) and for tumour sample analysis (16, 17). MS-MLPA has several advantages compared to other techniques for DNA methylation profiling: (1) MS-MLPA is simple to perform; (2) No bisulphite conversion of sample DNA is required; (3) methylation and copy number changes of numerous genes can be analysed simultaneously; (4) DNA isolated from paraffin-embedded tissue can be used for analysis. MS-MLPA differs from the standard MLPA technique in only a few aspects, the main difference being that each MS-MLPA reaction generates two samples for capillary electrophoresis analysis. Following the hybridisation, the reaction is divided into two tubes. One tube is processed as a standard MLPA reaction and provides information on DNA copy number changes only (not shown in Fig. 4). The other tube is incubated with the methylationsensitive HhaI endonuclease while the hybridised probes are simultaneously ligated. Hybrids of probes and unmethylated sample DNA are digested by the HhaI enzyme (Fig. 4 – step 2). These digested probes cannot be exponentially amplified by PCR and hence will not generate a signal when analysed by capillary electrophoresis. In contrast, if the sample DNA is methylated, the methylated probe–sample DNA hybrids are protected from digestion by HhaI. In this case, ligated probes will generate a signal (Fig. 4 – steps 3 and 4). The analysis of MS-MLPA results consists of two steps: (1) determining copy numbers by comparing the undigested reaction of each sample with that of reference samples (identical to standard


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Fig. 4. Outline of the MS-MLPA technique. 1. The sample DNA is denatured and incubated with a mixture of MLPA probes. Each probe consists of two oligonucleotides, which hybridise to directly adjacent target sequences. MS-MLPA probes contain a recognition site of a methylation-sensitive restriction endonuclease (e.g., HhaI or HpaII), reference probes do not. 2. The reaction is split into two tubes. One tube is processed as a standard MLPA (only ligation; not shown), and the reaction in the other tube (shown here) is ligated and digested with the methylation-sensitive restriction enzyme HhaI (ligation + digestion). Hybrids of probes with unmethylated sample DNA are digested. 3. Each non-digested probe has a ligation product of a unique length which is amplified exponentially by PCR. 4. For each sample, two peak patterns are produced: one from the ligation-only reaction (for determining copy number changes – not shown), and one from the ligation and digestion reaction (for methylation profiling – shown). The peak patterns are analysed by comparing them to those obtained on reference samples. Left peak pattern : ligation and digestion reaction of a healthy control showing only the 13 reference probes not containing a HhaI restriction site. The other 26 probes all detect a sequence containing a HhaI restriction site which is unmethylated in normal bloodderived DNA. The probe-sample hybrids are thus digested, and hence, the MS-MLPA probes are not visible in this sample. Right peak pattern : ligation and digestion reaction of a tumour sample. The extra four peaks visible here are due to the fact that some of the sequences detected were methylated in this tumour-derived DNA, protecting the probe-DNA hybrid from endonuclease digestion.

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MLPA) and (2) determining methylation patterns by comparing each undigested sample to its digested counterpart. Additional information about the MS-MLPA materials, reaction, and data analysis can be found in the sections below.

2. Materials 2.1. Sample Treatment

2.2. MLPA Reaction 2.2.1. DNA Denaturation and Hybridisation of MLPA Probes

2.2.2. Ligation Reaction

MLPA does not require a special method or kit for DNA extraction. Good results have been obtained with salting out protocols, phenol/chloroform extraction, and kits from various commercial suppliers (see Notes 1–3). It is strongly recommended to use within a single experiment only samples that have been extracted by the same method. 1. SALSA MLPA probemix (MRC-Holland, http://www.mlpa. com): A mixture of up to 50 SALSA MLPA probes and several control fragments are present, such as the four Q (quantity) fragments, stable for at least 1 year when stored at −20°C. 2. SALSA MLPA buffer (MRC-Holland, http://www.mlpa. com): Contains 1.5 M salt, buffer and additives; stable for at least 1 year when stored at −20°C; does not always freeze at −20°C, rather viscous and does not mix easily. 1. Ligase-65 enzyme solution (MRC-Holland, http://www. mlpa.com): Stable for at least 1 year when stored at −20°C. Solution in 50% glycerol which will not freeze at −20°C. 2. Ligase-65 buffer A (MRC-Holland, http://www.mlpa.com): Contains coenzyme NAD, which is required by the Ligase-65 enzyme, stable for at least 1 year when stored at −20°C, sensitive to repeated freezing/thawing and may deteriorate after more than 20 freeze/thaw cycles. 3. Ligase-65 buffer B (MRC-Holland, http://www.mlpa.com): Contains the salts required by the Ligase-65 enzyme, stable for at least 1 year when stored at −20°C.

2.2.3. PCR Reaction

1. SALSA PCR buffer (MRC-Holland, http://www.mlpa.com): Stable for at least 1 year when stored at −20°C. 2. SALSA PCR primers (MRC-Holland, http://www.mlpa. com): Contains one fluorescently labelled and one unlabelled PCR primer and dNTPs, stable for at least 1 year when stored at −20°C, light sensitive. SALSA PCR Forward primer (Labelled): 5′*GGGTTCCCT AAGGGTTGGA-3′. SALSA PCR Reverse primer (Unlabelled): 5′-GTGCCAGC AAG ATCCAATCTAGA-3′.


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3. SALSA Polymerase enzyme solution (MRC-Holland, http:// www.mlpa.com): Similar to Taq polymerase, stable for at least 1 year when stored at −20°C, solution in 50% glycerol which will not freeze at −20°C. 4. SALSA Enzyme dilution buffer (MRC-Holland, http://www. mlpa.com): Stabilises the polymerase/primers dilution, stable for at least 1 year when stored at −20°C. 2.2.4. Modification MLPA Reaction for MS-MLPA: Ligation and Digestion Reaction

2.3. Fragment Separation

1. Materials of the standard MLPA ligation reaction. 2. HhaI restriction endonuclease (Promega, 10 units/ml).

1. High-resolution electrophoresis system with fluorescent detection, e.g., instruments used for DNA sequencing, such as Applied Biosystems and Beckman capillary sequencers. 2. High-quality formamide (e.g. Hi-Di Formamide, Applied Biosystems). 3. Fluorescently labelled DNA size standard. For Applied Biosystems sequencers, the fluorescent label of the size standard depends on the installed filter set: LIZ-500 (preferred), or ROX-500, or TAMRA-500. For the Beckman CEQ sequencers, use the D1-600 marker. 4. Fragment analysis software, e.g. Coffalyser (MRC-Holland), Genemapper (AB), CEQ Fragment analysis (Beckman), Fragment profiler (Megabase).


1. Fragment analysis software, e.g., Coffalyser (MRC-Holland), Genemapper/Genescan (AB), CEQ Fragment analysis (Beckman), Fragment profiler (Megabase). 2. Raw data files from sequencer, e.g. .fsa (AB); .scf, .cqf, or .esd (Beckman); .rsd (Megabase). 3. Raw data evaluation checklist (see Fig. 5 and http://www. mlpa.com). 4. Peak pattern evaluation flow chart (see Fig. 6 and http:// www.mlpa.com).

2.5. Data Analysis

Various tools for MLPA data analysis are available. For more information, see http://www.mlpa.com.

2.5.1. Coffalyser MLPA Software

Coffalyser software, developed at MRC-Holland, is recommended for the analysis of MLPA data. All data normalisation steps are built-in functions of Coffalyser. Furthermore, Coffalyser corrects for systematic probe bias and size to signal drop (sloping), which often occurs in capillary sequencers. Coffalyser software can be downloaded free (http://www.mlpa.com) and is updated regularly.

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Eijk - Van Os and Schouten Raw data evaluation - checklist

Yes

No

1. Do you see a high baseline? 2. Do you see high background signals and/or shoulderpeaks? 3. Is the size standard pattern unusual, e.g. different heights, sloping, broad peaks? 4. Are all peaks very low? Lower than 5 % of the fluorescence maximum of the sequencer? 5. Is there too much signal sloping? 6. Are spiky peaks present? 7. Are high peaks above the fluorescence maximum of the sequencer present? 8. Have the peaks of the longer MLPA probes a much broader base than of the shorter fragments? 9. Do you see spectral pull up/pull down patterns in the colours? 10. Do you see irregular current patterns or decreasing current?

Fig. 5. Raw data evaluation checklist. You can proceed with peak pattern evaluation when all questions above are answered with NO. In case, one or more questions have been answered with YES, see the guidelines on “http://www. mlpa.com – MLPA procedure – Raw data evaluation” on how you can recognize problems and how to improve fragment separation.

3. Methods 3.1. Experimental Set Up

1. Select test and reference samples that have been extracted by the same method and that preferably have been derived from the same type of tissue (see Note 1). 2. Use at least three reference samples in each MLPA run. When using more than 21 samples, add 1 additional reference sample for each 7 samples. Reference samples should be spread randomly over the sample plate to minimise variation (see Note 4). 3. Include a no-DNA control reaction. This is an MLPA reaction on H2O or TE instead of DNA (see Note 5). 4. When available, inclusion of positive control samples is recommended, but not essential. It is possible to use DNA from cell lines as positive control sample, but take into account that cell lines may have acquired additional copy number changes, including gains of complete chromosomes.

3.2. Sample Treatment

1. Extract the DNA from a blood sample or other tissues. MLPA does not require a special method or kit for DNA extraction (see Notes 1 and 2). 2. Dissolve the DNA in TE (10 mM Tris–HCl pH 8.2 + 0.1 mM EDTA). The pH of the DNA preparation should be around 8.0 or 8.5 to prevent depurination during the initial heat treatment at 98°C.


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Peak pattern evaluation flow chart Go back to Sample treatment, MLPA reaction or Fragment separation (see www.mlpa.com > MLPA procedure). 1. Control fragment 92 nt + probe amplification products present? (see note 20)

Primer-peak or primer dimers visible (20-60 nt region)?

NO

YES All Q-fragments visible? (6470-76-82 nt) (see note 22)

YES 2. Peak signals of Q-fragments (64, 70, 76 & 82 nt): larger than half the signal of the ligation-dependent peak (92 nt)? (see note 22)

NO 4. Extra peaks visible in no DNA control reaction? (see notes 5, 37 )

YES

NO

1. Check fluorescent label (see note 21). 2. Check size standard peaks. If absent, repeat electrophoresis. 1. PCR failed. Repeat PCR using same ligation reaction (see note 15). 2. Probemix was not added. 1. No DNA present (see note 23). 2. Ligation reaction failed (see note 24).

Amount of DNA too low (see note 23). Repeat MLPA reaction with more DNA.

YES Peak signals of D-fragments (88 YES and/or 96 nt) smaller than 1/3 of the signal of the 92 nt control fragment? (see note 27)

NO 5. Any extra peaks visible?

NO

YES

NO 3. Unexplainable loss of some signals and increase of others?

NO

NO

Incomplete denaturation (see note 27). 1. Use less sample DNA when possible; this reduces the contaminants which may cause denaturation problems. 2. And/or include 5% glycerol in the 5 µl DNA sample. 3. Or prolong 5 min 98oC sample step to 40 min.

1. Incomplete hybridization: check heated lid and thermocycler settings (see note 25). 2. Overloading capillaries in some apparatus (e.g.ABI3700) may result in signals in neighbouring capillaries. Use less PCR product or reduce injection time (see note 33). 3. Denaturation of injection mixture before the fragment separation was omitted (see note 26). YES

6. Excessive primer-dimer formation visible? (seenote 28) NO 7. Split peaks visible in most MLPA probe signals? (see note 28)

Prepare Polymerase mix on ice and divide over a strip to permit multichannel pipetting; use ice block to keep strips cold. Put a mix of ligation product and PCR buffer in the thermocycler and heat to 60°C. At 60°C, add polymerase mix using a multichannel pipette. Start PCR immediately (see note 30).

YES

1. DNA may have been degraded. Rerun MLPA amplification products using high quality mineral oil and formamide. 2. Denaturation of injection mixture was omitted see (note 26). 2. High salt concentration in the injection mixture may cause secondary structure. Do not use more PCR product than ~1/10 of the total volume of the injection mixture.

YES

NO

MLPA and size standard peaks low and broad? 8. All peak signals too low? (lower than 5% of maximum detectable signal) (see note 28).

YES

Replace capillaries or gel (see note 29). YES 1. Increase capillary injection time or injection voltage (see note 33). If this doesn’t help: 2. Repeat PCR, using 40 cycles; reduce time between addition of polymerase mix and start of PCR. if this doesn’t help: 3. Repeat capillary electrophoresis and increase amount of PCR product.

NO

NO 9. One or more peak signals too high (off-scale)?

Repeat capillary electrophoresis (see note 33): decrease amount of PCR product injected (see note 31) decrease capillary injection time and/or voltage

YES

NO 10. Check for sloping: are peaks YES of long MLPA probes more than 3 times lower than those of shorter MLPA probes? (see note 35) NO 11. Are there large differences in relative peak areas between different samples that do not make sense? (see note 28) NO

YES

YES Does the size standard (ROX, TAMRA, LIZ, D1) show the same sloping? NO

Check run current: this should be a flat line; if not, check separation buffer & injection mixture for air bubbles. Formamide (see note 32), gel (see note 29) or capillaries may need to be replaced. Capillaries overloaded; try using less PCR product. Rerun MLPA products after diluting the MLPA products

1. Test the evaporation during the hybridization reaction (see note 34). 2. DNA samples may contain substances that inhibit the PCR reaction (see note 36). 3. For extensive information see www.mlpa.com > MLPA procedure > Raw data evaluation 1. 2. 3. 4.

Cause Cause Cause Cause

1: 2: 3: 4:

impurities in the DNA sample (see note 36). use of extremely large amounts DNA (old/evaporated/viscous samples). overloading of capillaries has saturated the fluorescent detection device (see note33). peak broadening due to deteriorated gel or capillaries (see note 29).

Continue with data analysis(see www.mlpa.com > MLPA procedure > Data analysis) .

Fig. 6. Peak pattern evaluation flow chart. Only data passing the raw data evaluation and peak pattern evaluation can be used for data analysis.

3. Aliquot reference/control samples and store the DNA at −20°C. DNA is very stable when dissolved in TE, but storage at 4°C for several years is not recommended. Contamination with micro-organisms and/or moulds can deteriorate your samples. 4. In a pre-PCR location, measure DNA stock concentrations if necessary.

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5. If needed, dilute the DNA in TE (10 mM Tris–HCl pH 8.2 + 0.1 mM EDTA) to working stock of 10–20 ng/ml tubes (see Note 3). 3.3. MLPA Reaction

1. In a pre-PCR location, label 0.2 ml tubes (strips or plates).

3.3.1. DNA Denaturation and Hybridisation (Day 1)

2. Add 5 ml of DNA working stock to each tube (or TE/H2O as a no-DNA control reaction) (see Note 3). 3. If needed, spin the strips or tubes briefly in a centrifuge. 4. Place the tubes in a thermocycler and start the MLPA programme (see Notes 6 and 7): denature sample DNA for 5 min at 98°C. Samples should be cooled down to 25°C before opening the thermocycler. 5. Thaw probemix and MLPA buffer. Always vortex thawed buffers and probemix briefly before use (see Note 8). 6. Prepare a hybridisation master mix, add for each reaction: 1.5 ml MLPA buffer + 1.5 ml probemix. Mix well by pipetting or vortexing (see Notes 8 and 11). 7. When the thermocycler reaches 25°C, add this 3 ml hybridisation master mix to each sample tube. Mix well by pipetting up and down. 8. Continue the thermocycler programme: incubate for 1 min at 95°C, then for 16 h at 60°C (see Notes 6, 7, and 9).

3.3.2. Ligation Reaction (Day 2)

1. In a pre-PCR location, thaw Ligase-65 buffer A and Ligase-65 buffer B. Vortex briefly. 2. Prepare a Ligase-65 master mix, add for each reaction: 3 ml Ligase-65 buffer A + 3 ml Ligase-65 buffer B + 25 ml dH2O. Mix by vortexing (see Notes 8, 10, and 11). 3. Add for each reaction 1 ml of Ligase-65 to the Ligase-65 master mix and mix well by pipetting up and down (see Note 12). 4. Continue the thermocycler programme: hold at 54°C (see Note 6). 5. When the samples are at 54°C, add 32.0 ml of the Ligase-65 master mix to each reaction tube and mix by pipetting up and down (see Notes 13 and 14). 6. Continue the thermocycler programme: 15 min incubation at 54°C (for ligation), followed by 5 min at 98°C for heat inactivation of the ligase enzyme and then hold at 15°C (see Note 6, 15).

3.3.3. PCR Reaction (Day 2)

1. In a pre-PCR location, thaw SALSA PCR buffer, SALSA PCR-primers, and SALSA Enzyme Dilution buffer. Vortex briefly (see Notes 8, 16, and 17). 2. Label new tubes for the PCR reaction.


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3. Prepare a PCR buffer mix, add for each reaction: 4 ml SALSA PCR buffer + 26 ml dH2O. Mix briefly by vortexing (see Notes 11 and 16). 4. Add 30 ml of the PCR buffer mix to each new tube. 5. At room temperature, transfer 10 ml of each ligation product to its corresponding PCR tube. 6. If needed, spin the strip tubes briefly in a centrifuge with a rotor for 0.2 ml strip tubes. 7. Place tubes into the thermocycler. 8. Prepare a polymerase master mix on ice, add for each reaction: 2 ml SALSA primer + 2 ml SALSA enzyme dilution buffer + 5.5 ml dH2O. Mix well by pipetting up and down (see Note 10 and 11). 9. Add 0.5 ml of SALSA Polymerase for each reaction to this polymerase master mix and mix well by pipetting gently up and down. Do not vortex (see Note 12). 10. Continue the thermocycler programme: hold at 60°C (see Note 6) and place the PCR tubes in the thermocycler. 11. Add 10 ml of polymerase master mix to each PCR tube while in the thermocycler at 60°C. Mix by pipetting gently up and down (see Note 13). 12. Close the tubes and continue the thermocycler programme immediately after the addition of the polymerase master mix: 35 cycles: 30 s 95°C; 30 s 60°C; 60 s 72°C. End with 20 min incubation at 72°C and then hold at 15°C (see Notes 6, 16, 17, 18, and 19). 3.3.4. Modification MLPA Reaction for MS-MLPA: Ligation and Digestion Reaction

1. In a pre-PCR location, thaw Ligase-65 buffer A and Ligase-65 buffer B. Vortex briefly. 2. Dilute ligase buffer A, add for each reaction: 3 ml Ligase-65 buffer A + 10 ml dH2O. Mix by vortexing. 3. Add 13 ml of the ligase buffer A dilution to each tube. Mix well by pipetting up and down. 4. Label a second 0.2-ml tube for each sample with sample initials, probemix number, and the mark digestion. 5. Transfer 10 ml from the first tube to the second tube. 6. Prepare a Ligase-65 master mix in two steps. First, mix for each reaction: 1.5 ml Ligase-65 buffer B + 8.25 ml dH2O. Mix by vortexing (see Notes 8, 10, and 11). Second, add 0.25 ml of Ligase-65 for each reaction. Mix well by pipetting gently up and down (see Note 12). 7. Prepare a Ligase-Digestion master mix in two steps. First, mix for each reaction: 1.5 ml Ligase-65 buffer B + 7.75 ml dH2O. Mix by vortexing (see Notes 8, 10, and 11). Second, add for

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each reaction 0.25 ml Ligase-65 enzyme + 0.5 ml HhaI enzyme. Mix well by pipetting gently up and down (see Note 12). 8. Continue programme on the thermocycler: Incubate both tubes in the thermocycler at 49°C for at least 1 min (see Note 7). 9. While at 49°C, add 10 ml Ligase-65 mix to the first tube (copy number test). Mix well. 10. While at 49°C, add 10 ml Ligase-Digestion mix to the second tube (methylation test). Mix well. 11. Continue programme on the thermocycler: incubate both tubes for 30 min at 49°C, then heat for 5 min at 98°C and hold at 15°C (see Notes 7 and 14). 3.3.5. Modification MLPA Reaction for MS-MLPA: PCR Reaction

1. In a pre-PCR location, thaw SALSA PCR buffer, SALSA PCR-primers, and SALSA Enzyme Dilution buffer. Vortex briefly (see Notes 8, 16, and 17). 2. Prepare a PCR buffer mix, add for each reaction: 2 ml SALSA PCR buffer + 13 ml dH2O. Mix by vortexing (see Notes 11 and 16). 3. Label new tubes for PCR with same sample initials, probemix number and mention whether it contains digested or undigested sample. 4. Add 15 ml of the PCR buffer mix to each tube. 5. At room temperature, transfer 5 ml of each ligation reaction and ligation–digestion reaction to its corresponding PCR tube. 6. If needed, spin the strip tubes briefly in a centrifuge with a rotor for 0.2-ml strip tubes. 7. Place tubes into the thermocycler. 8. Prepare a polymerase master mix on ice, add for each reaction: 1 ml SALSA primers + 1 ml SALSA enzyme dilution buffer + 2.75 ml dH2O. Mix well by vortexing (see Notes 10 and 11). 9. Add 0.25 ml of SALSA Polymerase for each reaction to this polymerase master mix and mix well by pipetting gently up and down. Do not vortex (see Note 12). 10. Continue thermocycler programme to the 60°C hold step (see Note 7). 11. Add 5 ml of polymerase master mix to each PCR tube while in the thermocycler at 60°C. Mix by pipetting (see Note 13). 12. Continue the thermocycler programme and proceed to PCR: 35 cycles: 30 s 95°C; 30 s 60°C; 60 s 72°C. End with 20 min incubation at 72°C and then hold at 15°C (see Notes 7, 16, 17, 18, and 19).


3.4. Fragment Separation

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The amount of MLPA PCR reaction, the run conditions, and the fluorescent label required for analysis all depend on the type of capillary sequencer or gel electrophoresis instrument used. For a proper fragment separation, the initial instrument settings for fragment separation should be optimised empirically. The initial settings for different instrument types and instructions on how to optimise fragment separation can be found on http://www.mlpa.com. For an ABI-Prism 3100 (Avant) Genetic Analyzer, ABI-3130 & ABI-3130XL Genetic Analyzer: 1. Mix 0.7 ml of the PCR reaction + 0.3 ml ROX or 0.2 ml LIZ internal size standard + 9 ml HiDi formamide (ABI nr.4311320). Seal the injection plate with plate sealing film. 2. Incubate the mix for 2 min at 80°C. Hold at 4°C for 5 min. 3. Place the injection plate in your capillary device and run the appropriate run module. 4. Initial Injection voltage: 1.6 kV; Initial Injection time: 15 s. For other electrophoresis parameters, the default setting for fragment analysis can be used. 5. The use of POP4 polymer and 36-cm capillaries is preferred. 6. When using POP7 gel, decreasing your Run Voltage to 10 kV and Run Time to 2,000 s may improve peak shape and thereby calculation of peaks area.


1. Before size calling, evaluate the electropherograms of the raw data according to the raw data evaluation checklist (see Fig. 5). 2. Rerun the MLPA products if needed (see Subheading 3.4). 3. Evaluate your sized-called electropherogram according to the peak pattern flow chart below (see Fig. 6).

3.6. Data Analysis

Data that has passed the Raw data and Peak pattern evaluation can be used for data analysis. Normalisation of electrophoresis results is essential for obtaining useful MLPA data; therefore, data analysis forms a crucial component of the MLPA procedure. More information can be found on http://www.mlpa.com.

3.6.1. Data Normalisation

Many different methods exist for the normalisation of MLPA data, and it is impossible to single out one strategy that works best for all probemixes (see Notes 38 and 39). Each product description of a SALSA MLPA kit purchased from MRC-Holland describes the best method to normalise your data. The majority of SALSA MLPA probemixes can be normalised following the method described below (see Note 40). 1. Intra-sample normalisation: divide the peak area of each probe’s amplification product by either (a) the total peak area

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(or peak height; see Note 41) of only the reference probes (block normalisation, see Note 42) or (b) the combined peak areas of all probes (population normalisation). 2. Inter-sample normalisation: divide these intra-normalised probe ratios of a sample by the average intra-normalised probe ratio of all reference samples. This provides a probe ratio for each probe per sample. 3.6.2. Additional Information Data Analysis MS-MLPA

1. Intra-sample data normalisation (all samples): divide the signal of each probe by the signal of every reference probe in that sample, thus creating as many ratios per probe as there are reference probes. Subsequently, the median of all these produced ratios per probe should be taken; this is the probe’s Normalisation Constant. This Normalisation Constant can then be used for sample to reference sample comparison, both for copy number and digestion determination. 2. The Inter-sample normalisation of MS-MLPA kits consists of two parts: (1) determining copy numbers by comparing different undigested samples (all MLPA kits; see Subheading 3.6.1), and (2) determining methylation patterns by comparing each undigested sample to its digested counterpart (MS-MLPA kits only; see below). The second part is unique for MS-MLPA kits and serves to semi-quantify the percentage of methylation within a given sample. 3. Methylation status of MS-MLPA probes (see Note 43) is calculated by dividing (a) the Normalisation Constant of each MS-MLPA probe obtained on the digested patient sample by (b) the Normalisation Constant of each MS-MLPA probe obtained on the corresponding undigested sample. Multiplying this value by 100 gives an estimation of the percentage methylation. Aberrant methylation can then be identified by comparing the methylation status of one or more MS-MLPA probes in the sample in question to that obtained on reference samples.


1. Probe ratios below 0.7 are usually regarded as an indication of a heterozygous deletion (copy number change from two to one allele) and probe ratios above 1.3 of a duplication (copy number change from two to three alleles) (see Notes 44–46). 2. To determine whether the produced ratios are reliable, sufficient knowledge about the MLPA technique and the studied application is essential. A result is more plausible when: (a) The overall standard variation per probe is low.


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(b) Few or no abnormalities are found in a patient cohort for a disease in which copy numbers changes are known to be rare. (c) Probes for adjacent exons show a decreased or increased signal, e.g. exons 2–5, indicating a multi-exon deletion or duplication. (d) The same result is obtained in a new experiment using less (minimum 20 ng) DNA and/or using different reference samples. When less DNA is used, possible contaminants which may influence the probe signal are diluted. (e) A single exon deletion is found in a gene with large introns. The frequency of single exon deletions is, in general, higher in genes with large introns than in small genes with very short introns. 3. A result is unlikely to be reliable when: (a) Probes for non-neighbouring exons show a decreased or increased signal, e.g., exon 3 and 17. (b) In the same sample, one or more reference probes show an abnormal copy number. (c) Many abnormalities are found in a patient cohort of a disease in which copy numbers changes are known to be rare. 4. Confirmation of results (a) It is strongly recommended to confirm MLPA results by another method, in particular, when only one probe shows a reduced or increased signal. Point mutations and (non-pathogenic) polymorphisms, even when located at 20 nt distance from the probe ligation site, can result in a decreased probe signal by destabilisation the binding of the probe oligonucleotides to the sample DNA. Furthermore, the signal of some probes is more influenced than other probes by impurities in the DNA sample or small differences in procedure (e.g., evaporation). 3.7.1. Additional Information Result Interpretation MS-MLPA

1. Each probemix contains several MS-MLPA digestion control probes detecting a sequence that is unmethylated in most tissues. An absent signal for these probes indicates a complete digestion by the restriction endonuclease. 2. Methylation changes can be identified by the appearance of a peak after HhaI digestion that is absent in the digested reference DNA (promoter regions of tumour suppressor genes) or by a change from 0.5 in 1.0 or zero in case of imprinted regions.

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4. Notes 1. In case, MLPA reactions are performed on old samples, or on samples that have been extracted using different methods, an increased probe variation is often observed. An extra purification step may help to produce more reliable results. Samples used to study methylation with MS-MLPA should always be derived from the same tissue, as methylation can vary between different types of tissue. Old viscous (partly evaporated) samples should be sufficiently diluted. 2. Substantial amounts of salt are present in DNA samples purified by the default program of the Qiagen EZ1 BioRobot, its predecessor Geno-M6, as well as by the Qiagen M48 and M96 systems. This high salt concentration prevents complete denaturation of the CG-rich parts of the sample DNA (CpG islands), thereby compromising MLPA and other methods. Incomplete DNA denaturation can cause false positive MLPA results. The EZ1 system has an alternative wash procedure which provides a good-quality sample DNA without salt. 3. It is recommended to use 50–100 ng in 5 ml of human chromosomal DNA, although an acceptable range is 20–500 ng DNA in 5 ml. If the stock concentration is less than 10 ng/ml, use the available concentration; please note that results may be less reliable when less than 20 ng in 5 ml sample DNA is used. The DNA amount can be considered sufficient when the average signal of the four Q-fragments is lower than onethird of the probe peak signals. The EDTA concentration in the DNA sample should not exceed 1 mM. Concentration of samples by evaporation/Speedvac may result in high EDTA concentrations that affect the MLPA reaction. The sample volume should not exceed 5 ml. The volume of the reaction is important for the hybridisation speed, which is mainly influenced by the probe and salt concentration. 4. Reference samples might not be essential when a large number (>20) of independent samples (different families) are run at the same time, AND the chance for each individual probe to be deleted or duplicated in a sample is low ( MLPA procedure > Troubleshooting. 6. Thermocycler program for the complete MLPA procedure: (a) Denaturation and hybridisation 1. 98°C

5 min

2. 25°C

Hold

3. 95°C

1 min

4. 60°C

Hold

(b) Ligation reaction 5. 54°C

Hold

6. 54°C

15 min

7. 98°C

5 min

8. 15°C

Hold

(c) PCR reaction 9. 60°C

Hold

10. 35 cycles 95°C 60°C 72°C

30 s 30 s 60 s

11. 72°C

20 min

12. 15°C

Hold

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7. Modified thermocycler programme MS-MLPA procedure: (a) Denaturation and hybridisation 13. 98°C

5 min

14. 25°C

Hold

15. 95°C

1 min

16. 60°C

Hold

for

the

complete

(b) Ligation and digestion reaction 17. 49°C

Hold

18. 49°C

30 min

19. 98°C

5 min

20. 15°C

Hold

(c) PCR reaction 21. 72°C

Hold

22. 35 cycles 95°C 60°C 72°C

30 s 30 s 60 s

23. 72°C

20 min

24. 15°C

Hold

8. SALSA MLPA probemix and buffers can be mixed by vortexing or repeated pipetting at room temperature before use. The MLPA buffer is viscous and does not mix easily. Please note that all solutions that have been frozen are not homogeneous after thawing! During the freezing process, all salts, including oligonucleotides, are excluded by the freezing water and are concentrated at the bottom of the tube. 9. The hybridisation time should be between 12 and 24 h for reproducible results. We recommend 16–20 h, but hybridisation of probes to their targets is nearly complete after 12 h. In case, the hybridisation reaction is interrupted due to power failure, just place the tubes again at 60°C and continue until the total hybridisation time is between 16 and 24 h. 10. Ligase-65 master mix and polymerase master mix should be made less than 1 h before use and stored on ice. All reagents should be returned to −20°C after use.


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11. Prepare master mix and buffer mix for all reactions within an experiment to minimise sample-to-sample variation. Prepare 5–10% extra to allow for pipetting errors. 12. Enzyme solutions are supplied in 50% glycerol and remain fluid at −20°C. Enzyme solutions should never be vortexed but should be mixed with water and buffers by repeated (gentle) pipetting up and down. 13. When performing MLPA on a large number of samples, multi-channel pipettes are recommended. In particular, the time between addition of polymerase mix and the start of the PCR reaction should be as short as possible. 14. In case, a large number of samples are run simultaneously, the ligation incubation period of the first tubes will be longer. This has no influence on the results. 15. Ligation reaction products can be stored at 4°C for up to 1 week. For longer periods, storage at −20°C is recommended. 16. To avoid contamination, use different micro-pipettes for performing MLPA reactions and for handling MLPA PCR products. After the PCR reaction, do not open the tubes near by the thermocycler. 17. MLPA is less prone to PCR-contamination problems than normal PCR reactions. In a normal PCR reaction, contamination with less than 100 copies of a previous PCR product carrying the same primers will often result in a false-positive signal. In MLPA PCR reactions, however, such amounts of contaminating PCR products are outcompeted by the large numbers of amplifiable fragments that are present at the start of the MLPA PCR reaction. In a typical MLPA reaction, 50 ng human DNA is used, containing 17,000 haploid genome copies. The number of ligated probes will be close to this. At the start of the MLPA-based PCR reaction, there will, therefore, be ~15,000 copies of each of the up to 50 different ligation products, meaning that usually, any contaminating PCR product will be outcompeted. 18. We recommend running 35 PCR cycles. However, it is possible to reduce the amount of cycles to 32. A lower number of PCR cycles will result in a very minor improvement in the linearity of relative probe signal with target sequence copy number. DNA samples containing less than 20 ng of human DNA may require up to 37 PCR cycles. 19. PCR reaction products can be stored at 4°C for at least 1 week and at −20°C for longer periods. The amount of reagents in each kit is sufficient for ~120 PCR reactions, making it possible to repeat some PCR reactions. As the fluorescent labels used are light-sensitive, the PCR products should be stored in a dark box or be wrapped in aluminium foil.

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20. Every SALSA MLPA probemix contains a 92 nt control fragment, which recognises a 2q14 DNA sequence. In most probemixes, this fragment is included with the D-fragments (see Notes 22 and 27). 21. For Applied Biosystems instruments, FAM-labelled primers are used; for Beckman CEQ sequencers, Cy5.0-labelled primers are used and for LICOR sequencers IR800-labelled primers. 22. All SALSA MLPA probemixes (MRC-Holland) contain four DNA Quantity control fragments (Q-fragments), and most of them also contain three DNA Denaturation control fragments (D-fragments; see Note 27). The four Q-fragments (64, 70, 76, 82 nt) are not ligation-dependent and are present in small quantities. When the MLPA reaction has been successful and more than 100 ng sample DNA is used, the Q-fragments are not clearly visible. If the peaks of all four Q-fragments are larger than half the size of the peaks of the D-fragments and the longer MLPA probes (Fig. 7), either the ligation reaction has failed (see Note 24) or the amount of sample DNA is less than 20 ng (see Note 23). In either case, the MLPA results are not reliable. In case, no Q-fragments AND no probe amplification products are present, the PCR reaction has failed (see Note 19). 23. Optical density (260 nm) measurements often overestimate the DNA concentration. In case of doubt, agarose gel electrophoresis can be used to check the presence of DNA. Usually, 20 ng DNA will be visible on ethidium bromidestained agarose gels and will give a smear in the 10,000 bp range. Please note that in samples with very low DNA concentrations, the peak size of all four Q-fragments will be increased. In case, only one increased peak is visible in the 64–82 nt range, it is not a Q-fragment but a non-specific amplification product or long primer-dimer, that has the same mobility as one of the Q-fragments.

Fig. 7. Effect of DNA quantity on Q-fragments. MLPA results with 5 ng (a), 10 ng (b), 20 ng (c) of DNA. Peaks of the Q-fragments decrease when the amount of sample DNA added is increased; primer peak (x). Primer-dimer peaks are often present at ~50–80 nt.


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24. The ligation reaction is very robust. Over 90% of the ligation is complete within 2 min of incubation at 54°C. Problems with the hybridisation or ligation reaction will usually result in variable peak patterns, in which most amplification products will still be present, although of variable height. Evaporation during the hybridisation reaction will often result in peak patterns with considerable sloping. 25. Some probes hybridise faster to their targets than others. The concentration of probes in the probemix has been adjusted to give almost complete hybridisation of each probe after the 16 h incubation. Some causes of incomplete hybridisation are: (1) wrong thermocycler programme, e.g. incubation at 60°C for only 16 min instead of 16 h, (2) the amount of probemix or MLPA buffer used was insufficient, (3) defective heated lid, resulting in evaporation of all fluid during overnight incubation (see Note 34), (4) no denaturation of sample DNA before hybridisation. Other causes of unexplainable decrease and/or increase of certain probe signals can be an incomplete sample denaturation (see Note 27) or an incomplete denaturation of the formamide/PCR reaction/size standard mix before capillary electrophoresis (see Note 26). 26. Brief heating (e.g., 2 min at 80°C) of the formamide/PCR reaction/size standard mix before the capillary electrophoresis is essential. If this heating step is omitted, sequences with a high Guanine and Cytosine content may remain doublestranded, whereas other sequences may denature and thus give a normal peak without this step. Be aware that doublestranded fragments will result in low peaks at unexpected locations. 27. Most SALSA MLPA probemixes contain three D-fragments (denaturation fragments) at 88, 92, and 96 nt. These D-fragments are DNA- and ligation-dependent, just like normal MLPA probes. They will be visible when ligation was successful and DNA amount was sufficient and properly denatured. The 88 nt and 96 nt D-fragments are synthetic MLPA probes that recognise a sequence very close to a strong CpG island. CpG islands have a very high CG content, rendering them extremely difficult to denature. In case, the 88 or the 96 nt fragment peak is much lower ( Troubleshooting. 29. The gel used for capillary electrophoresis can easily deteriorate, especially after prolonged exposure to temperatures over 25°C. When the instrument is not used daily, remove the gel from the instrument after use and store at 4°C. In case, the peaks of the size standard are low and broad, it is almost certain that the problem lies in the capillaries and/or gel. 30. The standard MLPA protocol in which the polymerase mix is added at 60°C requires that the PCR starts as soon as possible after addition of the polymerase mix to the sample. In case, large numbers of samples are tested, it is more convenient to prepare the PCR reactions on ice. After addition of the polymerase mix to the last samples, the PCR reactions should be transferred to a preheated thermocycler (60–72°C) and the PCR should be started immediately. For large numbers of samples, the use of multi-channel pipettes reduces the time required to start the PCR and will result in improved results. 31. Peak sizes do not increase linearly with the use of increasing amounts of MLPA PCR products in the formamide/PCR


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reaction/size-standard mix. Addition of a larger volume of the MLPA PCR products also increases the salt concentration in this formamide/PCR reaction/size-standard mix, causing reannealing of the DNA fragments. When an increase in peak heights is desired, increasing the injection time or injection voltage might be more effective (see http://www.mlpa.com – MLPA procedure – Fragment Separation) 32. Formamide can become acidic when it is stored for a long period at room temperature. This can result in depurination and fragmentation of the DNA upon heating. Store the formamide at −20°C. 33. For each type of instrument, the detection range for optimal quantification of fluorescent units is different. The signal of some types of instruments is saturated at a fluorescence level of 8000 (ABI-310/3100/3130), while other instruments (Beckman CEQ) use fluorescence levels that range up to 180,000 arbitrary units. Preferably, the highest peaks should not exceed 75% of the instrument’s maximum, and the majority of the peaks should be in the 10–50% range. 34. Excessive evaporation can be checked by the incubation of 8 ml water overnight at 60°C. The next morning, at least 5 ml water should still be present at the bottom of the tube. In case of excessive evaporation, try a different brand of tubes, increase or decrease the pressure of the heated lid on the tubes, check whether the heated lid works properly, or try using mineral oil on top of the PCR mix (e.g., Vapor-lock from Qiagen). At high temperatures, the plastic of the tubes is weak. As a result, deformation of tubes can occur when the pressure of the heated lid is too high, or when the pressure is adjusted while the tubes are at temperatures higher than 60°C. This can result in tube leaks and excessive evaporation. 35. Correction for signal sloping: In some runs, the peaks of the longer MLPA probes may be more than three times lower than those of shorter probes. This is called signal sloping. The most common cause of this phenomenon is the presence of impurities in the DNA sample, the use of old gel or capillaries, too high injection voltage (see 1.5 “Raw Data Evaluation ”), low-quality formamide, or sample evaporation during the hybridisation reaction (see Note 34). A correction for signal sloping should be included in the analysis when the signal sloping in a patient sample run differs from the signal sloping in the reference runs. The Coffalyser software is able to correct for this effect. 36. The PCR reaction in MLPA is more sensitive to sample contaminants than normal PCR reactions. Contaminants include phenol, ethanol, and salts. We recommend comparing only

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samples extracted/purified by the same method. Samples derived from paraffin-embedded formalin-fixed tissues should be compared to each other or to DNA from healthy tissue that has undergone a similar treatment. Problems with DNA purified by automated sample processing have also been reported (see Note 2). In case of doubts about DNA quality, it is useful to try a DNA sample from another lab or a commercially obtained DNA. It is often possible to clean contaminated samples by ethanol precipitation or silica-based clean-up kits. DNA losses during ethanol precipitation can be reduced by inclusion of 10 mg glycogen (Roche Diagnostics). In case, the Q-fragments are very low or absent, try using less sample DNA. 37. Background peaks can appear due to contamination. If the peaks in the no-DNA control have the same length as the normal peaks, first try to repeat the PCR reaction using the same ligation reaction. 38. It is recommended to normalise only samples that have been: (a) Extracted by the same method (b) Run within the same experiment (c) Run with the same probemix lot 39. Make sure you reset the bin settings on your sequencer software when using a different probemix lot, a different sequencer, or different run settings. 40. For the characterisation of tumours by MLPA, a more robust normalisation method is needed (18), as the target sites of some reference probes may be gained or lost. Intra-sample normalisation should be performed by dividing the signal of each target-specific probe by the signal of every single reference probe in that sample, thus creating as many ratios per target-specific probe as there are reference probes. Subsequently, the median of all these produced ratios per probe should be taken; this is the probe’s Normalisation Constant. Secondly, inter-sample comparison should be performed by dividing the Normalisation Constant of each probe in a given sample by the average Normalisation Constant of that probe in all the reference samples. 41. Peak area versus peak height: both peak area and peak height can be used to calculate probe signals. Theoretically, peak areas reflect the amount of fluorescence best. However, not all programs calculate the peak areas correctly, leading to false positives or negatives. This problem often occurs when the base of the peaks is wider or when the fragment analysis software separates a peak into a main peak and shoulder peak. In this case, calculating peak areas can lead to inaccuracies, and it is then better to use peak height.


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42. Please note that this type of normalisation assumes that no changes occurred in the genomic regions recognised by the reference probes. The robustness of this normalisation method depends on the number of reference runs, reference probes, chromosomal location of reference probes, and the origin of the reference samples. 43. An MS-MLPA probe targets a single specific HhaI site in a CpG island; if methylation is absent for a particular CpG-site, this does not necessarily mean that the whole CpG island is unmethylated! For samples containing both tumour and normal cells, MLPA experiments will indicate the average copy number or methylation status. 44. It is possible to find a copy number change with a reference probe. Each healthy person is expected to have an “unusual” copy number variation (CNV) for dozens of sequences. Results showing more than one copy number change of reference probes or non-neighbouring target-specific probes (e.g., exons 2 and 15) should be treated with caution, as this may indicate an unreliable MLPA reaction. Please note that blood cells may have acquired copy number changes that are not present in other tissues. We have encountered DNA blood samples with 13q14 loss and blood samples with a trisomy 12. Both these changes are often found in chronic lymphocytic leukaemia. 45. A polymorphism or point mutation in the probe’s hybridizing sequence can also result in a reduced peak area, even when located at a distance of 20 nt from the probe ligation site. Copy number changes detected by only a single probe will, therefore, always require confirmation by other methods. 46. Probe ratios of pathogenic mutations can be above 0.7 and/ or below 1.3. For example, a mosaic case of trisomy 21 has been detected with ratios of 1.1–1.15 of all eight chromosome 21 probes in the P095 aneuploidy SALSA MLPA kit. References 1. Schouten, J.P., McElgunn, C.J., Waaijer, R., Zwijnenburg, D., Diepvens, F. and Pals, G. (2002). Relative quantification of 40 nucleic acid sequences by multiplex ligation-dependent probe amplification. Nucleic Acids Res 30, e57. 2. Aretz, S., Stienen, D., Uhlhaas, S., Stolte, M., Entius, M.M., Loff, S., Back, W., Kaufmann, A., Keller, K.M., Blaas, S.H., Siebert, R., Vogt, S., Spranger, S., Holinski-Feder, E., Sunde, L., Propping, P. and Friedl, W. (2007). High proportion of large genomic deletions and a genotype phenotype update in 80 unrelated

families with juvenile polyposis syndrome. J Med Genet 44, 702–709. 3. Redeker, E.J., de Visser, A.S., Bergen, A.A. and Mannens, M.M. (2008). Multiplex ligationdependent probe amplification (MLPA) enhances the molecular diagnosis of aniridia and related disorders Mol Vis 14, 836–840. 4. Kanno, J., Hutchin, T., Kamada, F., Narisawa, A., Aoki, Y., Matsubara, Y. and Kure, S. (2007). Genomic deletion within GLDC is a major cause of non-ketotic hyperglycinaemia. J Med Genet 44, 3.

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5. Aldred, M.A., Vijayakrishnan, J., James, V., Soubrier, F., Gomez-Sanchez, M.A., Martensson, G., Galie, N., Manes, A., Corris, P., Simonneau, G., Humbert, M., Morrell, N.W. and Trembath, R.C. (2006). BMPR2 gene rearrangements account for a significant proportion of mutations in familial and idiopathic pulmonary arterial hypertension. Hum Mutat 2, 212–213. 6. Kluwe, L., Nygren, A.O., Errami, A., Heinrich, B., Matthies, C., Tatagiba, M. and Mautner, V. (2005). Screening for large mutations of the NF2 gene. Genes Chromosomes Cancer 42, 384–391. 7. Michils, G., Tejpar, S., Thoelen, R., van Cutsem, E., Vermeesch, J.R., Fryns, J.P., Legius, E. and Matthijs, G. (2005). Large deletions of the APC gene in 15% of mutationnegative patients with classical polyposis (FAP): a Belgian study. Hum Mutat 2, 125–134. 8. Taylor, C.F., Charlton, R.S., Burn, J., Sheridan, E. and Taylor, G.R. (2003). Genomic deletions in MSH2 or MLH1 are a frequent cause of hereditary non-polyposis colorectal cancer: identification of novel and recurrent deletions by MLPA. Hum Mutat 6, 428–433. 9. Depienne, C., Fedirko, E., Forlani, S., Cazeneuve, C., Ribaï, P., Feki, I., Tallaksen, C., Nguyen, K., Stankoff, B., Ruberg, M., Stevanin, G., Durr, A. and Brice, A. (2007). Exon deletions of SPG4 are a frequent cause of hereditary spastic paraplegia. J Med Genet 44, 281–284. 10. Beetz, C., Nygren, A.O., Schickel, J., AuerGrumbach, M., Bürk, K., Heide, G., Kassubek, J., Klimpe, S., Klopstock, T., Kreuz, F., Otto, S., Schüle, R., Schöls, L., Sperfeld, A.D., Witte, O.W. and Deufel, T. (2006). High frequency of partial SPAST deletions in autosomal dominant hereditary spastic paraplegia. Neurology 67, 1926–1930. 11. Eldering, E., Spek, C.A., Aberson, H.L., Grummels, A., Derks, I.A., de Vos, A.F., McElgunn, C.J. and Schouten, J.P. (2003). Expression profiling via novel multiplex assay

12.

13.

14.

15.

16.

17.

18.

allows rapid assessment of gene regulation in defined signalling pathways. Nucleic Acids Res 31, e153. Nygren, A.O., Ameziane, N., Duarte, H.M., Vijzelaar, R.N. et al. (2005). Methylationspecific MLPA (MS-MLPA): simultaneous detection of CpG methylation and copy number changes of up to 40 sequences. Nucleic Acids Res 33(14), e128. Bittel, D.C., Kibiryeva, N. and Butler, M.G. (2007). Methylation-specific multiplex ligation-dependent probe amplification analysis of subjects with chromosome 15 abnormalities. Genet Test 11, 467–475. Dikow, N., Nygren, A.O., Schouten, J.P., Hartmann, C., Krämer, N., Janssen, B. and Zschocke, J. (2007). Quantification of the methylation status of the PWS/AS imprinted region: comparison of two approaches based on bisulfite sequencing and methylation-sensitive MLPA. Mol Cell Probes 3, 208–215. Procter, M., Chou, L.S., Tang, W., Jama, M. and Mao, R. (2006). Molecular diagnosis of Prader-Willi and Angelman syndromes by methylation-specific melting analysis and methylation-specific multiplex ligation-dependent probe amplification. Clin Chem 52, 1276–1283. Jeuken, J., Cornelissen, S., Boots-Sprenger, S., Gijsen, S. and Wesseling, P. (2006). Multiplex ligation-dependent probe amplification: a diagnostic tool for simultaneous identification of different genetic markers in glial tumors. J Mol Diagn 4, 433–443. Hess, C.J., Errami, A, Berkhof, J., Denkers, F., Ossenkoppele, G.J., Nygren, A.O., Schuurhuis, G.J. and Waisfisz, Q. (2008). Concurrent methylation of promoters from tumor associated genes predicts outcome in acute myeloid leukemia. Leuk Lymphoma 49, 1132–1141. Coffa, J., van de Wiel, M.A., Diosdado, B., Carvalho, B., Schouten, J. And Meijer, G.A. (2008). MLPAnalyzer: data analysis tool for reliable automated normalization of MLPA fragment data. Cell Oncol 30, 323–335.

Chapter 9 Screening for Genomic Rearrangements by Multiplex PCR/Liquid Chromatography Claude Houdayer, Catherine Dehainault, Marion Gauthier-Villars, and Dominique Stoppa-Lyonnet Abstract Screening for large gene rearrangements has become established as an important part of molecular medicine; however, it is also challenging as these rearrangements range from an extra copy of a complete chromosome(s) to deletion or duplication of a single exon. In this chapter, we describe a versatile and robust method to assess exon copy number, called multiplex PCR/liquid chromatography (MP/LC) assay. Multiple genomic fragments are amplified under semiquantitative conditions using unlabeled primers, then separated by ion-pair reversed-phase high-performance liquid chromatography, and quantitated by fluorescent detection using a postcolumn intercalation dye. The relative peak intensities for each target directly reflect DNA copy number. This technique can be used not only to screen intronic, exonic, and intergenic parts of the genome but also for transcript quantitation. MP/LC appears to be an easy, versatile, and cost-effective method, which is particularly relevant to DHPLC users, as it broadens the spectrum of available applications on a DHPLC system. The authors describe a detailed protocol for large rearrangement screening in the RB1 gene. Key words: Multiplex PCR/liquid chromatography assay, Genomic rearrangements, DHPLC, Mutation detection

1. Introduction Genomic disorders, also termed rearrangement-based disorders, are caused by an alteration of the genome which can lead to the loss or gain of a dosage-sensitive gene or disruption of a gene (1). This group of disorders is distinguished from conventional Mendelian diseases in that the phenotype does not result from a point mutation, but rather from larger alterations of the genome. Such alterations range from extra copy of a complete chromosome(s) to deletion or duplication of a single exon. Duchenne/Becker Bimal D.M. Theophilus and Ralph Rapley (eds.), PCR Mutation Detection Protocols, Methods in Molecular Biology, vol. 688, DOI 10.1007/978-1-60761-947-5_9, © Springer Science+Business Media, LLC 2011

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muscular dystrophy is a striking example, as about 70% of cases of the disease are due to large deletions or duplications (2, 3). More generally, a 10–20% figure is observed for other disease-linked genes such as RB1, MSH2, MLH1, MSH6, or CFTR (4–8). Chromosome rearrangements are not random events, but result from predisposition to rearrangement due to the existence of a specific genomic context. Both homologous and nonhomologous recombinations can, therefore, occur at certain hotspots throughout the genome. Shaw and Lupski (9) speculated that the same genome flexibility that has enabled man to evolve relatively rapidly also makes the human species more susceptible to rearrangements associated with disease. Since the first comprehensive review on this topic (10), the number of characterized genomic disorders has continued to expand (9), which is why screening for gross rearrangements has become established as an important part of molecular diagnosis, although it requires dedicated tools as conventional PCR-based methods are mainly designed for point mutation detection. Consequently, such rearrangements go undetected because they are hidden by the normal allele or the other altered allele in dominant or recessive diseases, respectively. Chromosomal duplications or deletions can be detected using standard and molecular cytogenetic approaches such as high-resolution karyotyping, fluorescent in situ hybridization (FISH), or color bar coding on combed DNA ((11)and http://www.genomicvision.com). These techniques are being challenged by arrayCGH using PAC and BAC clones which has a higher throughput than FISH and is particularly useful to detect rearrangements not visible by standard karyotypic analysis (12). More recently, the so-called zoom-in microarray-CGH has reached high-resolution coverage close to 200–300 bp, allowing the detection of more discrete rearrangements that may involve a single exon (13). However, this powerful approach remains costly and can therefore not be applied in large-scale routine analyses. Hemizygosity studies based on segregation analysis of microsatellite markers require sampling of the parents and also depend on the informativeness of the markers. Transcript-based analysis may be an attractive alternative but is limited by RNA availability and could result in false-negatives due to nonsense-mediated decay (14). Real-time PCR strategies on genomic DNA are the methods of choice for DNA quantitation and therefore gene dosage. However, throughput remains limited by the small number of dyes currently available, making these strategies inappropriate for routine detection of exon deletion and duplication in large genes such as NF1, BRCA1, or RB1. An elegant multiplex ligation-dependent probe assay (MLPA) is widely used by diagnostic laboratories (see Chapter 8). It consists of relative quantification of up to 45 different DNA sequences in a single tube. Following probe hybridization, ligation,

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and PCR, amplification products are detected and quantified by capillary electrophoresis (15). This technique is easy to use when available as a manufactured kit for the gene of interest (see http:// www.mrc-holland.com) but proves to be time-consuming and difficult to implement for other applications (16). Lastly, various semiquantitative multiplex fluorescent PCR protocols have been used to detect gross rearrangements (17), culminating in the socalled quantitative multiplex PCR of short fluorescent fragments (QMPSF) (18). QMPSF has been successfully adapted to different genes (4, 19, 20) but requires purified fluorescently labeled oligonucleotides, with the attendant high cost. In this chapter, we describe a new, versatile, and robust method to assess DNA copy number. This method, called multiplex PCR/liquid chromatography assay (MP/LC), is based on the well-known properties of ion-pair reversed-phase high-performance liquid chromatography (IP-RP-HPLC) for separation and quantitation of nucleic acids (21) (see Chapter 3). First, a multiplex PCR with unlabeled primers enables simultaneous amplification of multiple genomic fragments under semiquantitative conditions. Second, PCR products are separated by nondenaturing IP-RP-HPLC and, lastly, are quantitated by fluorescent detection using a postcolumn intercalation dye. The exceptional sensitivity of the dye allows the use of limiting PCR conditions to remain in the exponential phase of amplification. Consequently, relative peak intensities for each amplicon directly reflect fragment copy number. The strength of MP/LC lies in its simplicity and cost-effectiveness. Therefore, multiple custom applications dealing with DNA copy number and requiring a variety of primer sets can be assayed by MP/LC (22–28). Intronic, exonic, and intergenic parts of the genome can be screened as previously demonstrated in a study describing the mapping and sequencing of the breakpoints in a contiguous gene syndrome (29). MP/LC also proves useful in expression analyses to quantitate the relative expression of a gene of interest (30). MP/LC appears to be parti cularly relevant to DHPLC users since it broadens the spectrum of available applications on a DHPLC system. The use of MP/LC for high-throughput screening of RB1 large rearrangements is described below. Constitutional mutations of the RB1 gene are associated with a predisposition to retinoblastoma (MIM 180200). Retinoblastoma is the most common pediatric eye tumor, with an incidence of 1 in 15,000–20,000. It is essential to identify RB1 mutations to provide appropriate genetic counseling in retinoblastoma patients, but this represents an extremely challenging task and a good mutational model, as the vast majority of mutations are unique and spread over the entire coding sequence. Gross rearrangement screening is mandatory in RB1 analysis, as an average 13% of the mutational spectrum consists of large deletions (6, 31). We studied more than 500 consecutively

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diagnosed cases for constitutional RB1 rearrangement screening. Individual written consent for molecular analysis was obtained from each subject sampled or from their legal guardian(s).

2. Material 2.1. DNA and Controls

1. DNA calibrated to 50 ng/ml by UV spectrophotometric assay (e.g., Nanodrop). 2. Prepare 50 ng/ml dilutions of control DNA (mutant controls, i.e., with RB1 deletions or duplication, and normal controls). It is preferable to use control DNA extracted under the same conditions as the test DNA.

2.2. PCR (See Note 1)

1. AmpliTaq DNA Gold, 5 U/ml (PE Applied Biosystems). 2. Standard primers. 3. Microtubes, pipette tips. 4. Quality control: Carry out amplification of normal control and mutant control DNA in each multiplex, as well as amplification of an internal control amplicon in each PCR mix. The internal control amplicon should be preferably chosen on a chromosome that is different from that of the gene of interest, and in the case of tumor studies, a chromosome not subject to rearrangement must be used.

2.3. WAVE 3500HS (Transgenomic)

1. Sizing standard: WAVE® DNA Sizing Standard for WAVE Systems pUC18 HaeIII Digest (fragment sizes: 80, 102, 174, 257, 267, 298, 434, 458, and 587 bp). 2. Staining solution: Wave OPT HS staining.

3. Method 3.1. Preanalysis Step: Preparation of Primer Mixes and PCR

1. Dilute primers to 20 mM in TE 10:1 (Table 1). 2. Pool the forward and reverse primers of the same amplicon in a single tube. Each primer is added at a concentration of 10 mM. 3. Prepare primer mixes as indicated in Table 1. 4. Prepare five PCRs according to the conditions indicated in Table 2 and use the amplification protocols shown in Table 3. Amplify at least two normal control DNAs and two mutant control DNAs. 5. Following PCR, store at −20°C until elution.

AGCTTTTGAAAGCAGAGTCA

AFM-exon 9

TCTTTAATGAAATCTGTGCC

CTTATGTTCAGTAGTTGTGG

Exon 10

Exon 13

TGTATTTTTTTAATGACAATTCAG CTCCACATGCAAGTTTGAAAC

ATAATCTGTGATTCTTAGCC

BRCA1-exon 11

Exon 19

Exon 26

Multiplex 4

AGTAAGTCATCGAAAGCATC

CTTATTCCCACAGTGTATCG

Exon 20

H2O

TCCTCAGACATTCAAACGTG

Exon 22

AACGAAAAGACTTCTTGCAG

AAGAAACATGATTTGAACCC

TCTTACTTGGTCCAAATGCC

AGGATACTTTTGACCTACCC

TAAACAATCAAAGGACCGAG

TTTGCTTACATATCTGCTGC

TATACGAACTGGAAAGATGC

Exon 18

Multiplex 3

TGTATTTTTTTAATGACAATTCAG CTCCACATGCAAGTTTGAAAC

BRCA1-exon 11 GATATCTAAAGGTCACTAAG

CATTTGGTAGGCTTGAGTTTG

ATCCTTACCAATACTCCATCC

GTAACGGTAACAACCTGGAG

GGGTCTGATAGGGAAGACTC

Reverse

Exon 2

Multiplex 2

GCTCCAAGTTTGTTTTTGTT

Forward

Primer sequence (5¢ to 3¢)

Promoter

Multiplex 1

Pooled primers (10 mM)

Table 1 Sequence primer and multiplex preparation

9

22

14

22

24

6

12

b

b

Volume of pooled primers in the mix (ml)

0.05

0.2

0.31

0.34

0.08

0.17

0.8

0.8

0.5

0.8

0.5

0.8

Concentration (mM)a

(continued)

209

273

205

185

159

125

343

295

205

165

239

156

Amplicon size (bp)

Screening for Genomic Rearrangements by Multiplex PCR/Liquid Chromatography 131

CGCCATCAGTTTGACATGAG

CCTGCGATTTTCTCTCATAC

CAGTTTTAACATAGTATCCAG

GGCAGTGTATTTGAAGATAC

GGGAGGCTCTTTAGCTTC TTAGGACAGC

AGTAGTAGAATGTTACCAAG

Exon 27

Exon 7

Exon 3

Exon 12

BRCA1-exon 18

Exon 8

GCATGAGAAAACTACTATGAC

GTATTTATGCTCATCTCTGC

CACCCAAAAGATATATCTGG

GAGACAACAGAAGCATTATAC

AGCCTGGGCAAAACAGTGAG

ATCTAATGTAATGGGTCCAC

GGTTGCTAACTATGAAACAC

Exon 5

Exon 24

Exon 6

Exon 11

Exon 14

Exon 23

Exon 25

Multiplex 5

H2O

TGCATTGTTCAAGAGTCAAG

Forward


Exon 9


Table 1 (continued)

AGAAATTGGTATAAGCCAGG

CTTGGATCAAAATAATCCCC

AGCCTGGGCAAAACAGTGAG

CGTGAACAAATCTGAAACAC

ATTTAGTCCAAAGGAATGCC

TGAGGTGTTTGAATAACTGC

CTAACCCTAACTATCAAGATG

TACTGCAAAAGAGTTAGCAC

GAGACCCATTTTCCCAGC ATCACCAGC

AACTACATGTTAGATAGGAG

AGCATTTCTCACTAATTCAC

ATGTTTGGTACCCACTAGAC

CAGTCACATCTGTGAGAGAC

AGTTAGACAATTATCCTCCC

Reverse

12

12

8

16

10

10

12

25.2

23.4

12.6

23.4

36

14

14.4

22


0.07

0.07

0.05

0.09

0.06

0.06

0.07

0.13

0.07

0.13

0.20

0.08

0.08

0.12

Concentration (mM)a

299

289

268

247

224

212

196

382

358

311

282

257

238

223

Amplicon size (bp)

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AAACCTTTCTTTTTTGAGGC

AAAAATACCTAGCTCAAGGG

GGGAGGCTCTTTAGCTTCTT AGGACAGC

CAATGCTGACACAAATAAGG

Exon 21

Exon 17

BRCA1-exon 18

Exons 15 and 16

AGCATTCCTTCTCCTTAACC

GAGACCCATTTTCCCAGCAT CACCAGC

TGTTAAGAAACACCTCTCAC

TACATAATAAGGTCAGACAG

CCCAGAATCTAATTGTGAAC

Reverse

b

Directly add following the volumes indicated in Table 2 to the PCR

Control amplicons are italicizedaIn the final PCR volume; forward and reverse equimolar

H2O

GTAGTGATTTGATGTAGAGC

Forward


Exon 4


6

24

12

12

16

20


0.14

0.07

0.07

0.09

0.12

Concentration (mM)a

368

358

349

330

307

Amplicon size (bp)

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Table 2 PCR protocol for the five multiplexes PCR protocol for Multiplex 1 (gold 4 mM MgCl2) 25 ml final 1 reaction ( ml) H2O

7

10× gold

2.5

MgCl2 (4 mM final)

4

dNTP

4

Promoter

2

AFM exon 9 (control peak)

1.3

DMSO (7.5%)

1.9

Taq gold

0.3

DNA (50 ng/ml)

2

Total

25

PCR protocol for Multiplex 2 (gold 4 mM MgCl2) 25 ml final 1 reaction (ml) H2O

3.7

10× gold

2.5

MgCl2 (4 mM final)

4

dNTP

4

Exon 10

2

Exon 13

2

Exon 2

2

AFM exon 9 (control peak)

1.3

DMSO (5% final)

1.2

Taq gold

0.3

DNA (50 ng/ml)

2

Total

25


7.5 (continued)


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Table 2 (continued) 10× gold

2.5

MgCl2 (4 mM final)

4

dNTP

4

Primer mix

3.5

DMSO (5% final)

1.2

Taq gold

0.3

DNA (50 ng/ml)

2

Total

25

PCR protocol for Multiplex 4 (gold 6 mM MgCl2) 20 ml final

1 reaction (ml)

H2O

5.8

10× gold

2

MgCl2 (6 mM final)

4.8

dNTP

3.2

Mix primers

2

Taq gold

0.2

DNA (50 ng/ml)

2

Total

20


4.8

10× gold

2

MgCl2 (6 mM final)

4.8

dNTP

3.2

Primer mix

2

DMSO (5% final)

1

Taq gold

0.2

DNA (50 ng/ml)

2

Total

20

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Table 3 Touchdown protocol for multiplexes 1 and 3 and PCR protocol for multiplexes 2, 4, and 5 Multiplex 1 Denaturation (°C)

95

95

95

95

95

95

95

95

Annealing (°C)

60

59

58

57

56

55

54

53

Elongation (°C)

72

72

72

72

72

72

72

72

Number of cycles

2

2

2

3

3

4

4

5

Denaturation (°C)

95

95

95

95

95

95

95

95

Annealing (°C)

51

50

49

48

47

46

45

44

Elongation (°C)

72

72

72

72

72

72

72

72

Number of cycles

2

2

2

3

3

4

4

5

Multiplex 3

Multiplexes 2, 4, and 5 Denaturation (°C)

95

Annealing (°C)

55

Elongation (°C)

72

Number of cycles

23

Denaturation and annealing steps take 30 s, the elongation step takes 45 s. An initial denaturation step of 95°C for 5 min and a final elongation step of 72°C for 5 min are needed.

3.2. Analysis Step: Elution of Amplified Products on Wave 3500HS

1. Elute the quality standards and Wave DNA Standard Sizing to ensure the good quality of the system and the column. 2. Connect the HSX module and the fluorescence detector (see Note 2). The modules are connected/disconnected in Wave software Navigator > Setup > Instrument > Module Setup. 3. Switch the two modules from “Available modules” to “Associated modules.” Reconfigure and reinitialize the system in Navigator Operator. 4. Check and, if necessary, adjust the Staining Solution volume. Staining Solution is stable for 3 weeks at room temperature. 5. Purge the fluorescence system before each use. 6. In Wave Navigator v1.7.0 software, create a new tray name in the desired project.


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Table 4 Gradients used for RB1 analysis Gradient Method name

Fragment sizes (bp)

%B, initial

%B, final

Duration (min)

Multiplex 1

156, 239

47.3

61.8

6.3

Multiplex 2

165, 205, 295, 343

47.3

61.8

6.3

Multiplex 3

125, 159, 185, 205, 273

47.3

61.8

6.3

Multiplex 4

209, 223, 238, 257, 282, 311, 358, 382

56.6

62.7

8.4

Multiplex 5

196, 212, 224, 247, 268, 289, 299, 307, 330, 349, 358, 368

54.3

60.6

9.6

Associate the method corresponding to the PCR (see Table 4). At the beginning of elution, program 2 blanks (use the same method as the samples, volume = 0 to equilibrate the system). 7. Run the analyses. 3.3. Analysis Step: Interpretation of Data in Wave Navigator v1.7.0 (build 25) Software

1. Select the Tray Name.

3.3.1. To Visualize the Results

2. In the table, select the lines of samples to be examined and click on to display the results to the “Analysis” window, or directly in the “Analysis” window, click on “Select Results” underneath the chromatogram, choose the Tray Name in the corresponding project, select the samples to be examined and click on “Add Selected Results.”

3.3.2. To Interpret the Results

1. Analyze all profiles at the same time: tick the profiles of interest and tick “Show all.” Always align the profiles of interest with the various normal and mutant Controls (Fig. 1a–e). 2. Use “mutation calling” to align profiles on the intensity of the control peak. 3. Click on the “mutation calling” icon. In the “normalization” part, select the selected peak in “Peak compare” and enter the limits of the peak corresponding to the amplicon control with which the intensities must be aligned. 4. In the “mutation calling” part, enter the interval of analysis of the fragments. 5. Do not change the “clustering” part and the “discrimination” part (leave the default parameter: automated). 6. Click on “Perform analysis” and perform interpretation.

Fig. 1. (a–e) MP/LC chromatograms for the five different multiplexes used in RB1 analysis. X-axis: retention time in min; Y-axis: fluorescence intensity. Exons under study are indicated at the top of the corresponding peaks. The chromatograms represent the normal and deleted controls, as indicated by arrows. Profiles are superimposed and then normalized using the control peak. A whole-gene deleted control is used except for Multiplex 4 for which a deletion from promoter to exon 17 was used. This figure shows the good resolution of the three first peaks that are situated close together. Adapted from ref. (28).


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4. Notes 1. PCR: Manipulate on ice. Mixes are prepared in a fume hood and DNAs are added on a dedicated laboratory bench top. Pipette tips are fitted with a cotton filter, and the pipettes are used specifically for the preparation of mixes and manipulation of DNA. Precautions are essentially designed to protect manipulation. 2. HSX module: In addition to the eight classical modules, the Wave 3500HS is equipped with a HSX (ICA-2000) module allowing incorporation into DNA of a fluorescent intercalator present in the Staining Solution. It is linked to a fluorescence detector (detector HSD L-7485). Incorporation and detection of fluorescence are performed after UV detection when the modules are connected. 3. GC-rich regions: The 5¢ part of the RB1 gene (promoter and exon 1) is particularly rich in GC, which can make it difficult to analyze for the detection of both point mutations and rearrangements. Due to the high degree of similarity of the amplified region, nonspecific intrastrand base-pairing tends to occur during PCR, resulting in nonspecific PCR. Consequently, some teams do not analyze this region which results in incomplete analysis, as mutations of the promoter region and exon 1 have been well documented. We have resolved these problems by adding dimethylsulfoxide (7.5% final volume) to the reaction medium. 4. DNA quality and quantity: Semiquantitative multiplex PCR techniques are robust techniques but highly dependent on the quality of the DNA studied. Degraded DNA can be responsible for loss of proportionality between signal intensity and copy number, particularly for large fragments, making the analysis uninterpretable. Contamination of DNA by phenol will have an even greater effect because it generates a random fluctuation of signal intensity. Phenol-free extraction techniques should therefore be preferred (perchlorate/chloroform or column-based commercial kits), or a system ensuring the absence of contamination by phenol such as the gel lock extraction system, which uses a gel-barrier system (Eppendorf). It is also essential to adjust all DNA samples studied to a suitable working concentration, classically 50 ng/ml. If the DNA concentration is too high, for example, the proportionality between signal intensity and copy number will be lost, particularly for small fragments. DNA calibration can be performed with: (a) a tube spectrophotometer (unsuitable for large series), (b) the NanoDrop from NanoDrop technologies (which has the advantage of tracing

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the spectrum of the sample), or (c) a plate reader (rapid, but reading at only one wavelength at a time). In our experience, the use of fluorescent dyes for the assay, such as PicoGreen (Molecular Probes), is unnecessary for these applications. Finally, buccal samples are poorly adapted to these analyses, including for screening for a mutation already identified in a relative, as DNA is often present in a low concentration and difficult to calibrate. In our experience, duplex PCR can be designed, but simultaneous amplification of a greater number of amplicons does not provide reliable results. Due to the importance of the quality, quantity, and calibration of DNA solutions, laboratories often prefer to extract DNA locally and therefore ask to receive whole blood. 5. A classical trap in the interpretation of these techniques concerns the false-positive results generated by a PCR primer mismatch. Each deletion of a single exon must, therefore, be systematically checked by another technique (long-range PCR, RNA studies, real-time PCR, for example) and/or by shifting the primers. Finally, duplication of an isolated exon is the most difficult case to characterize. The ideal situation is, therefore, to have a duplicated control of the sequence of interest, for example, DNA from a case of trisomy 13 in RB1 analyses. References 1. Stankiewicz P, Lupski JR (2002) Genome architecture, rearrangements and genomic disorders. Trends Genet 18:74–82 2. Koenig M, Hoffman EP, Bertelson CJ, Monaco AP, Feener C, Kunkel LM (1987) Complete cloning of the Duchenne muscular dystrophy (DMD) cDNA and preliminary genomic organization of the DMD gene in normal and affected individuals. Cell 50:509–517 3. Den Dunnen JT, Grootscholten PM, Bakker E, Blonden LA, Ginjaar HB, Wapenaar MC, van Paassen HM, van Broeckhoven C, Pearson PL, van Ommen GJ (1989) Topography of the Duchenne muscular dystrophy (DMD) gene: FIGE and cDNA analysis of 194 cases reveals 115 deletions and 13 duplications. Am J Hum Genet 45:835–847 4. Audrezet MP, Chen JM, Raguenes O, Chuzhanova N, Giteau K, Le Marechal C, Quere I, Cooper DN, Ferec C (2004) Genomic rearrangements in the CFTR gene: extensive allelic heterogeneity and diverse mutational mechanisms. Hum Mutat 23:343–357 5. Bombieri C, Bonizzato A, Castellani C, Assael BM, Pignatti PF (2005) Frequency of large

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CFTR gene rearrangements in Italian CF patients. Eur J Hum Genet 13:687–689 Houdayer C, Gauthier-Villars M, Lauge A, Pages-Berhouet S, Dehainault C, CauxMoncoutier V, Karczynski P, Tosi M, Doz F, Desjardins L, Couturier J, Stoppa-Lyonnet D (2004) Comprehensive screening for constitutional RB1 mutations by DHPLC and QMPSF. Hum Mutat 23:193–202 Charbonnier F, Olschwang S, Wang Q, Boisson C, Martin C, Buisine MP, Puisieux A, Frebourg T (2002) MSH2 in contrast to MLH1 and MSH6 is frequently inactivated by exonic and promoter rearrangements in hereditary nonpolyposis colorectal cancer. Cancer Res 62:848–853 Gille JJ, Hogervorst FB, Pals G, Wijnen JT, van Schooten RJ, Dommering CJ, Meijer GA, Craanen ME, Nederlof PM, de Jong D, McElgunn CJ, Schouten JP, Menko FH (2002) Genomic deletions of MSH2 and MLH1 in colorectal cancer families detected by a novel mutation detection approach. Br J Cancer 87:892–897 Shaw CJ, Lupski JR (2004) Implications of human genome architecture for rearrangement-


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familial retinoblastoma. Hum Mutat 28: 284–293 31. Richter S, Vandezande K, Chen N, Zhang K, Sutherland J, Anderson J, Han L, Panton R, Branco P, Gallie B (2003) Sensitive and efficient detection of RB1 gene mutations enhances care for families with retinoblastoma. Am J Hum Genet 72:253–269

Chapter 10 Mutation Surveyor: Software for DNA Sequence Analysis Jayne A.L. Minton, Sarah E. Flanagan, and Sian Ellard Abstract Advances in high-throughput sequencing techniques had presented a significant challenge to the processing capabilities of genetic laboratories. However, recent developments in the field of semi-automated mutation detection have revolutionised the task of mutation detection. This chapter provides user information for one commercially available program, Mutation Surveyor. The software is manufactured by SoftGenetics (Pennsylvania, USA) and provides an accurate and efficient program for detecting sequence variants. The chapter focuses on the methodology of setting up GenBank files as reference files and provides information on analysis parameters and data processing. Key words: Mutation Surveyor, Sequence analysis, Mutation detection, GenBank, SoftGenetics

1. Introduction The completion of the human genome project has assisted in the identification of gene mutations causative of human diseases. Mutations in over 2,000 genes have now been identified in patients with more than 3,000 different disease phenotypes (1). For the clinicians and their patients, it is becoming increasingly important to obtain a genetic diagnosis, as identifying the genetic aetiology of a disease may influence clinical management and will provide information regarding risk to future pregnancies. Mutation detection has previously employed approaches such as heteroduplex analysis by denaturing high-performance liquid chromatography (dHPLC) analysis or conformation-sensitive capillary electrophoresis (CSCE) (see chapters13 and 18). With the advent of high-throughput capillary sequencers and sequence analysis software, direct sequencing provides an accurate method for single gene analysis. High-throughput sequencing is now available with capillary sequencers such as the ABI 3730xl, which can Bimal D.M. Theophilus and Ralph Rapley (eds.), PCR Mutation Detection Protocols, Methods in Molecular Biology, vol. 688, DOI 10.1007/978-1-60761-947-5_10, © Springer Science+Business Media, LLC 2011

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analyse 96 samples at a time (Applied Biosystems, Warrington, UK). With a standard read length of 700 bp, the ABI 3730xl can generate 2,304 reads per day, which equates to >1,600 kb (http://www.AppliedBiosystems.com). The newly found ability to produce these large volumes of data has paved the way for the generation of commercially available automated sequence analysis software programs. These programs provide an accurate and efficient means to detect heterozygous and homozygous mutations in a high-throughput system. This chapter focuses on the Mutation Surveyor Software version 3.20, which is a commercially available computer program for sequence assembly and analysis. The program is manufactured by SoftGenetics (Pennsylvania, USA) and can be used with either fluorescent dideoxyterminator or fluorescent primer-based chemistries from both gel and capillary sequencing systems. A recent study by Ellard et al. (2) has demonstrated that Mutation Surveyor is a highly sensitive method for mutation detection. The authors carried out a large study investigating semi-automated single direction sequence analysis for the detection of heterozygous base substitutions and demonstrated that Mutation Surveyor had a sensitivity of >99.57%. The confidence that Mutation Surveyor provides in semi-automated analysis of unidirectional sequencing reduces the costs of sequencing considerably, saving on both consumables and labour. Setting up Mutation Surveyor GenBank Sequence files is straightforward and once these have been generated, the program can analyse hundreds of reads in a matter of seconds. The output files are clearly tabulated which aids in the interpretation of the results, and mutations and polymorphisms can be named according to the Human Genome Variation Society (HGVS)-approved nomenclature (http://www.hgvs.org/). The features of the program have resulted in Mutation Surveyor being the sequence analysis software program of choice for institutions across the world including the Mayo Clinic and Foundation (USA), National Cancer Institute (USA), Rikken Institute (Japan), Sanger Centre (UK), Max Planck Institute (Germany), National Health Service (UK), Dana Farber Cancer Institute (USA), Johns-Hopkins School of Medicine (USA) (Press release; 9 April, 2007; Softgenetics). The aim of this chapter is to provide information for users regarding the setting up of the GenBank Sequence files utilised by the program, parameters for analysis, and options for data output. More detailed information regarding Mutation Surveyor Software can be found on the SoftGenetics website (http://www.softgenetics.com/mutationSurveyor.html) and in the operation manual that accompanies the computer program.

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2. Materials 1. Reference sequence (downloaded from http://www.ncbi. nlm.nih.gov/). 2. Mutation Surveyor version 3.20 (SoftGenetics, Pennsylvania, USA).

3. Methods 3.1. Setting Up Mutation Surveyor 3.1.1. Creating a GenBank File

A GenBank reference sequence (.gbk) is used for the data analysis (see Note 1). GenBank files specific to a particular gene can be obtained from the Entrez Gene database, which is available through the NCBI website [http://www.ncbi.nlm.nih.gov/] (see Note 2): 1. Using the Entrez gene website, navigate to the home page for the gene of interest using the gene name and species as search criteria. 2. Click on the link “Map Viewer”, which can be found under the “Links” section. The map for the given gene will appear. Click on “dl” (download/view sequence), which is in the purple highlighted information section for the gene. 3. In the drop-down menu of the sequence format, choose GenBank and then click “Display”. 4. Further annotations to the GenBank file e.g. known variations/polymorphisms can be incorporated by clicking on “SNP”, followed by “Refresh”. 5. The annotated GenBank file can now be saved. To do this, select “text” from the “Send to” drop-down menu. Then, use the “Save As” option of the “File” menu to complete this task, making sure to choose the .txt file format in the “save as type” field.

3.1.2. Editing the GenBank File in Mutation Surveyor

1. Launch Mutation Surveyor and import the GenBank file into the “GBK File Editor”, which can be found in the Tools menu (see Note 3). 2. Click “Save” to save the reference file in a GenBank file format (.gbk). This file can now be uploaded directly into Mutation Surveyor as the GenBank sequence file for sequence analysis.

3.1.2.1. Defining the Region of Interest

Mutation surveyor will automatically create a region of interest (ROI) with a set number of nucleotides flanking either side of the exon. It is possible to redefine the ROI for individual reads (see Note 4):

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1. Within the GenBank file Editor, under the “Features2 tab”, set the ROI for each coding sequence (CDS) by entering the number of the 5′ nucleotide from which the ROI starts and in the “to” box place the most 3′ nucleotide number. The numbering system should correlate to the GenBank file. 2. Click on “Refresh” and re-save the annotated GenBank file. 3.1.2.2. Adding Known Variants to a GenBank File

The GenBank file should already be annotated with SNPs, which are recorded on the dbSNP database. Further, unreported sequence variants can be incorporated into the GenBank file so that during analysis they are flagged as known variants. To do this: 1. Identify the position of the variant in relation to the reference GenBank file. 2. From the “Tools” menu, select the “GBK File Editor” to open the relevant GenBank file. Select the “Features2” tab and add the nucleotide changes to the “variants” box (e.g. 123A/G(2271026) or 123R(2271026) with nucleotide changes corresponding to the GenBank file, represented in the standard IUB/IUPAC nucleic acid codes followed by the variants RefSNP accession ID (rs number)).

3.2. Data Processing 3.2.1. Importing Test and Reference Sequences

Once the GenBank files have been generated, the sample files are ready for analysis: 1. Launch Mutation Surveyor and from the File menu select “open files”. The “Open Files” dialog box that contains three panes will appear. 2. The reference GenBank file should be added to the top “GenBank Sequence” pane. To do this, click on the “Add” button and navigate to the saved GenBank file and then click “open”. The name of the reference GenBank Sequence file along with its path will appear in the “GenBank Sequence File” top pane. 3. The middle “Reference Files” pane can be used to specify a reference trace file (i.e. normal control) in which the user knows that no sequence variation is present. Using the same protocol as above, locate and import the reference files into the middle pane (see Notes 5 and 6). 4. The traces to be analysed by Mutation Surveyor are uploaded into the bottom “Sample Files” pane using the protocol set out above. 5. Once all the files have been added to the “Open Files” window, click “OK”. The files are then loaded and await processing.

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Prior to analysis, the analysis parameters should be defined: 1. Click on “Options” in the Process menu. 2. Under the Contig tab, ensure that “Genbank/reference comparison” is unticked, “BasePatch” is ticked and “Exclude 1st Base Difference” is ticked and set at 100 (see Note 7). 3. Under the Mutation tab, ensure that the “Allow computer to Delete Mutations” box is unticked. All other default settings can remain unchanged. 4. The Output tab provides the options for the information that will be available on the various mutation reports. We select the following boxes for display within the mutation report: Sample file name, Reference File Name, Lane Quality, Amino Acid and ROI Only. 5. Under the “Display” tab, tick the “Check” box that relates to the “Check 2D Small Peaks (mosaic) feature” (see Note 8). 6. Under the “2 directions” tab, ensure that the “1 Direction” setting is selected for unidirectional data or “2 directions” for bidirectional data and then click OK. 7. To begin processing, click on “Run” in the Mutation Surveyor tool bar. Once the files have been processed, the analysed data is presented in a hyperlinked table called the mutation report (see Notes 9–11).

3.3. Data Analysis

Each variant highlighted in the mutation report should be checked: 1. Click on a variant. This will take you to the electropherogram in a comparison window (see Note 12). 2. Once the variant has been viewed, right-click on the name of the mutation which can be found in the mutation box at the bottom of the window and select either “Confirm” or “Delete Mutation”. 3. Return to the Mutation Report, by clicking the link in the column on the left of the screen, and repeat steps 1–3 for each variant identified.

3.3.1. Mutation Detection and Designation 3.3.1.1. Viewing Insertion/ Deletion Frameshift Mutations

Frameshift mutations are hyperlinked to a “heterozygous indel detection” window in all output tables (see Note 13). If a frameshift mutation is identified, then: 1. Click on the hyperlink to view the trace. 2. Zoom in and scan the sample electropherogram for the presence of any additional peaks in the comparison trace downstream of the frameshift.

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3.3.1.2. Missense, Nonsense, and Splicing Mutations

Missense, nonsense, and splicing mutations will be highlighted in the mutation report and can be viewed by clicking on the variant. Nomenclature is defined according to the HGVS guidelines for sequence variants (http://www.hgvs.org/).

3.3.2. Visual Inspection

The whole ROI can be checked manually: 1. Click on the first sample in the mutation report. 2. Scroll to the start of the ROI (as indicated by the pink line) and then scan along the length of the sequence to check the quality of the sequencing data. 3. Return to the Mutation Report, by clicking the link in the column on the left of the screen, and repeat steps 1–3 for each individual sample file.

3.3.3. Semi-automated Sequence Analysis

After checking the Mutation Report (see Subheading 3.3), it is possible to generate a “HGVS” report, which will provide detailed tabulated information regarding the quality of the sequence: 1. Click on “HGVS Output” in the Reports menu. 2. A new window will appear. Ensure that only the “Reject Mutations with Zero Score in Another Direction” box is ticked and then click “OK”. 3. Within the HGVS report window, click on the Output settings symbol (denoted by a spanner overlying a hammer). Ensure that the Phred Threshold score is set at 20, the Trace Quality is set for the ROI and the signal-to-noise ratio (S/N) has a Quality Threshold of 25. It is also recommended that HGVS nomenclature be ticked (see Note 14). 4. Press “OK”. The HGVS table will now appear (see Note 15). 5. In the HGVS output table, check that the “ROI covered” box contains a “Yes” (see Note 16). 6. The “Bases below threshold” column shows any bases within the ROI that fall below the Phred score. These bases should be checked by examining the corresponding electropherogram along with any traces where the quality score for the ROI falls below 25; these are indicated by red highlighting.

3.4. Confirming and Reporting Mutations

Mutation Surveyor offers a variety of options for reporting sequencing analysis. Tables can be generated which list all or specific types of detected variations and can provide details about the traces and each mutation. Various nomenclatures are available to display mutations based on the guidelines set out by the HGVS or by standard genomic numbering. Reports can be generated and may be customised to include electropherograms of the variants. These features are available through the Reports menu.

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The analysed data can be saved in three main formats: as a Mutation Surveyor project, a link to the data files, or as output data tables (see Note 17). To save the data: 1. Go to file and save in the preferred format.

3.6. Other Features

There are many additional features that Mutation Surveyor offers for sequence analysis, ranging from sequence annotation to mosaic quantification. Information on each of these features is outside the realms of this chapter. However, this information and further details on analysis parameters can be found in the handbook that accompanies the Mutation Surveyor Software package.

4. Notes 1. The main advantage of using the downloadable GenBank files is that the user is able to utilise the data contained within the files. GenBank files are annotated with details regarding the coding regions of the gene, amino-acid numbering, regions of interest, and they also provide information on reported sequence variants and polymorphisms which the user can choose whether or not to download. Not all GenBank-reported variants have been validated; therefore, it is the responsibility of the user to ensure that the variants incorporated into the reference file are not pathogenic. This is highlighted in the example of autosomal recessive disorders where heterozygous carriers of mutations are unaffected. 2. If a GenBank file is not available for the gene of interest, it is possible to convert a text sequence into GenBank format. A simple text file can be pasted into a ReadSeq conversion tool window that can be found on the website http://www.ebi. ac.uk/cgi-bin/readseq.cgi. Of the “Output sequence format” drop-down menu, select the “GenBank” sequence format. A GenBank formatted text sequence is then generated and can be saved using the “Save As” option (.txt) of the “File” menu. This newly generated file can then be used in Mutation Surveyor’s GenBank file editor, where features such as the ROI can be defined and any known variants added, before saving it in a .gbk file format and using the file for analysis. 3. The sequence window displays the reference base sequence, with the annotated exons in red. This is depicted graphically by the yellow bar at the bottom with annotated sequence variants shown as lines above the bar. The annotated features from the GenBank file are extracted and presented in the user interface within the “Features2” tab. The interface allows the user to delete or add any further sequence annotation.

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4. The ROI can be expanded by the user to include additional sites of interest within the intron (e.g. branch sites) or can be decreased to allow for more concentrated analysis and eliminate analysis of sequence outside the user’s ROI. 5. The types of sequence data file that are accepted by Mutation Surveyor are ABI sequencer data files (.ab1/.abi), those for a standard file format generated from sequencing instruments (.scf) and those compressed into a .gz format. 6. Multiple reference sequence files can be uploaded at once into the reference and sample panes. This feature makes it possible to analyse multiple exons at a time and to analyse unidirectional and bidirectional sequencing. The majority of exons will be sequenced in the forward direction in a single reaction; however, larger exons may be amplified and sequenced in overlapping fragments, or conversely smaller exons with short intronic sequences may be amplified and sequenced in a single reaction. For each sequence, there should be a corresponding reference file. If no normal control traces are available, Mutation Surveyor can still carry out the analysis by generating a synthetic control trace based on the specified GenBank reference file generated. 7. The GenBank Reference Comparison determines the reference files used for the analysis. If this option is ticked, then the software will use the GenBank sequence as a reference file; if unticked, the program will utilise the reference files to compare sequences. The BasePatch allows the software to correct for poor mobility within the sample traces, and by setting the “Exclude first Base Difference” to 100 bp, the program will ensure that all sequences that contain overlapping regions are grouped into separate contigs. 8. The Mutation Surveyor default settings may miss low-level mosaic mutations. Low-level mosaic mutations can be detected within two directional sequence data by using the mosaic detection option in the settings (“Check 2D Small Peaks”). When this option is chosen, low-level mosaics will be highlighted by a green dot above the mutation electropherogram. 9. We advise that the mutation report be checked to confirm that all sample files are present and that each sample file matches the correct Genbank reference file. 10. The Mutation Report provides a quality score for each sample file and lists all of the variants identified by Mutation Surveyor. Known polymorphisms are highlighted in lilac boxes, with mutations written in red or blue text. Mutations written in red text are considered to be of low confidence, while those in blue are of high confidence.

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11. Mutations/polymorphisms are called when comparing a test sample trace to a reference control trace. The classification of a sequence variant is dependent on the satisfaction of four para meters: mutation height, overlapping factor, intensity dropping factor and the signal-to-noise ratio: (a) Mutation peak height is the maximum peak height of the mutation in the electropherogram. (b) Signal-to-noise ratio is used to determine the confidence of the peaks and calculated using a Gaussian distribution, assuming that the median value is the noise and the highest value is the signal. (c) Overlapping factor is a measure of relative shift of the two peaks at the mutation position in the horizontal direction. (d) Dropping factor is the drop in height of the normal peak at the position of the mutation relative to the neighbouring peaks. (e) Mutation score is used to call a mutation and rank its confidence level. The mutation score is a probability of error and is derived from the ratios of signal-to-noise level, the dropping factor, and overlapping factor, expressed as: The mutation score = −10 log (error probability)

  s / n  = 110 log erfc    √ 2    where erfc (x) is the complementary error function. Accuracy is defined as 100% minus the error percentage, where the highest possible confidence, 99.9%, corresponds to a mutation score of 30. A score of 20 corresponds to 99% accuracy; a score of 10 corresponds to 90% accuracy.

(f ) Poor-quality sequence from the beginning and the end of a sequence read can be trimmed. This is depicted as small vertical blue bars displayed in the mutation electropherogram. The portion of the sequence that is trimmed is based upon having a low signal-to-noise ratio. The Quality Trim is a default option that can be overridden in the Mutation Surveyor settings. 12. The comparison window includes an electropherogram of the sample sequence and the normal reference sequence. Differences between these electropherograms are represented in the trace comparison view. A yellow line in the trace comparison window highlights the exonic sequence, whilst a beige line denotes intronic sequence. The ROI is represented by a pink line. The annotated GenBank sequence is provided at the top of the window where information regarding nucleotide and amino-acid numbering can be found.

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13. Mutation Surveyor cannot call base substitutions that are contained within a heterozygous frameshift trace. For autosomal recessive disorders, it is, therefore, necessary to carry out an additional visual inspection for any heterozygous frameshift trace for patients in whom a second pathogenic mutation has not been identified, in case the second mutation has been masked by the frameshift. The “Heterozygous indel detection” feature allows a frameshift trace to be examined. Base substitutions within this region can be confirmed by sequence analysis of the reverse strand. 14. Phred is a base-calling program that assigns scores based on the probability that the base call is correct. A Phred score of 20 represents 99% likelihood that the call is correct. Mutation Surveyor uses a modified algorithm for determining Phred scores which employs peak spacing and peak ratio but not peak resolution. 15. The main HGVS-tabulated output measures are:

(a) Number: trace number by order of processing.

(b) Sample File: name of sample file.

(c) Reference File: name of the reference sequence file used in the analysis.

(d) Quality ROI (Entire): a measure of the average signal-tonoise ratio where a score of 50 represents ≤1% noise.

(e) Read Start : the number of the base at the start of the processed data.

(f) ROI Start : the number of the base at the start of the ROI.

(g) Read End: the number of the base at the end of the processed data.

(h) ROI End: the number of the base at the end of the ROI.

(i) ROI Covered Yes or No.

(j) Average Phred (ROI).

(k) Range of Phred (ROI).

(l) Bases Below Threshold (ROI).

(m) Quality (lane quality): it is a measure of the average signal-to-noise ratio where a score of 50 represents ≤2% noise.

(n) Mut#: the number of mutations found in this sample sequence, where −1 indicates “Bad Data”.

(o) Mutation: the abbreviated name for each mutation in HGVS format. 16. If the ROI has not been covered, the electropherogram should be manually inspected. It is likely that the sequence

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reaction has failed or that the sample sequence contains a frameshift mutation. 17. When saving the data as a Mutation Project (.sgp), all the data are preserved and compressed into a single file for viewing at a later date. This option allows the user to repeat the analysis under different settings, whereas the “File Link” (.prj) is an image file of the analysed data. The output data table can be saved in various formats (.txt, .xls, .htm, .xml) for export to programs such as Microsoft Excel. References 1. McKusick, V. A. (2007) Mendelian inheritance in man and its online version OMIM. Am J Hum Genet 80, 588–604.

2. Ellard, S., Shields, B., Tysoe, C., Treacy, R., Yau, S., Mattocks, C., Wallace, A. (2009) Semi-automated unidirectional sequence analysis for mutation screening in a clinical diagnostic setting. Genet Test 13(3), 381–386.

Chapter 11 Non-invasive Prenatal Diagnosis Cathy Meaney and Gail Norbury Abstract The discovery of cell-free fetal DNA in the maternal plasma of pregnant women has facilitated the development of non-invasive prenatal diagnosis (NIPD). This has been successfully implemented in diagnostic laboratories for Rhesus typing and fetal sex determination for X-linked disorders and congenital adrenal hyperplasia (CAH) from 7 weeks gestation. Using real-time PCR, fluorescently labelled target gene specific probes can identify and quantify low copy number fetal-specific sequences in a high background of maternal DNA in the cell-free DNA extracted from maternal plasma. NIPD to detect specific fetal mutations in single gene disorders, currently by standard PCR techniques, can only be undertaken for paternally derived or de novo mutations because of the background maternal DNA. For routine use, this testing is limited by the large amounts of cell-free maternal DNA in the sample, the lack of universal fetal markers, and appropriate reference materials. Key words: Cell-free DNA, Cell-free fetal DNA, Maternal plasma, Non-invasive diagnosis, Real-time PCR, Single gene disorders

1. Introduction Traditionally, invasive procedures such as amniocentesis and chorionic villus sampling (CVS) are used for prenatal diagnosis from about 11 weeks gestation; however, both are associated with a small but significant risk of miscarriage. The detection and quantification of cell-free fetal DNA (cffDNA) in the plasma from pregnant women (1) has provided a non-invasive approach for prenatal diagnosis and at an earlier stage than invasive analysis. The amount of cffDNA increases from early pregnancy of around 25 genome equivalents per millilitre (GE/ml) to 300 GE/ml in late pregnancy, corresponding to around 3% of the total cell-free DNA (cfDNA) and increasing to an average of 6% (3). The fetal cfDNA sequences are thought to be mostly less than 300 bp in size


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compared with maternal cfDNA being predominantly greater than 500 bp (4, 5) and are rapidly cleared from the maternal plasma after delivery (6). NIPD using cffDNA has been successfully used for prenatal screening of women from 7 weeks gestation for Rhesus D (RhD) blood group typing in at risk RhD-negative women (2, 7, 8) and sex determination for sex-linked disorders and congenital adrenal hyperplasia causing ambiguous genitalia. Currently the most widely used technology for cffDNA analysis is real-time quantitative PCR with fluorescently labelled probes providing a closed system to help reduce contamination. The fluorescent probes provide greater specificity and sensitivity than double-stranded DNA binding dyes, such as SYBR Green, that will produce a signal for both specific and non-specific PCR products. DNA amplification occurs when the target gene is present and is quantified by the threshold cycle (Ct), which is the PCR cycle at which fluorescence is significantly higher than the background level. The Ct value is directly proportional to the amount of target gene present and allows determination of the amount of the fetal or total (maternal and fetal) cfDNA. For fetal sex determination, this provides a sensitive technique for the detection of low copy numbers of fetal SRY male sequences against the high background of maternal cfDNA. Lack of amplification is suggestive of a female fetus but cannot be distinguished from a false-negative result arising from the failure to extract or detect the very low (single copy) levels of fetal cfDNA present in the sample aliquot. Amplification of a locus/house keeping gene, such as CCR5, will indicate if a low amount of total cfDNA is extracted and therefore likely that there will be inadequate amounts of fetal DNA in the plasma. Fetal ultrasound is an important adjunct test to help identify additional gestational sacs in multiple pregnancies that if discordant for the trait could potentially interfere with determining the correct fetal genotype. NIPD for single gene disorders is currently limited to the detection of paternally inherited or de novo fetal mutations and has been reported for various disorders including cystic fibrosis (9), achondroplasia (10), and b-thalassaemia (11–13). To help overcome the complication arising from the high background maternal DNA, techniques with high specificity and sensitivity have been used that include mass spectrometry (12), PCR using peptide nucleic acid (PNA) clamps (13), and enrichment of cffDNA by size fractionation (14). More recent technology such as microfluidics digital PCR may provide alternative and more sensitive methods for analysis of cffDNA but high costs may be a limitation (15). For routine diagnosis, however, more convenient and robust approaches are required. The use of small amplicons to improve the efficiency of amplification and the testing of multiple replicates may help to improve reliability. Development of appropriate reference materials and universal fetal markers to confirm the presence of fetal DNA will assist further in this analysis.

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Described below are methods for the extraction of cffDNA by both a manual and semi-automated procedure and for genotyping both by real-time PCR and by standard PCR and restriction analysis.

2. Materials 2.1. Maternal Blood Sample

1. Potasssium EDTA blood, 10–20 ml (biohazard).

2.2. Cell-Free DNA Extraction from Maternal Plasma

1. Absolute ethanol (highly flammable).

2.2.1. Qiagen QIAamp MinElute Virus Spin Kit (50)

2. Sterile-filtered water (see Note 1). 3. QIAamp MinElute Virus Spin Kit 50 (Qiagen cat 57704) includes reagents noted in items 3-10: QIAamp MinElute columns (50), store at 2–8°C. 4. Buffer AL (12 ml), store at room temperature. Care with handling as contains a chaotropic salt, guanidine hydrochloride (harmful irritant, do not mix with bleach). 5. Buffer AW1 (19 ml): add 25 ml of 96–100% ethanol and mix well each time before use. Working solution is stable for 1 year when stored at room temperature. Care with handling as contains a chaotropic salt, guanidine hydrochloride (harmful irritant, do not mix with bleach). 6. Buffer AW2 (13 ml): add 30 ml of 96–100% ethanol. Store at room temperature and mix well each time before use. 7. Buffer AVE (5 × 2 ml): store at room temperature. Contains sodium azide (highly toxic). 8. Protease resuspension buffer (6 ml), store at room temperature. Contains sodium azide (highly toxic). 9. Qiagen protease (lyophilised, store at room temperature) is dissolved in 1.4 ml protease resuspension buffer. Carefully mix to avoid frothing and store at 2–8°C for up to 1 year, mini mising extended periods at room temperature when in use. Care with handling as contains subtilisin (a sensitizer and irritant). 10. Carrier RNA (lyophilised, 310 mg): dissolve in 310 ml Buffer AVE to give a 1 mg/ml solution. Store single use aliquots at −20°C for up to 1 year to avoid freeze–thawing (see Note 2).

2.2.2. Qiagen EZ1 Virus Mini Kit (48) and EZ1 Robot

1. BioRobot EZ1 workstation and EZ1 Virus card (Qiagen). EZ1 Virus Mini Kit (Qiagen cat 955338) for 48 extractions includes the reagents listed below: 2. Reagent cartridges containing magnetic particles, lysis buffer, wash buffer, and RNase-free elution buffer are stored at room

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temperature. Care when handling cartridges as reagents contain guanidine salts (harmful irritant, should not be mixed with bleach), sodium azide (highly toxic), and ethanol and isopropanol (highly flammable). If precipitation of Buffer AL in reagent cartridge occurs then redissolve by warming at 37°C. 3. Protease resuspension buffer (6 ml), store at room temperature. Contains sodium azide (highly toxic). 4. Qiagen protease (lyophilised, store at room temperature) is dissolved in 4.4 ml protease resuspension buffer. Carefully mix to avoid frothing and store at 2–8°C for up to 1 year, minimising extended periods at room temperature when in use. Care with handling as contains subtilisin (a sensitizer and irritant). 5. Carrier RNA (lyophilised, 310 mg) is dissolved in 1000 ml Buffer AVE to give a 0.31 mg/ml solution. Store single use aliquots at −20°C for up to 1 year to avoid freeze–thawing (see Note 2). 2.3. Fetal Sex Determination Using Applied Biosystems 7300 Real-Time PCR System (AB7300)

1. TaqMan Universal PCR Master mix (no AmpErase UNG; uracil-N-glycosylase) contains the PCR components AmpliTaq Gold DNA polymerase and Buffer A with Passive Reference I Rox dye (Applied Biosystems, see Note 3). Store at 4°C. 2. TaqMan Gene Expression Assay SRY, 20× mix (Applied Biosystems, see Note 4) and TaqMan Gene Expression Assay CCR5, 20× mix (Applied Biosystems, see Note 5) include primers (18 mM each) and a TaqMan MGB probe (5 mM) labelled with a 5′ FAM reporter dye and a 3′ non-fluorescent quencher. Store in single use aliquots at −20°C. 3. Human pooled male genomic DNA (Promega) is diluted to 20 ng/ml and aliquots stored at −20°C. For the SRY and CCR5 standard curves, prepare tenfold serial dilutions from the 20 ng/ml aliquot in sterile water to 2, 0.2, and 0.02 ng/ml. Store at −20°C and use aliquots not more than three times. 4. Human pooled female genomic DNA (Promega) is diluted to 1 ng/ml and aliquots stored at −20°C.

2.4. Detection of De Novo or Paternal Mutations for Single Gene Disorders

See Subheading 2.2.1 for reagents.

2.4.1. Qiagen QIAamp MinElute Virus Spin Kit (50) 2.4.2. Standard PCR

1. AmpliTaq Gold PCR Master Mix (2×) includes AmpliTaq Gold DNA Polymerase (0.05 U/ml), Gold Buffer (100 mM KCl, 30 mM Tris–HCl, pH 8.0), 400 mM each dNTP and 5 mM MgCl2. Store at 4°C or −20°C (Applied Biosystems).


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2. AmpliTaq Gold DNA Polymerase (5 U/ml) with 25 mM MgCl2 solution and either 10× Buffer II (500 mM KCl, 100 mM Tris–HCl, pH 8.3) or 10× Gold Buffer (500 mM KCl, 150 mM Tris–HCl, pH 8.0). Store at −20°C (Applied Biosystems). 3. Deoxyribonucleotide triphosphates (dNTPs, 2 mM). 4. Appropriate primers (100 microM each). 5. Sterile-filtered water. 2.4.3. Restriction Enzymes, Agarose Gels, and Polyacrylamide Gel Electrophoresis

1. Bovine serum albumin (BSA, 100 mg/ml) and various restriction enzymes and 10× buffers stored at −20°C. 2. Agarose, standard. 3. AccuGel 40% (w/v) 19:1 acrylamide/bis solution (National Diagnostics; use care with exposure as this is a neurotoxin when unpolymerised). 4. N,N,N ′-tetramethyl-ethylenediamine highly flammable, see Note 6).

(TEMED,

Sigma,

5. Ammonium persulphate: prepare a 10% solution in water and store at 4°C. Use within 1 week. 6. Pre-mixed 10× TBE buffer (1.1 M Tris, 0.9 M boric acid, and 0.01 M ethylenediamine tetraacetic acid). 7. Ethidium bromide (10 mg/ml, powerful mutagen). 8. Glycerol loading dye: 50% glycerol/water, 0.25% bromophenol blue (irritant), and 0.25% xylene cyanol FF (irritant). Store at room temperature. 9. Molecular weight markers: 10 and 50 bp markers diluted to 50 ng/ml in water and glycerol loading buffer. Store at −20°C.

3. Methods 3.1. Maternal EDTA Blood Sample Collection and Separation

1. For cell-free DNA (cfDNA) extraction and analysis, generally 10–20 ml of maternal EDTA blood is required. Samples should be spun and plasma separated preferably within 12-24 h of collection (see Note 7). 2. Centrifuge the blood sample for 15 min at 3,000 × g at room temperature. 3. Carefully aspirate the top plasma layer into a new tube using a sterile pastette. The buffy coat layer contains the maternal white blood cells and can be carefully removed for the extraction of maternal DNA if required. 4. Centrifuge the plasma for another 15 min at 3,000 × g and carefully aspirate the top plasma layer into a fresh tube.

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5. Aliquot plasma into labelled microcentrifuge tubes. Keep aliquots at 4°C if to be tested that day otherwise store at −20 to −80°C. 3.2. Cell-Free DNA Extraction from Maternal Plasma 3.2.1. Manual cfDNA Extraction Using Qiagen QIAamp MinElute Virus Spin Kit (50)

Reference: QIAamp MinElute Virus Spin Kit instruction manual (16). 1. Prepare reagents as recommended in section 2.2.1 and in a safety cabinet if possible (see Note 8 & 9). 2. Add 50 ml Qiagen protease into a labelled 2-ml microcentrifuge tube and then transfer 400 ml plasma into this tube (see Note 10). 3. Prepare each time a fresh mix of Buffer AL and carrier RNA-Buffer AVE for the number of samples to be extracted according to Table 1. To avoid frothing, mix gently by inverting the tube. 4. Add 400 ml of Buffer AL/carrier RNA mix into the 2-ml tube and mix thoroughly by vortexing for 15 s. 5. Incubate at 56°C for 15 min then, to limit cross-contamination, pulse-spin to remove drops on the lid. 6. Add 500 ml of ethanol (96–100%) to the lysate and vortex for 15 s. Incubate at room temperature for 5 min then pulse-spin. 7. Pipette half of the lysate onto a QIAamp MinElute column (important not to wet rim to avoid cross-contamination) and centrifuge at 6,000 × g for 1 min. Discard the collection tube and filtrate, and put the column into a clean QIAamp 2-ml collection tube. Repeat this step with the remainder of the lysate and put in a clean QIAamp 2-ml collection tube. 8. Add 500 ml of Buffer AW1 to the QIAamp MinElute column and centrifuge at 6,000 × g for 1 min. Discard the collection

Table 1 Preparation of the Buffer AL and carrier RNA-Buffer AVE mix according to the number of samples to be extracted for the QIAamp MinElute Virus Spin method Sample number

Buffer AL volume (ml)

Carrier RNA-AVE volume (ml)

1

0.44

12.3

2

0.88

24.6

3

1.32

37.0

4

1.76

49.3


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tube and filtrate, and put the column into a clean QIAamp 2-ml collection tube. 9. Add 500 ml of Buffer AW2 to the column and centrifuge at 6,000 × g for 1 min. Discard the collection tube and filtrate, and put the column into a clean QIAamp 2-ml collection tube. 10. Add 500 ml of ethanol (96–100%) to the column and centrifuge at 6,000 × g for 1 min. Discard the collection tube and filtrate, and put the column into a clean QIAamp 2-ml collection tube. To dry the membrane of the column, centrifuge at 20,000 × g for 3 min. 11. Cut the lid off a 1.5-ml microcentrifuge tube and place the column into this tube. Add 75 ml of sterile water to the centre of the membrane and incubate at room temperature for 5 min (see Note 11). Centrifuge at 20,000 × g for 1 min and store cfDNA at 4°C or on ice until ready to use. 3.2.2. Semi-automated cfDNA Extraction Using Qiagen EZ1 Virus Mini Kit (48) and BioRobot EZ1

Reference: Qiagen EZ1 Virus Mini kit instruction manual (17). These instructions are specific for cfDNA extraction on the Qiagen BioRobot EZ1; however, other kits and robots can e used. 1. Prepare reagents as recommended in section 2.2.2 and in a safety cabinet if possible (see Note 8 & 9). A maximum of six samples can be extracted at once on the BioRobot EZ1. 2. Add 75 ml Qiagen protease solution and 10 ml carrier RNA to a 1.5-ml Qiagen EZ1 tube and mix gently by pipetting (do not vortex as it will froth). 3. Transfer 400 ml plasma into a labelled 2-ml EZ1 tube. 4. Insert the EZ1 Virus Card into the EZ1 card slot on the BioRobot EZ1 and then turn on the BioRobot EZ1. Wipe probes with ethanol. 5. Press “START” and from the Protocols menu select 400 ml plasma and an elution volume of 75 ml (see Note 12). 6. Remove the cartridge rack and sample tray from the BioRobot EZ1. Invert the reagent cartridge until the magnetic particles are mixed into the solution, slide the reagent cartridge into the cartridge rack and load back into the BioRobot EZ1. 7. For each sample, load an empty 1.5-ml EZ1 tube (discard lid) into the heating block at the back of the cartridge rack. 8. In row 1 of sample tray, load empty and labelled 1.5-ml EZ1 elution tubes (put lids aside). 9. In row 2 of sample tray, load EZ1 tip holders containing EZ1 filter tips. 10. In row 3 of sample tray, load 1.5-ml EZ1 tubes with 75 ml protease and 10 ml carrier RNA.

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11. In row 4 of sample tray, load labelled 2-ml EZ1 tubes containing 400 ml plasma. 12. Close the BioRobot EZ1 door and press “START” to begin the protocol. When the protocol has finished, replace the labelled lids onto the elution tubes and store at 4°C until ready to use. 3.3. Fetal Sex Determination Using Cell-Free DNA and Real-Time PCR Analysis 3.3.1. Applied Biosystems 7300 Real-Time PCR System (AB7300) and Plate Setup

These instructions are specific for performing real-time PCR using the AB7300 but can be adapted for use on other real-time PCR machines. 1. If possible setup the PCR reactions in a safety cabinet (see Note 13). 2. Prepare a master mix for the number of samples and controls testing for a 20-ml PCR using 1 ml SRY or CCR5 20× TaqMan Gene Expression Assay, 10 ml of TaqMan Universal PCR Master mix, and 4 ml sterile water. The volume of cfDNA added to the PCR can be varied if needed. 3. Pipette 15 ml of SRY master mix into a 96-well microtitre plate for the number of samples to be tested (×3–8 replicates each) and for the no template control (NTC, three replicates), male and female controls (three replicates each), and tenfold serial dilution male gDNA standards (three replicates for each standard). See an example of a plate setup in Fig. 1. 4. Pipette 15 ml of CCR5 master mix into a 96-well microtitre plate for the number of samples to be tested (×2–3 replicates each) and for the NTC (three replicates) and tenfold serial dilution male gDNA standards (three replicates for each standard). 5. Pipette 5 ml of cfDNA, control gDNA and water, and serialdiluted gDNA into the appropriate wells of the microtitre plate (see Note 14). 6. Place a plastic seal over the plate taking care not to touch the area covering the wells with gloves and ensuring there is no dust on the seal. Spin the microtitre plate in a plate centrifuge at 1,500 × g for 2 min to remove any bubbles. 7. Setup the plate worksheet on the computer software; start a new sample sheet then select and add the SRY detector for the SRY wells and CCR5 detector for the CCR5 wells, with the passive reference set to ROX (see Note 15). 8. For the cfDNA wells, type in the sample name and select “UNKNOWN,” for the water controls select “NTC,” for male and female controls select “UNKNOWN,” and for the male gDNA standard curve dilutions select “STANDARD” and add the concentration of each dilution.

Non-invasive Prenatal Diagnosis 1

2

3

4

5

6

7

8

9

10

11

163 12

A B UNKN UNKN UNKN UNKN UNKN UNKN

UNKN UNKN UNKN

UNKN UNKN UNKN UNKN UNKN UNKN

UNKN UNKN UNKN

C D E NTC

NTC

NTC

NTC

NTC

NTC

MALE

MALE

MALE

FEM

FEM

FEM

S1

S1

S1

S2

S2

S2

S3

S3

S3

S4

S4

S4

S1

S1

S1

S2

S2

S2

S3

S3

S3

S4

S4

S4

F G H

= SRY

= CCR5

Fig. 1. Example of a plate setup for real-time PCR analysis on an AB7300 machine of two cell-free DNA samples. Rows B and C (UNKN) = cfDNA samples; wells E1-3 (NTC) and E10-12 (NTC) = no template control; wells F1-3 (MALE) = 100 pg male gDNA control; wells F10-12 (FEM) = 100 pg female gDNA control; rows G and H = male standards for standard curve, where S1 = 100 ng male gDNA, S2 = 10 ng male gDNA, S3 = 1 ng male gDNA, and S4 = 0.1 ng male gDNA.

9. Load the plate onto the AB7300 System and press “start.” PCR conditions are: 1 cycle of 95°C for 10 min, 45 cycles of 95°C for 15 s, 60°C for 1 min, and 1 cycle held infinitely at 4°C. 3.3.2. Analysing Results on the Applied Biosystem 7300 Real-Time PCR System

1. Amplification data collected by the AB7300 Sequence Detector is analysed using the sequence detector system (SDS) software. After the run has completed, set the thres hold to 0.2 and analyse the run. 2. Both the report table of Ct values and the corresponding amplification graphs should be inspected and compared to confirm if there has been amplification or no amplification (see Figs. 2 and 3). 3. The run should be validated in accordance with internal criteria. Guidelines for the analysis and reporting of results are shown in Table 2. 4. The standard curves can calculate the amount of fetal and total DNA present in the cfDNA extract and are generated from creating a plot of Ct vs. log of the concentration (or copy number) of the tenfold serially diluted male gDNA.

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Fig. 2. (a ) Graph showing SRY amplification and control CCR5 amplification of the total cfDNA in the maternal plasma indicating a male fetus. (b ) Amplification with the CCR5 control and lack of amplification with SRY indicates a female fetus.

The gradient is generally between −3.3 and −4.5 and the R2 value greater than 0.99 indicate the accuracy of pipetting (see Note 16 and 17). 3.4. Detection of De Novo or Paternally Derived Disease Mutations

Confirmation of the presence of fetal DNA in the cfDNA extracted from plasma may first be undertaken by fetal sexing using real-time PCR analysis as in Subheadings 3.2 and 3.3 (see Note 18).


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Fig. 3. (a ) SRY amplification of the tenfold serial-diluted male gDNA standards and fetal cfDNA (amplification indicating male fetus) and (b ) corresponding standard curve of the threshold cycle (Ct) against the input log of the target quantity (ng) that allows for quantification of DNA in maternal plasma.

3.4.1. Extraction of Cell-Free DNA from Maternal Plasma

1. Extract cfDNA from maternal plasma using the manual Qiagen QIAamp MinElute Virus Spin Kit (50) as in Subheading 3.2.1. However, for each patient, 2 × 400 ml plasma aliquots may be extracted using two separate spin columns for each aliquot and the eluate pooled at the end of the procedure (see Note 19).

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Table 2 Guidelines for quality control validation and reporting of results Quality control validation

Results interpretation

Positive result

PCR amplification (eg. Ct 40 or 0) with SRY cfDNA replicates (and amplification with CCR5 cfDNA replicates)

Consistent with female fetus if SRY cfDNA replicates do not amplify, however, the presence of fetal DNA is not confirmed

Positive controls

PCR amplification (eg. Ct 40 or 0) with any SRY and CCR5 NTC replicates or SRY female replicates

Repeat if any NTC or female gDNA replicates amplify

Gradient (slope) of standard curves

Between −3.3 and −4.5

R2 value

Greater or equal to 0.99

cfDNA cell-free DNA, NTC non-template control using sterile water

2. Two times AW2 washes may be necessary to help remove any excess salt (see Note 20). 3. After the final spin at 20,000 × g for 3 min, add 55-ml sterile water to one of the dried spin columns and keep at room temperature for 5 min. Spin at 20,000 × g for 1 min. Take the 55 ml eluate and apply to the second dried spin column. Keep at room temperature for 5 min and spin at 20,000 × g for 1 min. Keep at 4°C until ready to use on that same day. 3.4.2. Standard PCR Using Cell-Free DNA

1. Prepare a master mix for the number of samples and controls testing for a 50 ml PCR volume with final concentrations of 1.5–3 mM MgCl2, 200 mM of each dNTP, 10 pmol of each primer, 1× AmpliTaq Gold Buffer or Buffer II, and 0.5–2.5 U AmpliTaq Gold DNA polymerase (see Note 21). 2. Mix thoroughly and pulse-spin. Add the appropriate amount of master mix to each PCR tube. 3. Add 1–10 ml of cfDNA and adjust water accordingly to make up to 50 ml. Add 1 ml of the appropriate control DNA (wild-type, positive, and NTC). 4. Place tubes in a heated lid thermal cycler and run a touch down PCR of 57 cycles: conditions of one cycle of 95°C for 5 min, then a temperature gradient of seven cycles


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(95°C for 15 s, start annealing temperature 7°C above the optimal primer annealing temperature and decrease by 1°C/cycle for 15 s, 72°C for 30 s). Follow with 50 cycles of 95°C for 15 s, the optimal primer annealing temperature for 15 s and 72°C for 30 s. Finish with 72°C for 10 min to complete the extension reaction (see Note 22). 3.4.3. Restriction Enzyme Digestion and Gel Electrophoresis

1. If restriction enzyme digestion is required for mutation detection, prepare a master mix for the number of samples and controls testing of the appropriate restriction enzyme, 10× buffer, BSA (if required) and sterile water for a final digest volume of 20–30 ml. Mix then add the appropriate amount of master mix to each digest tube. Add 10–20 ml of PCR (or water for the NTC control) and digest for 2-4 h or overnight at appropriate temperature. 2. Depending on the fragment size separation required, run on 3% agarose or 10–12% polyacrylamide gels. Prepare a 3% agarose gel by heating 450 g of agarose in 150 ml 1× TBE (dilute 100 ml of 10× TBE in 1 l of reverse osmosis water) and once cooled add 7.5 ml ethidium bromide. Run in 1× TBE running buffer at 100 V for 1–2 h. 3. For polyacrylamide gels, prepare a stock solution for 10% (or 12%) gels by mixing 125 ml (or 150 ml) AccuGel 40% (w/v) 19:1 acrylamide:bis acrylamide solution, 50 ml 1× TBE and make up to 500 ml with reverse osmosis water (see Note 23). 4. Wash two glass plates with detergent, rinse thoroughly with water, wipe with 70% ethanol, and place spacers between plates. Prepare a polyacrylamide gel by mixing 25-ml polyacrylamide stock solution with 200 ml ammonium persulphate and 34 ml TEMED (add in fume hood). Pour the mix between upright plates, insert the comb, and leave to set for 30–60 min. Once the gel is set, remove spacers and load into an upright gel tank, filling upper and lower chambers with 1× TBE running buffer. Remove the comb and wash wells with 1× TBE using a syringe. 5. Load 20–30 ml of the PCR/digest plus glycerol loading dye onto the gel, including 10 or 50 bp molecular weight markers. Run at 55 mA for about 1 h depending on the required size separation and stain for about 5 min with ethidium bromide (5 ml/100 ml 1× TBE). Xyalene cyanol runs at about 115 bp on a 10% gel and 70 bp on a 12% gel. 6. Detection of fetal DNA will be a faint band on the gel compared with the bands representing the total (maternal and fetal) cfDNA (see Figs. 4 and 5).

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Fig. 4. Detection of the paternally inherited torsion dystonia c.946delGAG (p.Glu302/303) mutation in the DYT1 gene from a maternal sample taken at (a ) 7 + 5 weeks gestation and (b ) 8 + 5 weeks gestation.

Fig. 5. Confirmation of the paternally inherited torsion dystonia c.946delGAG (p.Glu302/303) mutation in fetal tissue sample.


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4. Notes 1. The standard term “water” used in this text refers to sterilefiltered water that is appropriate for PCR procedures. All mixes and solutions are made with this water or, as stated, reverse osmosis water. 2. Carrier RNA enhances binding of small amounts of DNA and RNA to the silica and also takes up non-specific binding sites on the column therefore reducing the loss of DNA/RNA. Inclusion of carrier RNA may therefore increase the recovery of cffDNA. 3. Using TaqMan Universal PCR Master Mix with AmpErase UNG (uracil-N-glycosylase) and dUTP instead of dTTP may help to prevent carry-over contaminations of DNA from previous PCRs. An initial 2-min incubation at 50°C step at the beginning of the PCR will need to be included. 4. TaqMan MGB probes contain a minor groove binder at the 3¢-end that increases the melting temperature, permitting shorter probes, and therefore higher specificity. The TaqMan assays work at the annealing temperature of 60°C for ease of use. 5. CCR5 is the chemokine receptor gene present in both maternal and fetal genomes. Other control genes may be used, for example, b-actin or glyceraldehyde-3-phosphate dehydrogenase (GAPDH). 6. TEMED is best stored at room temperature in a desiccator. Use small bottles as after opening it may decline in quality and gels will take longer to polymerise. 7. Samples have been successfully analysed after several years of cfDNA storage at −20°C. Although cffDNA has been detected as early as 5 weeks gestation, the concentrations and sensitivity increase with advancing pregnancy. Analysis is therefore recommended from 7 weeks gestation and care must be taken with the possibility of multiple pregnancies or a vanished twin to ensure there is only one sac. 8. Use plasma aliquots only once as denaturation and precipitation of proteins occur when plasma is repeatedly freeze– thawed and may reduce the yield of cffDNA. To remove any precipitates in the thawed plasma sample, spin at 6,800 × g for 3 min and transfer supernatant to a new tube. 9. Prepare samples in a safety cabinet, wear powder-free gloves, use filter tips, and change tips between each transfer to avoid contamination. 10. QIAamp MinElute columns are optimised to use with a minimum volume of 200 ml plasma. However, 400 ml of plasma

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can be double loaded onto one column to increase the yield and the reagent volumes to add are therefore doubled. 11. The final eluate volume will be slightly less than that initially put onto the column so add slightly more than the required volume needed for downstream processes. 12. Final eluate volume can be 75, 100, 125, or 150 ml, however, using the lowest volume of 75 ml will concentrate the cfDNA extracted. 13. Real-time PCR reduces the risk of contamination as it is a closed system. It is, however, a very sensitive technique and if possible prepare SRY and CCR5 master mixes and subsequent loading onto a 96-well microtitre plate in a HEPA/UV PCR safety cabinet, preferably in a designated pre-PCR room. Prepare the master mixes before handling DNA (change gloves if necessary), use sterile filter tips, and decontaminate pipetmans using UV light for 20 min in HEPA cabinet after setup is completed. Minimise the exposure of the labelled DNA probe to light. 14. It is important to get the threshold cycle value for each replicate as close as possible for consistent results. Reverse pipetting can help to give more accurate and consistent amounts of DNA in each well, however, it is important not to take up too much extra cfDNA in the pipette tip due to the limited amount extracted. 15. The ROX passive dye is used to correct for pipetting inaccuracies; it does not take part in the PCR but provides an internal fluorescent reference that allows the reporter fluorescence to be normalised for non-PCR-related signal variation. 16. Due to the very low copy number of fetal DNA in maternal plasma, especially in the first trimester, accurate measurements may not be obtained. The concentration of DNA may be expressed as genome equivalents or genome copies per millilitre (GE/ml) and is calculated using the conversion factor of 6.6 pg of DNA per cell as described by Lo et al. (3) [Concentration in plasma (copies/ml) = Quantity (copies) of target in PCR determined by real-time PCR SDS detector × Vol DNA extracted/Vol DNA used in PCR × 1/Vol plasma extracted]. 17. Handling and addition of male gDNA to the microtitre plate wells after the addition of cfDNA and water (NTC) wells may help to reduce cross-contamination in wells and false-positive results. As the amount of fetal DNA is very low, testing multiple replicates and duplicate plasma samples, ideally at different gestations if under 9 weeks, is recommended as well as auditing the cell-free fetal sexing results with karyotyping, ultrasound, or birth outcome.


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18. Confirmation of fetal DNA will only be useful for male fetuses where there is amplification with SRY. Polymorphic marker analysis using real-time PCR may be undertaken to confirm fetal DNA, however, this is laborious and time-consuming and only informative in a small number of cases (18). 19. If necessary larger volumes of plasma and different extraction kits can be used to extract cfDNA. 20. If there are problems with amplifying the cfDNA and/or restriction digestion then undertaking two washes with AW2, which contains mostly ethanol, will help to remove any excess salt. 21. AmpliTaq Gold Master mix has all the PCR components pre-mixed and the AmpliTaq Gold DNA polymerase is a hot start enzyme activated at 95°C for 2–10 min with a 5¢–3¢ exonuclease activity that may help to reduce non-specific priming. Design primers to amplify a small PCR product under 200 bp as the majority of cffDNA fragments are thought to be less than 300 bp. 22. A high number of 57 PCR cycles are required as there is only a small amount of fetal DNA template. Touchdown PCR has a higher specificity and will help minimise mispriming and false amplification that may occur during standard PCR conditions (19). 23. The 10–12% stock solutions of polyacrylamide/bis, TBE and water, can be pre-made and stored in the fridge at 4°C in a bottle wrapped in foil as solution is light sensitive.

Acknowledgments Dr Kirsten Finning for her guidance in setting up the real time PCR analysis. Bhaneeta Mistry and Lighta Godinho for technical assistance. References 1. Lo YMD, Corbetta N, Chamberlain PF, Rai V, Sargent IL, Redman CW, Wainscoat JS. (1997) Presence of fetal DNA in maternal plasma and serum. Lancet 350, 485–487. 2. Finning KM, Martin PG, Soothill PW, Avent ND. (2002) Prediction of fetal D status from maternal plasma: introduction of a new noninvasive fetal RHD genotyping service. Transfusion 42, 1079–1085.

3. Lo YM, Tein MS, Lau TK, Haines CJ, Leung TN, Poon PM, Wainscoat JS, Johnson PJ, Chang AM, Hjelm NM. (1998) Quantitative analysis of fetal DNA in maternal plasma and serum: implications for noninvasive prenatal diagnosis. Am J Hum Genet 62, 768–775. 4. Li Y, Zimmermann B, Rusterholz C, Kang A, Holzgreve W, Hahn S. (2004) Size separation

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9.

10.

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12.

Meaney and Norbury of circulatory DNA in maternal plasma permits ready detection of fetal DNA polymorphisms. Clin Chem 50, 1002–1011. Chan KC, Zhang J, Hui AB, Wong N, Lau TK, Leung TN, Lo KW, Huang DW, Lo YM. (2004) Size distributions of maternal and fetal DNA in maternal plasma. Clin Chem 50, 88–92. Lo YMD, Zhang J, Leung TN, Lau TK, Chang AM, Hjelm NM. (1999) Rapid clearance of fetal DNA from maternal plasma. Am J Hum Genet 64, 218–224. Faas BH, Beuling EA, Christiaens GC, von dem Borne AE, van der Schoot CE. (1998) Detection of fetal RHD-specific sequences in maternal plasma. Lancet 352, 1196. Lo YM, Hjelm NM, Fidler C, Sargent IL, Murphy MF, Chamber-lain PF, Poon PM, Redman CW, Wainscoat JS. (1998) Prenatal diagnosis of fetal RhD status by molecular analysis of maternal plasma. N Engl J Med 339, 1734–1738. Nasis O, Thompson S, Hong T, Sherwood M, Radcliffe S, Jackson L, Otevrel T. (2004) Improvement in sensitivity of allele-specific PCR facilitates reliable noninvasive prenatal detection of cystic fibrosis. Clin Chem 50(4), 694–701. Li Y, Holzgreve W, Page-Chistiaens GC, Gille JJ, Hahn S. (2004) Improved prenatal detection of a fetal point mutation for achondroplasia by the use of size-fractionated circulatory DNA in maternal plasma – case report. Prenatal Diagn 24, 896–898. Chiu RWK, Lau TK, Leung T, Chow K, Chui D, Lo YM. (2002) Prenatal exclusion of beta-thalassaemia major by examination of maternal plasma. Lancet 360, 998–1000. Ding, C, Chiu RW, Lau TK, Leung TN, Chan LC, Chan AY, Charoenkwan P, Ng IS, Law HY,

Ma ES, Xu X, Wanapirak C, Sanguansermsri T, Liao C, Ai MA, Chui DH, Cantor CR, Lo YM. (2004) MS analysis of single-nucleotide differences in circulating nucleic acids: application to noninvasive prenatal diagnosis. Proc Natl Acad Sci USA 101, 10762–10767. 13. Li Y, Di Naro E, Vitucci A, Zimmermann B, Holzgreve W, Hahn S. (2005) Detection of paternally inherited fetal point mutations for beta-thalassaemia using size-fractionated cellfree DNA in maternal plasma. JAMA 293(7), 843–849. Erratum in: JAMA 2005 Apr 13;293(14), 1728. 14. Li Y, Zimmerman B, Rusterholz C, Kang A, Holzgreve W, Hahn S. (2004) Size separation of circulatory DNA in maternal plasma permits ready detection of fetal DNA polymorphisms. Clin Chem 50(6), 1002–1011. 15. Zimmerman BG, Grill S, Holzgreve W, Zhong XY, Jackson LG, Hahn S. (2008) Digital PCR: a powerful new tool for noninvasive prenatal diagnosis? Prenatal Diagn 28(12), 1087–93. 16. Qiagen method – http://www1.qiagen.com/ literature/handbooks/literature.aspx?id= 1000434. 17. Qiagen method – http://www1.qiagen.com/ literature/handbooks/literature. aspx?id=1000194. 18. Alizadeh M, Bernard M, Danic B, Dauriac C, Birebent B, Lapart C, Lamy T, Le Prise PY, Beauplet A, Bories D, Semana G, Quelvennec E. (2002) Quantitative assessment of hema topoietic chimerism after bone marrow transplantation by real-time quantitative PCR. Blood 99, 12. 19. Don RH, Cox PT, Wainwright BJ, Baker K, Mattick JS. (1991) ‘Touchdown’ PCR to circumvent spurious priming during gene amplification. Nucleic Acid Res 19(14), 4008.

Chapter 12 Automated DNA Sequencing Yvonne Wallis and Natalie Morrell Abstract Fluorescent cycle sequencing of PCR products is a multistage process and several methodologies are available to perform each stage. This chapter will describe the more commonly utilised dye-terminator cycle sequencing approach using BigDye® terminator chemistry (Applied Biosystems) ready for analysis on a 3730 DNA genetic analyzer. Even though DNA sequencing is one of the most common and robust techniques performed in molecular laboratories it may not always produce desirable results. The causes of the most common problems will also be discussed in this chapter. Key words: Fluorescent cycle sequencing, Applied Biosystems 3730 genetic analyzer, ExoSAP PCR clean-up, BigDye® sequencing chemistry, CleanSeq® post-sequencing clean-up, Ethanol precipitation, Sequencing troubleshooting

1. Introduction DNA sequencing is a technique used to precisely characterise the nature of sequence variants at the single base level. It can be used to characterise variants detected by pre-screening methodologies such as conformation-sensitive capillary electrophoresis (CSCE), denaturing high pressure liquid chromatography (dHPLC) and single-strand conformation polymorphism (SSCP) or it can be used as the screening tool itself. There are many advantages to using DNA sequencing as the primary screening tool: it is highly sensitive; the whole process is amenable to laboratory automation thereby facilitating high-throughput workflows; modern sequencing chemistries are robust and reliable tolerating a wide spectrum of template quality and producing even peak heights resulting in accurate heterozygote detection; it is possible to scale down reagents reducing the cost; it negates the need to optimise and maintain pre-screening techniques thereby simplifying laboratory Bimal D.M. Theophilus and Ralph Rapley (eds.), PCR Mutation Detection Protocols, Methods in Molecular Biology, vol. 688, DOI 10.1007/978-1-60761-947-5_12, © Springer Science+Business Media, LLC 2011

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processes and easy to use off the shelf semi-automated mutation detection software packages are available facilitating interpretation of large amounts of sequencing data. The vast majority of sequencing is based on Sanger chain termination chemistry usually performed within a cycle sequencing reaction using a PCR product as the template (1). During this reaction cycles of template denaturation, primer annealing, and extension are performed resulting in the incorporation of fluorescently labelled dideoxynucleotides (ddNTPs) into the sequencing products which can then be resolved by semi-automated capillary electrophoresis. Current Sanger chain termination chemistry can readily achieve read lengths of 1,000 bp and per base raw accuracies of more than 99% (2). Fluorescent cycle sequencing of PCR products is a multistage process: the PCR template is prepared and purified to remove unused PCR primers, unincorporated nucleotides (and PCR buffer if possible); the cycle sequencing reaction is then performed; unincorporated dye-labelled fluorescent dideoxyterminators are removed and the sequencing products are then ready to be resolved by capillary electrophoresis using a genetic analyzer. Following data collection sequencing analysis software interprets the fluorescent data to display it in the form of a sequencing chromatogram. Different methodologies are available to perform each of the stages of fluorescent cycle sequencing. This chapter will describe some of the more commonly used approaches and those most familiar to the authors: enzymatic removal of excess primers and nucleotides; BigDye® terminator cycle sequencing and removal of unincorporated dye terminators by the more traditional ethanol precipitation procedure as well as a more recent strategy using Agencourt® CleanSEQ® magnetic bead technology (this method is more amenable to automation facilitating high-throughput workflows). Automated DNA sequencing is one of the most common and robust techniques performed in molecular biology laboratories. Unfortunately, it may not always produce desirable results and problems may be difficult to pinpoint. Fortunately, most failed (or suboptimal) DNA sequencing results have only a fairly limited number of causes. The causes of the most common problems will also be discussed.

2. Materials 2.1. Purification of PCR Products Prior to Cycle Sequencing

1. PCR products (see Note 1). 2. ExoSAP-IT enzymes (GE Healthcare).

Automated DNA Sequencing

2.2. Cycle Sequencing Using BigDye ® Terminator Chemistry

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1. Purified PCR products. 2. BigDye® Terminator Cycle Sequencing Kit (v1.1 or v3.1, Applied Biosystems). 3. BigDye® 5× Terminator v1.1, v3.1 Sequencing Buffer (Applied Biosystems, see Note 2). 4. Forward and Reverse Sequencing Primers diluted to 3.2 mM (see Note 3). 5. Reagent grade water. 6. Sequencing plates (suitable for 3730 genetic analyzer, e.g. MicroAmp™ Optical 96-well Reaction Plates, Applied Biosystems).

2.3. Purification of Cycle Sequencing Products Using Agencourt ® CleanSEQ ® Kit

1. Agencourt CleanSEQ® dye-terminator removal kit (supplied by Beckman Coulter). Store at 4°C and do not freeze. It is important to mix the CleanSEQ® solution thoroughly before using to fully resuspend the magnetic beads (the solution should appear homogeneous). 2. 85% Ethanol (can be prepared in advance and stored at 4°C for 1 week). 3. Agencourt SPRIPlate® Magnetic Plate (96- or 384-well format). 4. Elution buffer (either reagent grade water or 0.1 mM EDTA, pH 8.0). See Note 4. 5. Agencourt® Direct Inject Magnet (96- or 384-well format, optional).

2.4. Purification of Cycle Sequencing Products by Ethanol Precipitation

1. 125 mM EDTA (diluted from a 500 mM stock solution, molecular biology grade from commercial supplier). 2. 100% Ethanol (see Note 5). 3. 70% Ethanol. 4. Hi-Di™ formamide (Applied Biosystems, see Note 6). Formamide is toxic; refer to the material data sheet.

2.5. Capillary Electrophoresis and Analysis Using Applied Biosystems 3730 Genetic Analyzer

1. 50-cm Capillary array (Applied Biosystems, 36 cm length capillary can also be used). 2. POP-7™ Polymer (Performance Optimized Polymer, Applied Biosystems). 3. Sequencing Buffer, 10× with EDTA (Applied Biosystems). 4. 3730 Data collection software (v3.0, supplied by Applied Biosystems with the genetic analyzer). 5. Sequencing analysis software (v5.2, supplied by Applied Biosystems with the genetic analyzer).

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3. Methods The following procedures may be performed either in tubes (0.2 or 0.5 ml) or in plates (96- or 384-well) depending on the sample throughput and level of automation required. The use of 96- and 384-well plates facilitates the use of robotic liquid handling equipment enhancing throughput, reproducibility, and quality of the sequencing process. 3.1. Purification of PCR Products Prior to Cycle Sequencing

To obtain high quality sequencing results it is important to remove excess dNTPs and primers from PCR templates prior to setting up the cycle sequencing reaction. Various methods are available to do this. One common approach is the deployment of the two enzymes Exonuclease I and Shrimp Alkaline Phosphatase (see Note 7 for an alternative technique). The ExoSAP protocol is a very simple way to clean up PCR products before sequencing. Exonuclease I removes excess primers, while the Shrimp Alkaline Phosphatase removes unincorporated dNTPs. 1. Add 2 ml ExoSAP-IT enzyme mix to 5 ml PCR product. It is important to keep the enzyme mix cool during this stage. 2. Mix and place on a thermal cycler at 37°C for at least 15 min, followed by 15 min at 80°C. Consumable costs may be reduced in two ways: first by diluting the ExoSAP-IT enzyme mix one in two with water prior to use and second by adding 1 ml of diluted mix to 5 ml of PCR product. 3. PCR samples can be stored at 4°C for up to 24 h (longer at −20°C).

3.2. Cycle Sequencing Using BigDye ® Terminator Chemistry

Applied Biosystems produce two versions of the BigDye® terminator cycle sequencing kits, v1.1 and v3.1 (see Note 8). Both provide a ready to use sequencing reaction premix to which the user must add a template and appropriate sequencing primer (see Note 3). Applied Biosystems recommend the preparation of either 10 or 20 ml final volume sequencing reactions containing 4 and 8 ml of the terminator mix, respectively. As a guide the following volumes are used to prepare 10 and 20 ml sequencing reactions: Reagent

Quantity for 10 ml Quantity for 20 ml

Cycle sequencing terminator mix

4 ml

8 ml

PCR template

Variable

Variable

Primer (either forward or reverse) 3.2 pmol

3.2 pmol

Reagent grade water

up to 20 ml

up to 10 ml


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It is possible to reduce the volume of the cycle sequencing terminator mix by diluting it with an appropriate volume of 5× BigDye® terminator sequencing buffer thereby reducing consumable costs (see Note 2). Many laboratories routinely use a one in 16 dilution (0.25 ml) of the terminator mix (with 1.875 ml 5× sequencing buffer in a final volume of 10 ml) without deterioration of sequence quality. Optimisation of buffer conditions for specific PCR fragments, however, is recommended. 1. Prepare a mastermix containing appropriate volumes of BigDye® terminator ready mix, primer, 5× BigDye® terminator sequencing buffer (if required) and water. It is important to mix well and centrifuge the sequencing mastermix at this stage (insufficient mixing may lead to the appearance of “ski-slope” sequencing data). If sequencing bidirectionally two mastermixes should be prepared, one containing the forward primer and the other containing the reverse primer. 2. Aliquot each mastermix to appropriate PCR templates (keep the mix cool during this stage). 3. Centrifuge the samples and place on a thermal cycler and cycle using the following conditions: 95°C for 1 min 30 Cycles of: 95°C for 30 s 50°C for 10 s (annealing temperature may be adjusted as appropriate) 60°C for 4 min Hold the reaction at 4°C until required. 4. It is recommended that the sequencing products are processed as quickly as possible to avoid excessive deterioration of sequencing quality; otherwise, store at −20°C. 3.3. Purification of Cycle Sequencing Products Using Agencourt ® CleanSEQ ® Kit

Unincorporated dye terminators must be completely removed before the samples can be analysed by electrophoresis. Excess dye terminators in sequencing reactions obscure data in the early part of the sequence and will therefore interfere with basecalling. The Agencourt® CleanSEQ® system utilizes SPRI® (Solid Phase Reversible Immobilisation) magnetic bead technology. It does not require centrifugation or filtration and can be performed manually or fully automated for high-throughput workflows. This technology tolerates a wider variability of PCR and sequencing reaction product quality than many other techniques including ethanol precipitation. The Agencourt CleanSEQ® procedure is performed in three stages as follows: (a) Selective binding of sequencing extension products to paramagnetic beads and separation of

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the beads with a magnetic field. (b) Washing of the beads to remove unincorporated dyes, nucleotides, salts, and other contaminants. (c) Elution of the purified sequencing product from the paramagnetic beads. If the sequencing reactions have not been performed in a plate they should be transferred to a plate (96- or 384-well formats) at this stage prior to starting the CleanSEQ® procedure. The clean-up can be carried out manually; however, the use of robotic liquid handling equipment is recommended. The suppliers of the CleanSEQ® beads recommend using a 1:1 ratio of sequencing reaction volume to beads. Post-sequencing clean-up costs are therefore significantly reduced when using reduced sequencing reaction volumes (5 and 10 ml). 1. For each 10 ml sequence reaction add 10 ml CleanSEQ® beads and 42 ml 85% ethanol (the volume depends on the initial sequencing reaction volume, see Note 9). 2. Mix by pipetting several times until the mixture is homogeneous. 3. Transfer the plate to a SPRIPlate® magnetic Plate for a minimum of 3 min. During this time the magnetic beads will form a ring around the side of the well and the supernatant will go clear. 4. Keep the sequencing plate on the magnet; remove and discard as much of the supernatant from each well as possible (it contains excess fluorescent dye and contaminants). Ensure that the pipette tip is placed at the bottom of the well when aspirating to avoid disturbing the ring of magnetic beads. 5. Add 100 ml of 85% ethanol to the wells and again leave the plate for a minimum of 30 s to allow the magnetic beads to settle. This must be performed while the plate is situated on the magnet, it is not necessary to mix or resuspend the beads during this step. 6. Repeat steps 4 and 5 for a total of two 85% ethanol washes. 7. Remove all of the supernatant from the wells and then leave the plate to air dry for a minimum of 1 min (on or off the magnet). Excessive drying can lead to degradation of the fluorescent dyes. 8. If using an Agencourt® direct inject magnet plate resuspend the cleaned sequencing products in 70 ml elution buffer and then briefly centrifuge the plate to remove any bubbles that may be present at the bottom of the wells (as they may interfere with electrokinetic injection). If not using an Agencourt® direct inject magnet plate then add 40 ml of elution buffer and allow the sample plate to separate on the magnet for 3–5 min or until the solution is clear. Transfer 35 ml of the clear sample into a new plate for loading on to a genetic analyzer.


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It is important to leave 5–10 ml of liquid behind to prevent transfer of beads in to the final plate. Residual beads can interfere with the injection, causing late starts or failed injections (if this occurs retransfer the samples away from the beads and reinject). 9. The products are now ready to be run on a genetic analyzer. If the samples are not to be loaded immediately seal the plate and store at 4°C for up to 24 h. If not loading within 24 h the products can be stored at −20°C (although freezing may compromise the sequencing quality). It is important to avoid long delays between sample preparation and loading on to the genetic analyzer (see Note 10). 3.4. Purification of Cycle Sequencing Products by Ethanol Precipitation

For most applications, the ethanol/EDTA precipitation method will give clean sequencing data. It is more labour intensive than the CleanSEQ® approach (but significantly cheaper) and therefore is appropriate for processing small sample numbers. The procedure below is appropriate for use with both the v1.1 and v3.1 BigDye® Terminator sequencing kits. As this method involves the use of EDTA it is important to remove any traces of alcohol to ensure the complete removal of salt residues and unincorporated dyes. Salt contamination will lead to drop-off in peak height. If this is a problem it is recommended to perform two 70% washes (rather than one as described below). 1. To each well add 2.5 ml of 125 mM EDTA and then 30 ml 100% ethanol. 2. Seal the plate (use an aluminium foil seal for best results; press the foil on to the wells to prevent leakage). 3. Invert the plate four to five times. 4. Leave at room temperature for at least 15 min to precipitate the extension products. 5. Place the plate in a centrifuge (with a plate adaptor) and spin at 2,000–3,000 × g for 30 min. It is important to proceed to the next step immediately. If this is not possible then spin the plate for an additional 2 min immediately before proceeding to the next step. 6. Carefully remove the foil seal (without disturbing the precipitates), invert the plate and place it on to a paper towel (folded to the size of the plate) sitting within the centrifuge plate adaptor. It is essential to keep the plate inverted during this step to prevent dislodging the pellet. 7. Pulse spin the plate up to 185 × g to ensure the complete removal of the supernatant. It is important to remove the supernatant completely as it contains dissolved unincorporated dye terminators.

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8. Add 30 ml of 70% ethanol to each pellet. 9. Place the plate in a centrifuge and spin at 1,650 × g for 15 min. It is important to proceed to the next step immediately. If this is not possible then spin the plate for an additional 2 min immediately before going to the next step. 10. Invert the plate and place it on to a paper towel (folded to the size of the plate) sitting within a centrifuge plate adaptor. It is essential to keep the plate inverted throughout this step to prevent dislodging the pellet. 11. Spin the plate at 185 × g for 1 min; start the timer when the rotor starts to move. 12. Air dry for 10–15 min before resuspending in 10 ml Hi-Di™ formamide buffer. Briefly centrifuge the plate to remove any bubbles that may be present at the bottom of the wells. The samples are now ready to run on a genetic analyzer. Once resuspended, samples in solution are more subject to degradation at room temperature, a process accelerated by exposure to heat, humidity, and light. The plates should therefore be sealed and run as soon as possible. Alternatively, the dry pellets can be stored in the dark at −20°C until required. 3.5. Capillary Electrophoresis and Analysis Using 3730 Genetic Analyzer

The purified sequencing products may now be loaded on to an Applied Biosystems 3730 genetic analyzer following instructions supplied by the manufacturer. During the sequencing run the 50 cm (or 36 cm) capillary array is filled with POP-7™ and the samples are then injected into the capillary array (a 10 s injection at 1.5 kV is suitable for the majority of samples). Samples are run at 60°C for an appropriate length of time (dependent on fragment length, the run can be skipped once all sequencing data has been collected). Each run requires an instrument protocol appropriate for the combination of capillary length, POP-7™, and BigDye® chemistry used, e.g. the Z-Dye set should be used for BigDye® v3.1 and the E-Dye set should be used for BigDye® v1.1. A run module is also required which defines the injection and running conditions. Analysis of sequencing data requires an analysis protocol. The default analysis protocol set up for the combination of 3730, BigDye® v3.1 (or BigDye® v1.1), and POP-7™ polymer using the KB basecaller works well for the majority of samples. Following electrophoresis, sequencing results are interpreted either by manual inspection of chromatograms or by using a more automated third party software such as Mutation Surveyor (Softgenetics) (see Chapter 10). Identifying the cause of a poor DNA sequencing result can often be very difficult as a particular sequencing problem may have many different underlying causes, or be the result of multiple


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interacting factors. Often the only way to work out the real cause of a particular problem is to perform a process of elimination. This process can be greatly simplified by visually examining both the raw and processed data chromatograms. It may be useful to include a control DNA template in a set of sequencing reactions to help determine whether failed reactions are the result of poor template quality or sequencing reaction failure. Applied Biosystems include a control template and primer in both the v1.1 and v3.1 BigDye® terminator cycle sequencing kits. Notes 11–16 describe the major causes of sequencing problems as well as how to identify them and solve the underlying problems.

4. Notes 1. The quality and quantity of DNA templates used for sequencing reactions determine the quality of sequence data produced. It is important to avoid residual salts, proteins, detergents, and RNA as they interfere with capillary electrophoresis and electrokinetic injection. Capillary electrophoresis is especially susceptible to the presence of residual salts, from template preparation (genomic DNA and PCR products), from cycle sequencing reactions, or from precipitation methods using salts. The negative ions in salts can be preferentially injected into the capillary array during electrokinetic injection, leading to lower signal. In addition, the negative ions compete and interfere with the injection of larger DNA extension fragments, leading to shortened read lengths. The quantity of PCR product used for cycle sequencing has a significant impact on sequencing data. Too little template reduces the signal strength and therefore the peak height of reaction products potentially resulting in bases that cannot be called. Excess template causes several problems as follows: it behaves similarly to proteins and accumulates in the capillary array adversely affecting data resolution and capillary array lifetime, and it results in the generation of short extension fragments which will be injected more efficiently resulting in “top heavy” peak characteristics and shortened reads. The following quantities of PCR product are recommended by Applied Biosystems for use in BigDye® sequencing reactions: 1–3 ng of 100–200 bp products; 3–10 ng of 200– 500 bp products; 5–20 ng of 500–1,000 bp products; and 10–40 ng of 1,000–2,000 bp products. However, it is generally impractical to accurately quantify every PCR fragment prior to sequencing (especially when sequencing large numbers of fragments of varying sizes simultaneously). As a rule

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of thumb, therefore, the authors follow a generic PCR/ sequencing protocol which works well for the majority of fragments ranging from 150 to 900 bp in size. The sequencing template is generated from 50 ng genomic DNA using 30 cycles of PCR. ExoSAP products are then diluted one in three with reagent grade water prior to sequencing set-up. 2. When using the BigDye® 5× sequencing buffer it is important to ensure that the final reaction volume of buffer is at a 1× concentration (the buffer concentration of the neat terminator mix is at 2.5×). Below are examples of the volumes of 5× sequencing buffer required when diluting the ready reaction mix by either one in two or one in four (into a final volume of 10 ml): Reagent

Quantity for one in two

Quantity for one in four

Terminator mix

2 ml

1 ml

5× sequencing buffer

1 ml

1.5 ml

PCR template

Variable

Variable

Primer (either forward or reverse)

3.2 pmol

3.2 pmol

Reagent grade water

Up to 10 ml

Up to 10 ml

3. The sequencing primer also helps to determine the quality of the sequencing data and therefore it is important to consider the melting temperature (ideally between 55 and 60°C) and GC content (ideally about 50%). The use of universal sequencing primers (complementary to 5¢ tails incorporated into PCR templates) facilitates the implementation of high-throughput workflows. The universal sequence is added to the 5¢ end of forward and reverse PCR primers as appropriate. The authors use the following pair of M13 universal sequencing primers (annealing temperature of 50°C): Forward primer TGTAAAACGACGGCCAGT Reverse primer CAGGAAACAGCTATGACC 4. The suggested elution buffers are 0.1 mM EDTA (pH 8.0) or reagent grade water. Water is used to give maximum signal strength while EDTA is used to lower the signal in cases where the signal is too strong. The appropriate elution buffer will vary depending on the sensitivity of the genetic analyzer, the amount of BigDye® used and the type of template. If using a direct inject magnet better results may be obtained using 0.05 mM EDTA in place of the 0.1 mM EDTA. 5. If absolute (100%) ethanol is used in the ethanol/EDTA precipitation procedures, it is important to remember that when exposed to air, 100% ethanol absorbs moisture and


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becomes more dilute over time, resulting in slight variations in concentration. 6. Deionised formamide is used to denature the DNA samples before placing them on to a genetic analyzer (e.g. Applied Biosystems 3730/3730xl DNA analyzers). Applied Biosystems supply and recommend the use of Hi-Di™ Formamide buffer for use on their instruments. Formamide purchased from other commercial suppliers may be supplied in glass bottles, which can produce contamination from minerals. In addition, formamide purchased from commercial suppliers is often contaminated with variable amounts of water and organic and inorganic ions. Water reacts slowly with formamide to produce formic acid and ammonia. The ionic products of this reaction cause two problems as follows: they compete significantly with the larger DNA ions for injection into the capillary array resulting in weaker signals, and they react with the DNA causing degradation of the sample. The author recommends storing™ formamide buffer at −20°C as 1 ml aliquots (sufficient for preparing one 96-well plate) to avoid multiple freeze–thaw cycles. 7. The Agencourt® AMPure® system is an alternative approach for the purification of PCR products prior to cycle sequencing based on similar technology to the CleanSEQ® system. Again it can be easily automated for either 96- or 384-well formats. The Agencourt® AMPure® system removes unincorporated primers, dNTPs, DNA polymerases, and salts used during PCR amplification that can interfere with downstream sequencing. According to manufacturers guidelines this product will routinely produce recovery rates greater than 90% for amplicons larger than 200 bp and greater than 85% for smaller PCR amplicons. 8. Two versions of the BigDye® terminator cycle sequencing kits are available. The v3.1 is recommended for mixed-base detection and long read protocols whereas the v1.1 kit is recommended for sequencing short PCR products using rapid electrophoresis run modules or applications where the sequencing read begins close to the primer. 9. There are two CleanSEQ® calculators available dependent on the sequencing reaction volume used: For 10 ml sequencing volumes: volume of 85% ethanol = 2.077 × (10 ml + sample volume). For 5 ml sequencing volumes: volume of 85% ethanol = 1.428 × (5 ml + sample volume). 10. It is important to avoid long delays between sample preparation and electrophoresis. Excessive delay may cause a breakdown of the DNA sequencing chemistry resulting in a rapid

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drop off in signal intensity of the raw data. This may therefore become a problem when loading large numbers of plates simultaneously, especially if plates are sitting in the analyzer in excess of 12 h. Causes of sequencing chemistry breakdown include low pH (particularly a problem for samples resuspended in pure water when exposed to air) and nuclease contamination. If sequencing chemistry breakdown is a problem samples should be resuspended in 50 mM EDTA, pH 7.4, just before injection and if possible avoid loading more than four to six sequencing plates on to the genetic analyzer at a time. 11. A failed DNA sequencing reaction results in a noisy or “messy” trace chromatogram composed of low quality score sequence peaks. The signal strength in the raw channel will usually be below 100. There are a number of causes of failed DNA sequencing reactions including: poor quality DNA; loss of the reaction during clean-up (a particular problem when using ethanol precipitation clean-up protocols); too much template DNA; use of incorrect or degraded primer; low quality water; “dead” sequencing chemistry which can occur if the BigDye® chemistry is stored under the wrong conditions or is freeze–thawed too many times (resulting in degradation of either the Taq DNA polymerase or dye-labelled nucleotides); or finally a blocked capillary (picked up when every trace using a particular capillary fails). 12. Sequencing traces composed of two or more peaks at each position (with good trace signal strength in the raw channel) will usually be the result of mixed DNA templates. Lower levels of mixed signal (below 20%) are normally base called sufficiently well by the KB basecaller; however, higher levels will result in a significant reduction in the quality scores such that the base-called sequence may not match the expected sequence. Causes of mixed template sequencing include: the presence of more than one template in the reaction (the most common cause of mixed template sequencing); the presence of two primers in the sequencing reactions (for example, forward and reverse sequencing primers); leftover primers in the PCR template; or using an annealing temperature that is too low in the sequencing reaction. 13. Short DNA trace read length problems occur when a sequencing reaction cannot routinely produce more than 650 high quality bases for samples run on 36 cm arrays or more than 750 bases for traces collected on 50 cm arrays under standard run conditions. The ends of these shorter than expected traces appear “messy” with “rough” looking peaks. Causes of short DNA sequencing read lengths include the following: too much or too little template DNA (DNA concentration can be evaluated by gel electrophoresis); excessive dilution of the


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BigDye® reagent (better results may be obtained by reducing the reaction volume rather than using high dilution factors of the BigDye® chemistry); too much primer; or the use of “dirty” template DNA. 14. Another common problem is premature loss of resolution of sequence traces manifesting as “blurry” trace chromatogram peaks occurring before base 500 for traces collected using 36 cm arrays or before base 650 for traces collected on 50 cm arrays. When this occurs nucleotide runs in the sequence appear as indistinct blobs rather than as individually separate peaks. Causes of blurry sequencing traces include capillary overload (normally caused by running “dirty” samples with large amounts of template contaminated with non-template genomic DNA, proteins, or salt) and high sequencing run voltages (the higher the run voltage the lower the resolution of the peaks in the trace file). 15. Sequencing traces normally start between scan number 1,800 and 2,200 for 50 cm capillaries and between scan 1,500 and 1,900 for 36 cm capillaries. Signals that appear later than this (after scan 3,200 for 50 cm traces and scan 2,700 for 36 cm traces) will often be problematic because the processed data is often of low resolution with wide peaks. Signal start delays are caused by capillary overload as a result of either too much template DNA or dirty template DNA (contaminated with proteins and/or salt as well as CleanSeq® magnetic beads). 16. Late “G” dye peaks at base positions 190 and 400 may appear as a result of the breakdown of the BigDye® sequencing chemistry before sample clean-up (i.e. before the leftover labelled nucleotides are removed). The earlier of the two “G” peaks is normally the largest; the 400 base G peak may not be visible. The major causes of breakdown of the BigDye® che mistry is excess heating or excessive freeze/thaw cycles. The problem of late “G” peaks is often only detected if the sample clean-up is not performed correctly (early dye “blobs” may also be present). If late “G” dye peaks are problematic avoid overheating the DNA sequencing reactions (keep the denaturation temperature in the sequencing thermocycling step below 96°C and under 20 s) and avoid excessive freeze/thaw cycles of the BigDye® chemistry, e.g. by aliquoting the BigDye® stock into smaller volumes. References 1. Sanger, F., Nicklen, S., and Coulson, A. R. (1977) DNA sequencing with chain terminating inhibitors. Proc. Natl Acad. Sci. USA 74, 5463–5467.

2. Shendure, J., and Ji, H. (2008) Nextgeneration DNA sequencing. Nat. Biotechnol. 26, 1135–1145.

Chapter 13 Phylogenetic Microarrays for Cultivation-Independent Identification and Metabolic Characterization of Microorganisms in Complex Samples Alexander Loy, Michael Pester, and Doris Steger Abstract High-throughput sequencing and hybridization technologies promise new insights into the natural diversity and dynamics of microorganisms. Among these new technologies are phylogenetic oligonucleotide microarrays (phylochips) that depend on the standard molecules for taxonomic and environmental studies of microorganisms: the ribosomal RNAs and their encoding genes. The beauty of phylochip hybridization is that a sample can be analyzed with hundreds to thousands of rRNA (gene)-targeted probes simultaneously, lending itself to the efficient diagnosis of many target organisms in many samples. An emerging application of phylochips is the highly parallel analysis of structure–function relationships of microbial community members by employing in vivo substrate-mediated isotope labeling of rRNA (via the isotope array approach). This chapter provides an introduction to phylochip and isotope array analysis and detailed wet-lab protocols for preparation, labeling, and hybridization of target nucleic acids. Key words: Microarray, Ribosomal RNA, Phylochip, Microbial diversity, Microbial diagnostics, Isotope labeling

1. Introduction Microarrays that consist of rRNA-based oligonucleotide probes (so-called phylogenetic microarrays or phylochips) are highperformance hybridization platforms for not only identification of many different phylogenetic groups of microorganisms but also their physiological profiling in parallel, a methodological option that is strongly needed considering the enormous microbial richness in many natural and biotechnological environments. Shortly described, phylochip-based identification (and under certain circumstances also quantification (1)) of target microorganisms begins with DNA or RNA isolation from a sample of interest. Bimal D.M. Theophilus and Ralph Rapley (eds.), PCR Mutation Detection Protocols, Methods in Molecular Biology, vol. 688, DOI 10.1007/978-1-60761-947-5_13, © Springer Science+Business Media, LLC 2011

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Prior to fluorescence labeling, PCR is often used for amplification of rRNA genes to improve detection sensitivity. Another possibility for the analysis of abundant members of a microbial community is PCR-independent, direct fluorescence labeling of environmental rRNA. After hybridization of the fluorescently labeled target nucleic acids with the microarray, probe signals are digitalized with a fluorescence imager. At last, diverse computational approaches can be applied for normalization, statistical analysis, and interpretation of the large amount of obtained data. Two types of phylochips can be distinguished based on the total number of probes. The high capacity of the microarray format is best exploited with high-density phylochips that carry several thousands of oligonucleotide probes (2). In contrast, only up to a few hundred oligonucleotides are immobilized on lowdensity phylochips. These usually custom-made microarrays target microorganisms that are defined by (a) their physiological capabilities (3), (b) their phylogeny or taxonomy (4), or (c) the environment where they live (5). A great benefit of low-density phylochips is that the sensitivity and specificity of almost each individual probe can be empirically determined by using an adequate set of reference organisms. This crucial information on the performance of a microarray is not economical to acquire for high-density phylochips, simply due to the large number of reference targets that would be required to test the different probes. The DNA microarray technique involves a series of different computational and laboratory-based steps, starting with the initial development of a microarray to its final application for the analysis of unknown samples. De novo development of a micro array, including establishment of target sequence databases, target group definition and probe design, and in silico/empirical evaluation of the new microarray (6), is not addressed here. Instead, this chapter presents already established lab protocols for microarray fabrication by spotting and for fluorescence labeling/ hybridization of target nucleic acids using three different methods. Furthermore, an introduction to the isotope array approach (7) provides guidance for phylochip-based structure–function analysis of complex microbial communities.

2. Materials 2.1. Microarray Fabrication

1. Aldehyde group-coated glass slides (e.g., CEL Associates, Houston, Texas). Prior to spotting, dust and other particles should be removed from slides by using a clean air flow. 2. Amino-modified oligonucleotide probes (5¢-NH2-(spacer)(diagnostic rRNA-targeted sequence)-3¢): rRNA-targeted probes are typically 15–25 nucleotide in length.

Phylogenetic Microarrays for Cultivation-Independent Identification

189

The 5¢-terminal nucleotide is aminated to allow for covalent coupling of the oligonucleotides to aldehyde groupcoated slides. Probes should additionally contain a 5¢-spacer element, e.g., a stretch of 30 dTTPs, to improve accessibility during hybridization (8). We use a probe concentration of 50–100 pmol ml−1 in 50% dimethylsulfoxide as spotting buffer (see Note 1). Probes are aliquoted in 384-polystyrol micro well plates (Ritter GmbH, Schwabmünchen, Germany). 3. BioRobotics MicroGrid spotter (Genomics Solutions, Ann Arbor, MI, USA). 4. Standard quill or split pins with approximately 100 mm tip diameter: Microspot pins 2500 (Genomics Solutions) or SMP3 pins (TeleChem International Inc., Sunnyvale, CA, USA). Larger pins, e.g., SMP15XB pin with tip diameters of 600 mm (TeleChem International Inc.), are needed for isotope array analysis. 5. 0.2% (w/v) sodium dodecyl sulfate (SDS). 6. 100% ethanol. 7. Phosphate buffer: 80:20 (v/v) mixture of 200 mM Na2HPO4 and 200 mM NaH2PO4, pH 7.2–7.4. 8. Phosphate-buffered saline (PBS): 130 mM NaCl, 5% phosphate buffer (v/v), pH 7.2–7.4. 9. Sodium borohydride (NaBH4) solution: Dissolve 1.0 g NaBH4 in 300 ml PBS. Add 100 ml of 100% ethanol to reduce bubbling. Prepare freshly prior to use. 2.2. PCR-Based Microarray Analysis: Hybridization of Labeled DNA 2.2.1. Polymerase Chain Reaction

1. Target group-specific 16 S rRNA gene-targeted primers (50 pmol ml−1). See Table 1 for a list of general bacterial and archaeal primers. 2. PCR reagents (e.g., Fermentas, http://www.fermentas.com): 10× Taq polymerase buffer, dNTP-Mix (dATP, dTTP, dGTP, dCTP: 2.5 mM each), MgCl2 (25 mM), and Taq-Polymerase (5 U ml−1). 3. PCR purification kit (e.g., QIAquick PCR Purification Kit, Qiagen).

2.2.2. Random Prime Labeling of Target DNA

1. DecaLabel DNA labeling kit (Fermentas), including Klenow Fragment, (without exonuclease activity, 5 U ml−1), decanucleotide in 5× reaction buffer, Mix C (dGTP, dATP, dTTP; each 0.33 mM), NTP Mix (0.25 mM of each nucleotide). 2. 1 mM dCTP. 3. 1 mM Cy3- or Cy5-labeled dCTP (GE Healthcare Life Sciences, USA), store at −20°C in the dark. 4. QIAquick Nucleotide Removal Kit (Qiagen).

S-D-Bact-0008-b-S-20

S-D-Bact-0008-c-S-20

S-D-Bact-0008-d-S-20

All 4 variants together

27f-YM-Bif

27f-YM-Bor

27f-YM-Chl

27f-YM+3

Most Bacteria

Most Bacteria

Target organisms

AGA ATT TGA TCT TGG TTC AG

Most Bacteria, not SR1, OD1

Most Bacteria and Archaea, not Fibrobacteres, Nanoarchaeum

Chlamydiales

AGA GTT TGA TCC TGG CTT AG Borrelia

AGG GTT CGA TTC TGG CTC AG Bifidobacteriaceae

AGA GTT TGA TYM TGG CTC AG Most Bacteria

ACG GGC GGT GTG TAC AAG

S-D-Bact-0008-a-S-20

27f-YM

CCT ACG GGI GGC IGC A

UNIV1389aR S-D-Univ-1389-a-A-18


I-341F

AGA GTT TGA TYM TGG CTC

GGY TAC CTT GTT ACG ACT T


BACT8F (616V)

Sequence (5¢-3¢)c

PROKA1492R S-*-Proka-1492-a-A-19 (1492R)

Full nameb

Short namea

186553

52702

135483

+ 114

+ 99

+ 39

135231

497706

136624

74.4%

78.8%

77.6%

94.9%

78.3%

Specificityd

Table 1 Selected PCR primers for amplification of almost complete small-subunit rRNA gene fragments from Bacteria and Archaea

(3)

(3)

(42)

(41)

(40)

Selected referencesc

190 Loy, Pester, and Steger

NTe

Most Archaea, not Korarchaeota and Nanoarchaeota

Most Archaea

Most Archaea, not Korarchaeota and Nanoarchaeota

4111

8365

12778

Most Archaea, not Nanoarchaeota 7197

Most Bacteria

86.6%

81.9%

61.3%

71.3%

(3)

(45)

(44)

(43)

(40)

b

a

Synonymous short names used in the literature are indicated in parentheses Name of SSU rRNA gene-targeted oligonucleotide primer based on the nomenclature of Alm et al. (46) c Please note that for some primers variants exist in the literature (differing in length and/or type and number of degenerate nucleotides) d Specificity was tested using the Ribosomal Database Project (RDP) probe match tool (Release 10 Update 4; 690,149 16 S rRNA sequences). Absolute and relative numbers of perfect-match hits are shown (47) e NT not tested. Specificity check was not possible because the probe binding site was outside the sequence range that is testable by the RDP probe match tool

ACG GGC GGT GTG TGC AAG

ACK GCT CAG TAA CAC GT

UNIV1389cR S-D-Univ-1389-c-A-18

S-D-Arch-0109-a-S-17

ARCH109F (A109F)

TCY GKT TGA TCC YGS CRG

TTM GGG GCA TRC IKA CCT

S-D-Arch-0008-a-S-18

ARCH8F (A3Fb)

CAK AAA GGA GGT GAT CC

ARCH1196R S-D-Arch-1196-a-A-18 (UA1204R)

S-D-Bact-1529-a-A-17

BACT1529R (630R)

Phylogenetic Microarrays for Cultivation-Independent Identification 191

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2.2.3. Microarray Hybridization

1. 10% (w/v) blocking reagent solution (Roche, Mannheim, Germany), store at −20°C. 2. 20× sodium citrate/sodium chloride (SSC; 3.0 M sodium chloride, 0.3 M sodium citrate). 3. 10% (w/v) SDS. 4. 10% (w/v) N-laurylsarcosine. 5. Formamide (high-grade, Carl Roth GmbH & Co., Karlsruhe, Germany), store at 4°C, handle under a fume hood (irritating, toxic). 6. 500 mM EDTA, pH 7.5. 7. 1 M Tris–HCl, pH 7.5. 8. Double-distilled water. 9. HybriWell™ Sealing System (e.g., HBW2222; 21.6 × 21.6 mm, loading volume: ca. 60 ml; Grace Bio-Labs, Bend, OR, USA). 10. ThermoTWISTER in combination with the PALMhybIV thermoblock (QUANTIFOIL Instruments GmbH, Jena, Germany).

2.2.4. Microarray Washing

1. 5 M tetramethyl ammonium chloride (TMAC), handle with care (toxic). 2. 10% (w/v) SDS. 3. 500 mM EDTA, pH 7.5. 4. 1 M Tris–HCl, pH 7.5.

2.3. PCR-Based Microarray Analysis: Hybridization of Labeled RNA 2.3.1. In Vitro Transcription

1. Template DNA for in vitro transcription is prepared by PCR with 16 S rRNA gene-targeted primers (Table 1). The forward primer must contain the promoter recognition site for T3-RNA polymerase (5¢-AAT TAA CCC TCA CTA AAG GG-3¢) at the 5¢end (see Note 2). 2. T3 DNA-dependent RNA polymerase (20 U ml−1), 5× T3 RNA polymerase transcription buffer (Fermentas). 3. Ribonucleotides: ATP, CTP, GTP, UTP (100 mM each, Roche). 4. RNase Inhibitor: RNasin, 40 U ml−1 (Fermentas). 5. 5 mM Cy3- or Cy5-labeled UTP (GE Healthcare Life Sciences), store at −20°C and keep in the dark. 6. 100 mM dithiothreitol (DTT). 7. RNase-free molecular-grade water, e.g., double-distilled water treated with diethyl pyrocarbonate (DEPC).

2.3.2. DNA Digestion

1. RNase-free DNase I (1 U ml−1), 10× DNase I reaction buffer containing 25 mM MgCl2, and 25 mM EDTA (Fermentas).

Phylogenetic Microarrays for Cultivation-Independent Identification 2.3.3. Cleanup of RNA

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1. RNase-free TE buffer (10 mM Tris–HCl and 1 mM EDTA prepared in DEPC-treated double-distilled water), pH 8.0. 2. 5 M DEPC-treated NaCl (NaClDEPC). 3. 100% ethanol, per analysis grade. 4. 70% ethanol (dilute 100% ethanol with DEPC-treated water). 5. RNase inhibitor: RNasin, 40 U ml−1 (Fermentas).

2.3.4. Fragmentation of Labeled RNA

1. 25 mM Tris–HCl pH 7.4, prepared in DEPC-treated doubledistilled water. 2. 100 mM DEPC-treated ZnSO4 (ZnSO4 DEPC). 3. 500 mM DEPC-treated EDTA (EDTADEPC).


1. RNase-free molecular-grade water. 2. 100× Denhardt’s solution, store at −20°C. 3. 20× SSCDEPC pH 7.0 (3.0 M sodium chloride, 0.3 M sodium citrate, DEPC-treated). 4. Formamide (high-grade), store at 4°C, handle under a fume hood (irritating, toxic). 5. 10% (w/v) DEPC-treated SDS. 6. HybriWell™ Sealing System (Grace Bio-Labs, Bend, OR, USA). 7. ThermoTWISTER (QUANTIFOIL Instruments GmbH, Jena, Germany).


1. 2× SSC/0.1% SDSDEPC: Prepare from 20× SSCDEPC and 10% SDS stock solution. 2. 0.1× SSC: Prepare from 20× SSCDEPC stock solution. 3. DEPC-treated double-distilled water.

2.4. PCR-Independent Microarray Analysis: Hybridization of Labeled, Native RNA

1. CyScribe Direct mRNA Labeling Kit (GE Healthcare Life Sciences). 2. 5 M NaClDEPC. 3. 100% ethanol, per analysis grade. 4. Dry ice. 5. 70% ethanol (dilute 100% ethanol p.a. with DEPC-treated double-distilled water). 6. Reagents for RNA fragmentation, hybridization, and microarray washing as outlined in Subheading 2.3.

2.5. The Isotope Array Approach

1. Radioactively labeled substrates (e.g., from GE Healthcare). 2. Reagents and media for incubation of environmental samples.

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3. Reagents for RNA labeling, fragmentation, hybridization, and microarray washing as outlined in Subheadings 2.3 and 2.4. 4. b-imager (BioSpace Mesure, Paris, France) and a highly sensitive scintillation membrane (Zinsser Analytic, Frankfurt, Germany).

3. Methods 3.1. Microarray Fabrication

Two main techniques for manufacturing DNA microarrays are available: in situ synthesis of oligonucleotides directly on the microarray surface (9–11) and spotting of presynthesized probes onto a solid microarray substrate. The spotting procedure employs a robotic device for depositing probes onto defined positions (i.e., in a grid-like fashion) on the microarray surface (12). A broad range of different surface chemistries are available for tethering of spotted probes and are described in detail elsewhere (13). We use a contact-spotting procedure for manufacturing low-density phylochips: 1. 5¢-amino-modified oligonucleotide solutions are spotted onto aldehyde-coated planar glass slides using a BioRobotics MicroGrid contact-arrayer with split pins (see Note 3). 2. Place freshly spotted microarrays in a humid chamber at room temperature for 24 h (see Note 4). 3. For removal of unbound probes, wash slides thoroughly in 0.2% SDS (i.e., for 2 min with vigorous agitation). Repeat this step. Thereafter, rinse slides thoroughly in double-distilled water (i.e., for 2 min with vigorous agitation). Repeat this step and allow slides to dry completely (approx. 5 min). For reduction of free aldehyde groups, agitate slides for 5 min in sodium borohydride solution. Immerse slides in ice-cold 100% ethanol to stop the reaction. Rinse slides for 1 min in 0.2% SDS. Repeat this step. Rinse slides for 1 min in doubledistilled water. Repeat this step. Finally, immerse slides for 2 s in 100°C hot double-distilled water and dry them by centrifugation (3 min, 290 × g). Microarrays are now ready for hybridization and can be stored in the dark at 25°C (see Note 5). 4. Perform quality control of spotted microarray (see Note 6).

3.2. PCR-Based Microarray Analysis: Hybridization of Labeled DNA

Nucleic acid extraction efficiencies from different samples, cell types, and organisms can vary drastically. Furthermore, nucleic acids isolation protocols for complex samples such as soils or sediments must also ensure proper removal of substances that interfere with subsequent chemical or enzymatic reactions.


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In order to maintain good comparability, the same (set of ) extraction method(s) should be used for all samples in a microarray experiment. Low nucleic acid extraction efficiencies or low abundance of target populations may require target amplification by, e.g., whole-community genome amplification (14, 15) and/or selective PCR assays (16), to generate sufficient target for sensitive microarray hybridization. Fluorescence labeling of PCR products can be achieved in several ways. Amplicons are labeled during the PCR reaction using fluorescently labeled primers (17–19) (see Note 7), Cy-dye labeled dNTPs (20, 21), or amino-allyl dNTPs for subsequent tagging with Cy-dye molecules (5). Alternatively, amplicons are labeled after PCR by random prime labeling (3, 4, 22) (see Note 8). A protocol for the latter case is described in the following section. 3.2.1. Polymerase Chain Reaction

1. Use environmental genomic DNA or DNA from reference organisms as a template. 2. PCR amplification of 16 S rRNA genes is carried out in 50 ml reaction mixtures containing 1× PCR buffer, 2 mM MgCl2, dNTP mix (0.25 mM each), forward and reverse primers (0.5 mM each), and 2.5 U of Taq polymerase (see Note 9). 3. The size and amount of the PCR product is assessed by agarose gel electrophoresis. 4. Purify PCR products by using the PCR purification kit according to the manufacturer’s instructions (use double-distilled water for final elution of DNA). Determine the DNA concentration, e.g., by using a photometer. If necessary, vacuum-dry the purified PCR products to increase concentration (see Note 10).

3.2.2. Random Prime Labeling of Target DNA

1. Purified PCR products are labeled using the DecaLabel DNA labeling kit and Cy3- or Cy5-labeled dCTPs. Reaction mixtures (total volume 45 ml) contain 200 ng of purified PCR product and 10 ml of decanucleotides in reaction buffer and are denatured at 95°C for 10 min and placed immediately on ice. Add 3 ml of deoxynucleotide Mix C (without dCTP), 0.5 ml of Cy-dCTP, 0.5 ml of dCTP, and 1 ml of the Klenow fragment. Incubate the labeling reaction mixture at 37°C for 45 min. Subsequently, denature the reaction mix for 10 min at 94°C and place immediately on ice. Add 1 ml of the Klenow fragment and incubate the labeling reaction mixture again at 37°C for 45 min. Complete labeling by addition of 4 ml of dNTP-mix and incubation at 37°C for 60 min (see Note 11). 2. To remove the enzyme and unincorporated deoxynucleotides and decanucleotides, the labeling mixture is purified with the QIAquick Nucleotide Removal Kit according to the manufacturer’s instructions with the exception that during the final step DNA is eluted with 50 ml double-distilled water.

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3. Determine DNA concentration and labeling density, e.g., by using a photometer (guiding value ~50 pmol Cy dye/mg DNA). Aliquot labeled DNA at defined target concentrations. Finally, vacuum-dry DNA and store in the dark at −20°C. 3.2.3. Microarray Hybridization

The hybridization and washing of microarrays can be performed either by using a fully-automated hybridization station (5) or manually (3, 4). In the following, a manual, nonstatic hybridization protocol is described. 1. The hybridization buffer consists of 500 ml of 20× SSC, 20 ml of 10% N-laurylsarcosine, 4 ml of 10% SDS, 200 ml of 10% (w/v) blocking reagent stock solution, and x ml formamide (see Note 12). Adjust hybridization buffer with double-distilled water to a final volume of 2 ml (see Note 13). 2. Prepare hybridization solution by dissolving the labeled, vacuumdried DNA in the required volume of hybridization buffer. 3. The HybriWell Sealing System in combination with the temperature- and agitation-controlled ThermoTWISTER device is used for hybridization (see Note 14). 4. Carefully apply the adhesive HybriWell chamber over the area of the slide that contains the spotted probes and make sure that an air-tight seal is formed. 5. Place the microarray on the aluminum plate of the ThermoTWISTER and set the hybridization temperature. 6. During prewarming of the slides, the hybridization solution is denatured for 10 min at 95°C. Subsequently, the hybridization solution is pipetted into one of the ports of the HybriWell chamber. The chamber must be free of air bubbles. The amount of hybridization solution depends on the size of the HybriWell chamber. Close the ports with seal pads and set the ThermoTWISTER to the desired agitation speed (e.g., SP3). Incubate overnight for 18 h.


1. All washing steps are performed in 50 ml polypropylene tubes. 2. To prepare the washing buffer, mix 30 ml of 5 M TMAC, 2.5 ml of 1 M Tris–HCl, 200 ml of 500 mM EDTA, 500 ml of 10% SDS, and adjust with double-distilled water to a final volume of 50 ml. 3. After hybridization, remove HybriWell chamber and immediately immerse the microarray in 50 ml of prewarmed TMAC washing buffer. Wash thoroughly for 5 min at the desired temperature (see Note 15). 4. Finally, rinse the microarray twice in ice-cold double-distilled water and dry by centrifugation (3 min, 290 × g).


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5. Proceed directly with scanning or store microarrays dry in the dark (e.g., in a slide box with a small desiccant bag) at ~20°C. 3.3. PCR-Based Microarray Analysis: Hybridization of Labeled RNA

3.3.1. In Vitro Transcription

An alternative approach for labeling PCR-amplified DNA makes use of in vitro transcription into RNA. Linear amplification during in vitro transcription produces large amounts of singlestranded target RNA. Single-stranded targets are preferred over double-stranded targets (e.g., fragments of double-stranded DNA are produced by random prime labeling of target DNA) because they are less prone (at least theoretically) to undesired molecular interactions during hybridization such as reannealing of the two fully complementary target strands (which competes with the formation of probe–target hybrids) or ‘probe hitchhiking’ (i.e., probe-bound targets form heteroduplexes with nontarget sequences via conserved sequence regions, resulting in unspecific probe signals) (23). For labeling, modified ribonucleotides can be directly incorporated into nascent RNA during in vitro transcription (24, 25), or a label can be chemically coupled to RNA after in vitro transcription (26). In the following section, a protocol for in vitro transcriptionbased labeling and hybridization of RNA is described (see Note 16). This protocol can be adapted for the direct analysis of extracted RNA by implementation of a cDNA synthesis step using reverse transcription (27). 1. PCR-amplification of template DNA with 16 S rRNA genetargeted primers (forward primer must contain T3-promoter sequence) and purification of PCR products are performed as outlined in Subheading 3.2. 2. Prepare reaction mixture by combining 4 ml 5× T3 RNA polymerase buffer, 2 ml 100 mM DTT, 1 ml 10 mM ATP, 1 ml 10 mM CTP, 1 ml 10 mM GTP, 0.75 ml 10 mM UTP, 0.5 ml 5 mM Cy3- or Cy5-UTP, 0.5 ml 40 U ml–1 RNasin, and 400 ng of purified PCR product. Adjust reaction volume to 18 ml using DEPC-treated double-distilled water. Add 2 ml 20 U ml–1 T3 RNA polymerase. Mix gently by vortexing and spin briefly. Incubate at 37°C for 5 h. Stop reaction by cooling on ice or proceed to the next step.

3.3.2. DNA Digestion

1. For degradation of template DNA, add 2 ml DNase I buffer and 2 ml RNase-free DNase I and incubate at 37°C for 15 min. Add 2 ml of 25 mM EDTA to stop enzymatic digestion.

3.3.3. RNA Cleanup

1. Adjust the volume to 100 ml with TE buffer. Add 10 ml 5 M NaCl and mix well by inverting the tubes (not by vortexing). Add 300 ml of ice-cold 100% ethanol and mix well

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(not by vortexing). Incubate at –20°C overnight for precipitation of RNA. Centrifuge for 30 min at maximum speed (23,645 × g) at 4°C and wash pellet with 500 ml of ice cold 70% ethanol. Air-dry the pellet to remove residual ethanol (see Note 17). Resuspend the pellet in 50 ml TE buffer and add 0.5 ml RNasin (40 U ml–1). 2. Determine RNA concentration spectrophotometrically. 3. Optional: Run a denaturing gel to check integrity of transcript RNA. 4. Store RNA at −20°C or −80°C (see Note 18). 3.3.4. Zn2+-Based Fragmentation of Labeled RNA

1. Adjust RNA concentration with RNase-free molecular-grade water to 80 ng ml–1. Mix 50 ml of labeled RNA (4 mg in total) with 1.43 ml 1 M Tris–HCl pH 7.4 and 5.71 ml 100 mM ZnSO4. Incubate reaction mix at 60°C for 30 min (see Note 19). Stop reaction by adding 1.43 ml 500 mM EDTA to chelate Zn2+ ions. Add 1 ml of RNasin (40 U ml–1). 2. Determine RNA concentration spectrophotometrically and proceed directly with hybridization or store RNA at −80°C.


1. Except for the hybridization buffer, hybridization of labeled RNA is generally performed as outlined above for labeled DNA. 2. Prepare hybridization buffer by mixing 300 ml 20× SSC, 10 ml 10% SDS, 10 ml 100× Denhardt’s reagent, x ml formamide, and y ml target RNA. Adjust to 1 ml with RNase-free moleculargrade water (see Note 12). 3. Directly before hybridization, the hybridization mix is incubated for 1 min at 65°C.


1. All washing steps are performed in 50-ml polypropylene tubes. 2. Remove HybriWell hybridization chamber and immediately immerse slide in 2× SSC/0.1% SDSDEPC. Wash by shaking at room temperature for 5 min. 3. Transfer slide in 0.1× SSCDEPC and wash slide by shaking at room temperature for 5 min. 4. Rinse slide thoroughly for 1 min in ice-cold DEPC-treated, double-distilled water. Dry slide by centrifugation (3 min, 290 × g). 5. Proceed directly with scanning or store microarrays dry in the dark (e.g., in a slide box with a small desiccant bag) at ~20°C.

3.4. Hybridization with Native RNA

Direct hybridization of environmental RNA extracts is the method of choice due to absence of error-prone amplification steps.


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However, given that the relative rRNA content of low-abundant community members will be below the detection limit of the microarray, microarray hybridization of native RNA, and thus the isotope array approach that relies on this strategy (see next section), is generally restricted to the most abundant microorganisms (>1%) in a sample. 1. Extract total RNA from your sample. See Subheading 3.2 for general recommendations. Evaluate RNA purity and quality by gel electrophoresis. 2. The CyScribe Direct mRNA Labeling Kit is used according to the manufacturer’s instructions for direct chemical labeling of RNA with Cy3 or Cy5 dyes. This kit covalently labels guanine residues at the N7 position and thus does not interfere with subsequent base-pairing. 3. 0.5 mg of RNA is used for one labeling reaction. 4. To remove excess dye, labeled RNA is precipitated with 0.1 volume of 5 M NaCl and 3 volumes of 100% ethanol at −80°C (on a dry ice–ethanol mixture). RNA is pelleted by centrifugation (30 min, 23,645 × g, 4°C), washed with 70% ethanolDEPC, air-dried in the dark and resuspended in 50 ml of RNAse-free molecular-grade water. 5. All subsequent steps are performed as described in Subheading 3.3, starting with the RNA fragmentation step. 3.5. The Isotope Array Approach

The isotope array approach exploits rRNA-targeted microarray technology for tracing the assimilation of a radioactively (14C) labeled substrate by members of complex microbial communities (28). By making use of the multiple probe hybridization format of a microarray, many phylogenetically different microorganisms with a defined metabolic ability can be identified in a single hybridization experiment. Proof-of-principle applications of the isotope array include the analysis of ammonia-oxidizing bacteria (29) and denitrifying bacteria within the order Rhodocyclales (M. Hesselsoe, A. Loy, in preparation) in activated sludge. 1. The isotope array approach starts with the incubation of an environmental sample with a 14C-labeled substrate. The incubation conditions and time (e.g., type of carbon source, electron donor and acceptor, etc.) determine the physiological function of interest and thus the functional microbial guilds that are under investigation. Microorganisms that use the provided labeled substrate as carbon source will incorporate the 14C-label into their biomass (including their rRNA). 2. After incubation, total RNA is extracted and hybridized to a 16 S rRNA-targeted microarray following the procedure described in Subheading 3.4 (see Note 20).

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3. Following the analysis of the fluorescence signal of the individual microarray probes, also their radioactivity signal is recorded with a b-imager by using a highly sensitive scintillation membrane (Zinsser Analytics, Germany) and chemoluminescence detection. The analysis time on the b-imager is usually around 24 h. The radioactivity for each spot and background is quantified in disintegrations per minute (dpm) using b-Vision software (BioSpace Mesures). By taking into account the incubation time of the 14C-substrate with the sample, the ratio between the radioactivity and fluorescence signal of the microarray probe spots provides a quantitative measure of how efficiently and how fast a probe-defined population has incorporated the 14C-label into its rRNA. 3.6. Data Analysis

Diverse image acquisition devices (i.e., microarray scanners) and software tools are available for recording and subsequent computational analysis of fluorescence signals from microarrays. We use the GenePixTM Personal 4100A scanner and the GenePix Pro software for initial image acquisition and analysis (Molecular Devices, Sunnyvale, CA, USA). As exemplified by the protocols outlined above, a diagnostic microarray experiment often involves the analysis of one sample per hybridization (single-color experiment, in contrast to two-color microarray experiments that are typically performed for gene expression analysis). The required scanning resolution is generally dependent on the area of the probe spot because a minimum of 100 pixels per spot are needed for adequate statistics. Digital image analysis begins with the definition of each microarray spot by superimposing a grid of individual circles on the fluorescence image. The mean or median signal intensities are quantified for each probe spot and its local background and used for calculation of a signal-to-noise ratio. The simplest of all further possible analysis strategies considers a probe as positive if its signal-to-noise ratio is above an arbitrarily defined or empirically determined threshold; different signal-tonoise ratio calculations should be compared to determine the method with the lowest number of false-positive and false-negative results (30). Presence or absence of target organisms in a sample is subsequently inferred from presence or absence of the respective phylogenetic probe (set). Besides these basic analysis options, novel bioinformatic approaches have been recently developed for the analysis of one-color hybridization data from diagnostic microarrays. For example, the web application PhyloDetect offers a likelihood-based method for scoring presence or absence of groups of target organisms that can be distinguished by a given diagnostic microarray probe set (31). Microbial communities in environmental samples are generally a complex conglomeration of distantly and closely related organisms. Therefore, even optimized probe/array designs and


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experimental conditions cannot exclude cross-hybridization, which potentially masks the presence of low-abundant target organisms. Another new approach to data analysis from diagnostic microarrays promises a possible solution to this problem (32). A free energy (∆G)-based analytical predictor of theoretical crosshybridization is combined with an optimization algorithm that debunks perfectly-matched probe–target signals from the actual recorded microarray signals. These signals are typically biased by spurious contributions such as cross-hybridization, unequal binding of different perfectly matched probes, background fluorescence, and stochastic variability. The application of this novel mathematical approach in concert with an oligonucleotide microarray targeting the 16 S and/or 23 S rRNA of selected microorganisms allowed the accurate identification of members of the genus Vibrio and quantification of their rRNA molecules in marine samples at their very low natural abundance. It remains to be demonstrated whether this ingenious mathematical approach, including the analytical cross-hybridization predictor, is transferable to other microarray formats that use probes of different lengths and other target labeling procedures. Indeed, there is disagreement on the ability of free energy (∆G)-based mathematical models to predict fluorescence signal intensities of microarray probes (33).

4. Notes 1. Probe–target hybridization efficiency is also a function of probe density (i.e., the number of probe molecules per probe spot area) and the nature (i.e., single or double-stranded RNA or DNA) and length of the labeled target. Pretesting serial probe dilutions and different spotting buffers guarantees optimal probe density. 2. Location of the T3 promoter sequence at the forward primer yields in vitro transcribed RNA with the same orientation as native rRNA. 3. It is generally recommended to operate the spotting device in a clean room, under constant temperature (~20–22°C) and humidity (~50–55%). Regular cleaning of the spotting device (e.g., removal of dust) and especially of the spotting pins (e.g., prior to each use) improves array quality. For low-density microarrays, we recommend using a single pin to avoid pin-to-pin variability (34). 4. Directly after spotting, slides can be simply dried or kept under humid conditions before washing with different buffers (35). After comparing several treatments, we recommend incubation in a humid chamber for efficient cross-linking

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between spotted probes and the aldehyde-coated glass slide. A humid chamber can easily be prepared by filling doubledistilled water into a plastic box. Direct contact between the slides and the water must be avoided, e.g., by using a slide rack. Close the box tightly. 5. Slides can be used up to 3 months of storage, but their quality decreases with storage time. 6. The poly-dTTP spacer of each probe can be exploited as a universal target site for a fully complementary 5¢-Cy3 or 5¢-Cy5-labeled poly-dATP oligonucleotide (e.g., 12mer) for quality control and potentially also for normalization of spotto-spot variability. For quality control, selected slides from a spotting run are hybridized with 2 pmol polyA-oligo at 42°C for 1.5 h (any of the hybridization buffers described in this chapter can be used, but without formamide). Subsequently, slides are washed for 1 min in ice-cold double-distilled water and dried by centrifugation (3 min, 290 × g) prior to scanning. 7. This might lead to a bias called the “position of label” effect (36). 8. Random prime labeling generates short labeled fragments that show reduced secondary structure formation and thus higher probe binding affinities. 9. Use high template concentrations, low PCR cycle numbers, and perform and pool multiple PCRs for each template to ameliorate PCR biases (37). 10. To increase DNA yield at the final step, apply 25 ml of doubledistilled water on the column and wait for 5 min before the centrifugation step. Repeat this step. Alternatively, several PCRs from the same template can be prepared and pooled. The DNA binding capacity of a column from the QIAquick PCR Purification Kit enables purification of up to approximately 100 ml of pooled PCR product. 11. In contrast to previous publications (3, 16), an optimized version of the random prime labeling protocol, yielding significantly higher probe signals after microarray hybridization, is described here. Optimization of the labeling reaction included the adjustment of the ratio of unlabeled and Cy-labeled dCTPs and the implementation of an additional denaturation step. 12. For a newly developed diagnostic microarray, it is not possible to predict a certain formamide concentration, hybridization/ washing temperature, and template concentration which ensures best sensitivity and specificity. Prior to the application of a microarray for the analysis of unknown samples, these parameters need to be carefully evaluated by using defined reference targets (6).


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13. SDS should be added at last to avoid precipitation of buffer reagents. 14. There are different ways to perform microarray hybridization. Generally, continuous mixing of the hybridization solution, by using hybridization devices like the ThermoTWISTER, modified BellyDancer laboratory shaker (38), or fully automated hybridization stations (39), improves the specificity and reproducibility of hybridization results. Alternatively, hybridization can be carried out under static, diffusionlimited conditions without active mixing in self-made (http:// cmgm.stanford.edu/pbrown/mguide/HybChamber.pdf) or purchased (e.g., ArrayIt, TeleChem International Inc.) chambers. In this case, the microarray is placed in the hybridization chamber, overlaid with approximately 20 ml of hybridization mixture and covered with a clean cover slip (rinsed in 100% ethanol and air-dried). 25 ml of hybridization buffer is filled in each of the four cavities in the hybridization chamber. The lid is closed tightly, and the hybridization chamber is incubated in a water bath or hybridization oven at the desired temperature (3). Washing is performed as described in Subheading 3. 15. To prevent cross-hybridization, avoid cooling of the microarray between the hybridization and the washing step by transferring the slides swiftly into the prewarmed washing solution. Washing solutions can be reused for approximately ten slides. 16. General considerations when working with RNA: RNases must be inactivated or removed from all reagents and equipment. It is not sufficient to autoclave tips, tubes, and solutions to inactivate RNases. Incubate all solutions with 0.1–0.2% (v/v) diethylpyrocarbonate (DEPC) under a fume hood (toxic) for 24 h to inactivate RNases. Subsequently, solutions should be autoclaved to destroy DEPC. Solved reagents with primary amino groups, e.g., Tris–HCl cannot be treated with DEPC directly but are prepared with RNasefree water instead. Surfaces should be cleaned with RNase Away (Molecular BioProducts, San Diego, CA, USA). Glassware can be baked at 180°C for 6 h, and plasticware (e.g., tubes) can be treated with 3% H2O2. 17. Do not dry by using a vacuum centrifuge. The RNA pellet should also not be dried completely as this will make it difficult to be dissolved again. 18. For short-time storage (up to 2 months), RNA can be kept at −20°C. For long-term storage, RNA samples should be divided into aliquots and kept at −80°C. Repeated freeze-and thaw cycles may deteriorate RNA quality. 19. Do not mix during incubation because water condensation on the lid of the tube is included in the optimized protocol.

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20. Phylochips used for isotope array analysis must consist of larger probe spots (min. 500 mm in diameter) due to the lower resolution limit of the b-imager (29).

Acknowledgments We acknowledge Stephanie Füreder, Martin Hesselsoe, Sebastian Lücker, Manuel Hofer, and Michael Wagner for their contributions to the development of the microarray protocols presented in this chapter. This work was supported by grants of the German Bundesministerium für Bildung und Forschung (BMBF, BIOTA 01LC0621D) and the Austrian Science Fund (P18836-B17 and P20185-B17) to AL and by a stipend of the Alexander-vonHumboldt foundation to MP. References 1. Wagner M., Smidt H., Loy A., and Zhou J. (2007) Unravelling microbial communities with DNA-microarrays: challenges and future directions Microb Ecol 53, 498–506. 2. DeSantis T. Z., Brodie E. L., Moberg J. P., Zubieta I. X., Piceno Y. M., and Andersen G. L. (2007) High-density universal 16S rRNA microarray analysis reveals broader diversity than typical clone library when sampling the environment. Microb Ecol 53, 371–83. 3. Loy A., Lehner A., Lee N., Adamczyk J., Meier H., Ernst J., Schleifer K.-H., and Wagner M. (2002) Oligonucleotide microarray for 16S rRNA gene-based detection of all recognized lineages of sulfate-reducing prokaryotes in the environment. Appl Environ Microbiol 68, 5064–81. 4. Lehner A., Loy A., Behr T., Gaenge H., Ludwig W., Wagner M., and Schleifer K. H. (2005) Oligonucleotide microarray for identification of Enterococcus species. FEMS Microbiol Lett 246, 133–42. 5. Neufeld J. D., Mohn W. W., and de Lorenzo V. (2006) Composition of microbial communities in hexachlorocyclohexane (HCH) contaminated soils from Spain revealed with a habitat-specific microarray. Environ Microbiol 8, 126–40. 6. Loy A., and Bodrossy L. (2006) Highly parallel microbial diagnostics using oligonucleotide microarrays. Clin Chim Acta 363, 106–19. 7. Wagner M., Nielsen P. H., Loy A., Nielsen J. L., and Daims H. (2006) Linking microbial community structure with function: fluorescence in situ hybridization-microautoradiography and

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Chapter 14 Prenatal Detection of Chromosome Aneuploidy by Quantitative-Fluorescence PCR Kathy Mann, Erwin Petek, and Barbara Pertl Abstract QF-PCR refers to the amplification of chromosome-specific polymorphic microsatellite markers using fluorescence-labelled primers, followed by semi-quantitative analysis of the products on a genetic analyser to determine copy number and/or imbalances of specific chromosomal material. This approach is now widely used for rapid prenatal diagnosis of the common trisomies. In addition, it can successfully detect maternal cell contamination and mosaicism in prenatal material. Key words: QF-PCR, Prenatal, Trisomy, Down syndrome, Mosaicism, Maternal cell contamination

1. Introduction Autosomal chromosome aneuploid pregnancies that survive to term, namely, trisomies 13, 18, and 21, account for 89% of chromosome abnormalities with a severe phenotype (1). They are normally detected by full karyotype analysis of cultured cells. The average UK reporting time for a prenatal karyotype analysis is approximately 14 days (2), and in recent years, there has been increasing demand for more rapid prenatal results with respect to the common chromosome aneuploidies, to relieve maternal anxiety and facilitate options in pregnancy. The rapid tests that have been developed negate the requirement for cultured cells, instead directly test cells from the amniotic fluid (AF) or chorionic villus sample (CVS), with the aim of generating results within 48 h of sample receipt. In the past, interphase fluorescence in situ hybridisation (FISH) (3, 4) was the method of choice in many genetic laboratories usually because the expertise and equipment was readily available. However, a quantitative-fluorescence PCR (QF-PCR)-based approach is more suited to a high-throughput diagnostic service Bimal D.M. Theophilus and Ralph Rapley (eds.), PCR Mutation Detection Protocols, Methods in Molecular Biology, vol. 688, DOI 10.1007/978-1-60761-947-5_14, © Springer Science+Business Media, LLC 2011

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and, following a number of pilot studies (5–9), is now used as a clinical diagnostic service in many European laboratories (10–14). It may be used as a stand-alone test for some referral indications or an adjunct test to full karyotype analysis, which subsequently confirms the rapid result and scans for other chromosome abnormalities not detected by the QF-PCR assay. 1.1. Principle

QF-PCR refers to the amplification of chromosome-specific polymorphic microsatellite markers using fluorescence-labelled primers, followed by semi-quantitative analysis of the products on a genetic analyser to determine copy number and/or imbalances of specific chromosomal material. Tetranucleotide repeat markers are used to minimise PCR-generated “stutter bands” (amplified sequences that are 1–3 repeat units smaller than the true allele size) and both mitotic and meiotic allele instability. Where a microsatellite marker is heterozygous, the ratio of its allele peak areas represents a disomic (1:1) or trisomic (2:1, 1:2 or 1:1:1) chromosome complement (see Fig. 1). A marker is uninformative if only a single peak is observed. Due to allele size heterogeneity and differences in sample type and quality, the amplification of a single marker relative to other markers in the assay may vary greatly. Thus, a comparison of allele peak areas between markers, as an indicator of chromosome copy number, is not recommended. Comparison of allele peak areas within a single locus is more resilient to the effects of DNA quality and the plateau phase of the PCR (15). The procedure described here utilises a “one-tube test,” where 16 markers are co-amplified in one multiplex reaction (see Table 1 and Fig. 1). Five markers are used for both chromosomes 13 and 21, and six for chromosome 18. A separate sex chromosome

Fig. 1. A Genotyper profile of a trisomy 21 sample amplified in the QF-PCR multiplex and analysed on a 3100 genetic analyser. Size in base pairs is shown on the horizontal axis and fluorescent units on the vertical axis. Peaks are labelled with marker name. Heterozygous markers for chromosomes 13 and 18 exhibit two allele peaks with peak areas in a 1:1 ratio. Markers represent trisomy 21 as either three allele peaks in a 1:1:1 ratio (D21S1411) or 2:1 (D21S1435 and D21S1437) or 1:2 (D21S1409 and D21S11) allele ratios.

Location

13q12.1

13q13.3

13q31.1

13q21.33

13q12.12

18q22.1

18q22.3

18p11.31

18q12.3

18q11.2

18q12.3

Marker name

D13S252 F R

D13S305-F -R

D13S628-F -R

D13S634-F -R

D13S325-F -R

D18S386-F -R

D18S390-F -R

D18S391-F -R

D18S535-F -R

D18S819 F R

D18S978-F -R

0.667

0.78

0.92

0.75

0.75

0.875

0.75

0.812

0.688

0.75

0.85

Heterozygosity

180–230

370–450

450–500

190–235

340–415

320–417

235–315

355–440

425–474

418–482

260–330

Size range (bp)

Table 1 Details of primers used in the trisomy multiplex

NED-GTAGATCTTGGGACTTGTCAGA GTCTCCCATGGTCACAATGCT

PET-CTTCTCACCTGAATTACTATGGT TTTGTAATCGATCTACCACAGTT

6-F-CAGCAAACTTCATGTGACAAAAGC CAATGGTAACCTACTATTTACGTC

CTGGCTAATTGAGTTAGATTACAA VIC-GGACTTACCACAGGCAATGTGACT

CTCCAACCTCACTTGAGAGTA NED-GGTCAATAGTGAATATTTGGATAC

VIC-TGAGTCAGGAGAATCACTTGGAAC CTCTTCCATGAAGTAGCTAAGCAG

VIC-CTGTGCTATCTCCTCCAACG GTTTGAAAGATAGGCCATGCAG

6-F-GGCAGATTCAATAGGATAAATAGA GTAACCCCTCAGGTTCTCAAGTCT

NED-TGGATGAATACGCCACTTTTC TGGTTAAAAGATTGCCAAGGAG

TGGTTATAGAGCAGTTAAGGCAC VIC-GCCTGTTTGAGGACCTGTCGTTA

PET-GCAGATGTACTGTTTTCCTACCAA AGATGGTATATTGTGGGACCTTGT

Primer sequences 5¢-3¢

0.18

1

0.5

0.08

0.2

0.7

0.175

0.38

1.7

1

0.675

(continued)

Final concentration (mM) of each primer

Prenatal Detection of Chromosome Aneuploidy by Quantitative-Fluorescence PCR 209

21q21.1

21q21.1

21q21.2

21q22.3

21q21.3

D21S11-F -R

D21S1437-F -R

D21S1409 F R

D21S1411-F -R

D21S1435-F -R

0.75

0.933

0.81

0.84

0.9

Heterozygosity

160–200

256–345

160–220

283–351

220–283

Size range (bp)

6-F-CCCTCTCCAATTGTTTGTCTACC ACAAAAGGAAAGCAAGAGATTTCA

ATAGGTAGATACATAAATATGATGA NED-TATTAATGTGTGTCCTTCCAGGC

PET-GGAGGGGAATACATTTGTGTA TTGCCTCTGAATATCCCTATC

6-F-CTACCACTGATGGACATTTAG CTGGAGGGTGTACCTCCAGAA

6-F-TTTCTCAGTCTCCATAAATATGTG GATGTTGTATTAGTCAATGTTCTC

Primer sequences 5¢-3¢

0.3

0.75

0.5

0.88

0.45


Size ranges given are those used in Genotyper version 3.7 (Applied Biosystems). Heterozygosity values are based on our cohort and may vary in other populations. All markers are tetranucleotide repeats

Location

Marker name

Table 1 (continued)

210 Mann, Petek, and Pertl

Xp22.3

Xp22.3

Xq13.1

Xq26.2

Xq26.2

DXS6807 F R

DXS1283E F R

DXS981 F R

DXS1187 F R

XHPRT

Xp22.2 Yp11.2

F R

F R

AMEL

SRY

Yq11.223

–

–

–

0.87

0.74

0.78

0.72

0.86

0.89

0.7

Het

323–370

248 bp

106 112

240–280

350–420

265–300

125–170

225–260

295–340

300–380

Allele size range (bp)

PET-CAA GGA TCC AAA TAA AGA ACA GAG A GGT TAT TTC TTG ATT CCC TGT G

NED-AGT AAA GGC AAC GTC CAG GAT TTC CGA CGA GGT CGA TAC TTA

NED-CCC TGG GCT CTG TAA AGA ATA GTG ATC AGA GCT TAA ACT GGG AAG CTG

PET-ATG TGG TCT TCT ACT TGT GTC A GTG TGT GGA AGT GAA GGA TAG

VIC-TACTGGAGGTGAGGGTTGTG TGGGCTGCCCAGATACAACT

VIC-ATG CCA CAG ATA ATA CAC ATC CCC CTC TCC AGA ATA GTT AGA TGT AGG

VIC-CAG CTA CTC AAT GAA AAG CC TGA TGG AGA AAG TCA CTG AAC

6-FAM-CTC CTT GTG GCC TTC CTT AAA TG TTC TCT CCA CTT TTC AGA GTC A

NED-AGT TTA GGA GAT TAT CAA GCT G CCC ATA CAC AAG TCC TCA AAG TGA

6-FAM-TCTCCCTTATTTGTGGTTTTGC AGCAGTTCTCCCTTATCCAC

Sequence 5¢-3¢

0.4

0.4

0.2

0.8

1.6

0.6

0.22

0.22

0.8

1.6


Size ranges given are those used in Genotyper version 3.7 (Applied Biosystems). Heterozygosity (Het) values are based on our cohort and may vary in other populations. DYS448 is a hexanucleotide repeat. DX6807, DXS981, DXS1187, XHPRT, DXS7423, and DXYS267 are tetranucleotide repeats, whilst DXS1283E is a dinucleotide repeat. AMEL and SRY are not polymorphic

DYS448 F R

Xq21.31 Yp11.31

DXYS267 F R

Yp11.31

Xq28

DXS7423 F R

F R

Location

Name

Table 2 Details of primers used in the sex chromosome multiplex

Prenatal Detection of Chromosome Aneuploidy by Quantitative-Fluorescence PCR 211

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Fig. 2. (a) A Genotyper profile of a normal male sample. Y-specific sequences are present (AMEL-112 bp, SRY, DYS448), X and Y sequences are present in an equal ratio (AMEL and DXYS267) and all X chromosome markers show a single allele consistent with, though not diagnostic of, a single X chromosome. (b) A Genotyper profile of a normal female sample. Y-specific sequences are absent, and heterozygous markers for the X chromosome exhibit two allele peaks with peak areas in a 1:1 ratio.

multiplex can be used for sexing purposes and to screen for sex chromosome aneuploidy (see Table 2 and Fig. 2) (16). Markers are located along the length of each chromosome to increase the chance of detecting unbalanced chromosome rearrangements. 1.2. Potential Problems 1.2.1. Maternal Cell Contamination

Evidence of a second genotype, as shown by inconsistent dosage ratios for each chromosome and/or extra allele peaks, usually indicates contamination of the sample by maternal cells (see Fig. 3), although it may rarely represent a twin or chimera. Maternal cell contamination (MCC) is usually associated with blood-stained AF samples, although the degree of blood-staining should not in itself be used as an indicator of MCC as blood cells may be foetal or maternal in origin. Usually, samples exhibiting MCC are accompanied by some degree of blood staining, although this ranges from a pale pink cell pellet and clear liquor to a deep-red coloration of the whole fluid. When the majority genotype shows consistent normal or abnormal results with no inconclusive allele ratios, then the

Prenatal Detection of Chromosome Aneuploidy by Quantitative-Fluorescence PCR

213

Fig. 3. A Genotyper profile of a sample exhibiting a high level of maternal cell contamination. The characteristic triallelic pattern, where the foetal-specific and maternal-specific allele peak areas combine to equal the shared foetal/maternal allele, is observed in markers D21S11, D18S391, D18S386, D13S305, D21S1411, D13S628, D13S252, D18S819.

result may be reported, although it may be advisable to confirm the origin of the majority genotype by analysis of a maternal blood sample. We have found the level of blood staining in the amniotic fluid cell pellet to broadly correlate with the level of maternal genotype; the majority genotype from pellets with fewer red blood cells is consistently foetal in origin. However, when the presence of two genotypes causes allele ratios to skew outside of the normal or abnormal range (see Subheading 3.5 in step 1), it is recommended that the QF-PCR results are not interpreted due to the increased risk of a misdiagnosis. In these cases, if a rapid result is required and one of the genotypes is determined as foetal in origin, either by sexing or by genotype analysis of a maternal sample, interphase-FISH may then be used if the analysis takes into account the foetal to maternal ratio; the number of analysed cells can be increased to account for those that are maternal, or a sex chromosome probe can be co-hybridised with an autosome probe and only the male cells analysed. In our sample set from London, UK, approximately 10% of AF samples are found to have two genotypes although allele ratios vary considerably, proportional to the relative contribution of each cell line. The majority of these exhibit a very low level second genotype and can be reported as normal, whilst approximately 2% of AF samples in cohorts from both London and Graz, Austria exhibit second genotypes that prevent confident interpretation of allele ratios and are therefore reported as unsuitable due to MCC. The detection of maternal cells in a blood-stained amniotic fluid sample should not discredit the karyotype analysis of cultured cells. Subsequent genotype analysis of cultured cells from

214


Fig. 4. A Genotyper profile of a sample exhibiting trisomy 21 mosaicism. All informative chromosome 21 markers show either an unequal triallelic pattern (D21S11 and D21S1411) or unequal diallelic ratios (D21S1435, D21S1437 and D21S1409). Chromosome 13 and 18 markers are normal. The presence of triallelic chromosome 21 markers is consistent with a meiotic non-disjunction event followed by trisomy rescue to generate the normal cell line.

samples showing MCC normally demonstrates a single genotype, consistent with the selection and growth of foetal cells and loss of maternal lymphocytes during the culture process (17). In those samples with a mixed female/male cell population evident on the QF-PCR analysis of uncultured cells, a single male genotype is usually detected on testing cultured cells. However, the presence of two genotypes in samples where no blood staining is evident may indicate a maternal tissue plug. In this case, maternal cells may grow in culture, and therefore, genotype analysis of cultured cells may be used to identify MCC. 1.2.2. Mosaicism

The problem of mosaic genotypes and karyotypes in prenatal samples is well documented, particularly in CVS. With respect to QF-PCR, two issues are relevant: the levels of mosaicism detectable by the QF-PCR technique (see Fig. 4) and the degree of concordance between the test result and the foetal genotype. The first of these can only be determined by the analysis of samples (both postnatal and prenatal) exhibiting mosaicism for one of the tested regions. The generation of “artificial mosaics” by the mixing of two genotypes in known measures represents a chimera rather than a mosaic genotype. The presence of a triallelic result is consistent with a meiotic non-disjunction event generating the trisomy cell line, whilst the absence of a triallelic result indicates the possibility of a normal conception followed by a mitotic non-disjunction event, although the likelihood of this depends upon the number of markers used in the assay; using five or ten markers for chromosome 21 resulted in 13.9 or 6.7% of abnormals exhibiting only diallelic results, respectively (unpublished data). Analysis of mosaic cases in our sample set found that


215

a minimum level of 15% trisomy mosaicism could be detected if a triallelic allele pattern was observed, and 20% trisomy mosaicism if only dialleic ratios were present (18). Thus, a mitotic error occurring in a disomic foetus may be harder to detect, due to the absence of a third allele. Indeed, QF-PCR identified only one of three trisomy 18 or 21 mosaics described by Pertl et al. (8); this mosaic case was also triallelic, in this case for chromosome 21. Discrepancies between the QF-PCR and karyotype results have recently been described (19, 20). These have been shown to be due to mosaicism and confinement of cell lines to different regions of the tested sample. In order to minimise such discrepant results, it is recommended that DNA is prepared from dissociated cells prepared from 5 to 15 mg of cleaned villi. Although only a small aliquot of this cell suspension is required, all major cell lines in both the mesoderm and cytotrophoblast should be represented and detected by QF-PCR. Karyotype analysis of cell populations that are subsequently cultured from this cell suspension should minimise discrepancies between the two techniques. Chorionic villi consist of an outer cytotrophoblast layer and internal mesoderm. The mesoderm layer is derived from a later foetal cell lineage, whereas the cytotrophoblast is derived from a much earlier lineage and is thus less representative of foetal tissue (21). DNA prepared from dissociated cells represents cells from both the cytotrophoblast layer and mesenchymal core (22). Interphase-FISH results are also thought to represent both the cell layers. In contrast, karyotype analysis of direct CVS preparations concerns only the cytotrophoblastic line, whilst culture conditions primarily lead to expansion of the mesoderm cell line, resulting in a final karyotype that is more representative of the foetus. In summary, care should be taken in the interpretation of trisomic prenatal results derived from CVS material in the absence of a triallelic result demonstrating a meiotic origin to the trisomy cell line. 1.2.3. Submicroscopic Duplications

Partial chromosome duplication may be identified by QF-PCR analysis by the presence of both normal and abnormal marker results on one chromosome. This pattern may indicate a cytogenetically visible abnormality (11) or one that is submicroscopic. If the most distal or proximal markers are duplicated, then this may indicate the unbalanced product of a reciprocal translocation. However, in our experience, the presence of a single abnormal marker result, whilst all other informative results are normal, is most likely to represent a submicroscopic duplication (SMD) (12). In the majority of cases, analysis of parental samples shows these to be inherited. Inherited SMDs are unlikely to be clinically significant and can be categorised as copy number variants (23). According to the UK Best Practice Guidelines for the Diagnosis of Aneuploidy (http://www.cytogenetics.org.uk/prof_standards/professional_ standards.htm), if an abnormal marker result is flanked by normal

216


markers and if a SMD involving the marker has been observed in a normal individual, the result can be classed as a probable SMD and not reported. For all other single marker abnormalities, it is necessary to establish the inheritance of the SMD. 1.2.4. Primer Site Polymorphisms

Primer site polymorphisms (PSPs) are known phenomena of PCR assays (24). Sequence differences between the genomic DNA and primer can result in complete or partial allele dropout (ADO) due to reduced or nonhybridisation of the primer to genomic DNA. Partial ADO in a normal sample can either give an abnormal diallelic ratio consistent with trisomy for that region or an inconclusive ratio (see Subheading 3.5). Complete ADO in an abnormal sample can result in a normal diallelic ratio at that locus. In all cases of suspected ADO caused by PSPs, it is recommended to repeat the PCR at a lower annealing temperature (for example, 4°C lower than the standard temperature). This provides a less stringent environment for primer hybridisation, resulting in reduced ADO, as represented by a change in the allele ratio. If the follow-up tests are consistent with the presence of a PSP, the marker result should be failed and not used as part of the QF-PCR analysis, even if it shows a normal ratio; a PSP may cause an abnormal diallelic result to appear normal at a lower annealing temperature.

1.2.5. Somatic Microsatellite Mutations

Somatic changes in the length of a microsatellite sequence, due to DNA replication and proof-reading errors, may be visible as an unequal triallelic result, where the areas of the two lowest alleles combine to equal the highest allele (Fig. 5), or as skewed diallelic ratios. The characteristic triallelic pattern represents two cell lines that have one common allele and a second allele of different lengths.

Fig. 5. A Genotyper profile of a sample exhibiting a somatic microsatellite mutation (SMM) at marker D13S305. The characteristic triallelic result is evident where the two lowest allele peaks representing the two mosaic cell lines combine to equal the higher allele present in both cell lines. All other informative markers are normal.


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QF-PCR analysis of AF samples found less than 0.1% to have a somatic microsatellite mutation (SMM) at a single locus, whilst SMMs were observed at a much higher frequency (4%) in individual whole villi (25). This high frequency is probably due to the small clonal cell population being analysed. QF-PCR analysis of digested/chopped villi containing a more diverse cell population (see Subheading 3.1.2) results in the identification of fewer SMMs. Interestingly, the majority of SMMs are single-repeat-unit expansion or contraction events. These somatic changes in repeat length are specific to the microsatellite and are not clinically significant. Thus, a single marker result, showing a characteristic triallelic pattern consistent with a SMM, where all other markers on that chromosome are normal, does not require further clarification. However, it is more difficult to class skewed diallelic results as SMMs; PSPs (see Subheading 1.2.4) and SMDs (Subheading 1.2.3) may be alternative explanations. It is important to distinguish between these events if a 1:2 or 2:1 allele ratio is observed, as these may represent a clinically significant imbalance. Once PSPs have been excluded, analysis of cultured cell populations may help to distinguish between a SMD and SMM. In the case of a SMM, the proportion of the two cell lines may change between uncultured and cultured cells, thus altering the allele ratio.

2. Materials 2.1. Sample Preparation

Sterile water (Sigma, Poole, UK) is used for all procedures. 1. Collagenase: Store powder at −20°C. Resuspend in Hank’s Balanced Salt Solution (HBSS) to a concentration of 2.5 mg/ mL. Store at −20°C. 2. 0.5% Trypsin EDTA: Store at at −20°C. 3. Chang D medium: Store at −20°C.

2.2. D NA Preparation

1. InstaGene Matrix (Bio-Rad, Hercules, CA): Store at 4°C.

2.3. P CR Set-Up

1. T.1E buffer: 10 mM Tris–HCl pH 7.6, 0.1 mM EDTA. Filter-sterilise and store at room temperature. 2. 10× Multiplex PCR Kit (QIAGEN, Hilden, Germany): Store at −20°C. Once thawed, store at 4°C. The kit contains Taq polymerase and dNTPs. 3. 5¢ Labelled fluorescent oligonucleotide primers: The authors use Applied Biosystems (Foster City, CA) primers, 5¢-labelled with 6-FAM, VIC, NED, or PET. Fluorescence-labelled primers should not be exposed to light for prolonged periods of time or repeatedly freeze-thawed (see data sheet). These

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are supplied as precipitates and are resuspended in T.1E buffer to give a working concentration of 100 mM. The primers are stored at −20°C. Aliquots in use are stored at 4°C. 4. Unlabelled oligonucleotide primers: Resuspend in T.1E buffer to a working concentration of 100 mM and store at −20°C. Although unlabelled primers are more stable than fluorescentlabelled primers, freeze-thawing should be avoided. Aliquots in use are stored at 4°C. 2.4. Analysis

1. Capillary-based genetic analyser and associated consumables (e.g. Applied Biosystems 3100): Other genetic analysers capable of fragment resolution, fluorescence detection and quantification can also be used and include Applied Biosystems capillary-based analysers, models 310, 3730, and 3700, Amersham-Pharmacia ALF, and MegaBase systems, although fluorescent labels may have to be substituted depending on the filter sets of each analyser. 2. Deionised formamide (toxic: refer to the material safety data sheet): HI-DI formamide (Applied Biosystems) is highly deionised, to minimise the breakdown of the fluorescent label. Store at −20°C in appropriate aliquots. 3. Genescan-500 LIZ size standard for 100–500 bp fragment analysis. 4. 96-Well plates. 5. Software: Genescan Analysis version 3.7 for fragment analysis, Genotyper version 2.5 (Applied Biosystems) for allele size-calling and labelling, and Microsoft Excel to calculate and tabulate allele dosage ratios.

3. Methods The processing of a number of prenatal samples at one time, and the risk of sample mix-up, necessitates stringent quality control procedures (see Note 1). In addition, care must be taken, as with all PCR-based tests, to avoid contamination of tested material with amplified products of previous reactions and external DNA (see Note 2). 3.1. Sample Preparation

Sample and DNA preparation (steps 1–5, Subheading 3.2) are carried out in a class II biological containment cabinet (see Note 3).

3.1.1. Amniotic Fluid

Between 10 and 20 mL AF is normally received in a 20-mL Sterile Universal Container. The sample is centrifuged at 200 × g for 10 min. Fluid is carefully removed using a 20-mL syringe to leave approximately 1 mL of fluid (equal to the top of the conical


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s ection of the Universal container). The cell pellet is resuspended in the fluid using a plastic Pasteur pipette, and approximately, 100 mL (2–3 drops) is transferred to a 0.5-mL Eppendorf tube. This aliquot (approximately 1/10 of the original AF sample) can be stored at 4°C until processed. The remainder can be cultured for subsequent karyotype analysis, as required. 3.1.2. Chorionic Villus (see Note 4) and Tissue Samples (see Note 5)

Maternal deciduas are removed from 5 to 15 mg of chorionic villi. Cleaned villi are digested with collagenase (final concentration 2.5 mg/mL) at 37°C for 35 min, followed by trypsin digestion (final concentration 2.5 mg/mL) at 37°C for 25 min. The addition of 2 mL of Chang D medium prevents further digestion. A single drop (approximately 100 mL) of the dissociated cell slurry is removed for QF-PCR analysis. This may be stored at 4°C. The rest of the sample is cultured for karyotype analysis. For tissues, a small section of whole tissue sample is isolated and can be stored at 4°C.

3.2. DNA Preparation (see Note 6)

This is a quick and simple procedure and can be successfully applied to a number of sample types and different sample qualities (see Note 7). As it is based on a boiling lysis protocol, aided by vigorous vortexing, the safety issues associated with phenol/chloroform extractions are also avoided. Following cell lysis, a commercial resin (InstaGene Matrix) removes trace metal contaminants that may inhibit the PCR. Prior to the DNA extraction, place the InstaGene Matrix on a magnetic stirrer and set at a medium speed for at least 5 min. 1. Pellet the cells/villus/tissue at 12,000 × g for 1 min in a microcentrifuge and carefully remove the liquid, leaving enough to resuspend the pellet (approximately 10–20 mL). For amniotic fluid samples, any blood staining in the cell pellet is noted as a percentage of whole pellet. If more than 40% of the pellet is red, follow steps 2 and 3. For all other samples, proceed with step 4. 2. Vortex to resuspend the cell pellet and add 200 mL of H2O to wash the sample (see Note 8). 3. Vortex the sample, pellet the cells and remove the wash solution as described in step 1 above. Resuspend the cells in the remaining wash solution by vortexing. 4. Add between 100 and 400 mL of InstaGene Matrix to the cells/villus using a wide-bore pipette tip, e.g. a Gilson p1000 tip and vortex (see Note 9). 5. Incubate at 100°C for 8 min. 6. Vortex again at high speed for 10 s and pellet the InstaGene Matrix at 12,000 × g for 3 min in a microcentrifuge. 7. Place the samples on ice to cool. 8. Store the DNA preparation at −20°C (see Note 10).

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3.3. PCR Set-up

Batches of PCR assays can be prepared in advance, tested and stored at −20°C. These are 20-mL aliquots of a master mix that contain all components except DNA which is added immediately prior to temperature cycling, to give a total volume of 25 mL. The final concentrations of the reaction components are 1× Multiplex PCR Kit and 2.5–42.5 pmol of each primer (see Tables 1 and 2), in a total volume of 25 mL (Note: DNA is added according to volume rather than concentration, as the concentration is not measured). 1. To a thin-walled PCR tube containing 20 mL of the master mix, add 5 mL of DNA solution (see Note 11), taking care not to disturb the InstaGene Matrix pellet. Mix by pipetting. 2. Add one drop of mineral oil if a heated lid is not being used and place it in the PCR machine. 3. Perform a PCR with the following cycling conditions: Taq polymerase activation and initial denaturation at 95°C for 15 min followed by 25 cycles of 94°C for 30 s, 58°C for 90 s, 71°C for 90 s (see Note 12), and a final incubation of 72°C for 20 min followed by storage at 10°C (see Note 13).

3.4. Analysis 3.4.1. PCR Product Preparation

Post-PCR clean-up to remove excess primers and free dye molecules is not normally carried out (see Note 14). 1. Prepare PCR products for analysis by the addition of 3 mL of product to 15 mL of HI-DI formamide in 96-well plates (see Note 15). 2. Denature at 95°C for 2 min and snap-chill on ice.

3.4.2. 3100 Analysis

Separate PCR products through a 36-cm capillary array filled with POP6 (see Note 16). A 10-s injection time is suitable for most samples (see Note 17). The running conditions are 60°C for 3,000 s.

3.4.3. Genotyper Analysis

Macros are used to label allele peaks with marker name, size, and peak area (Fig. 1) (see Note 18). The Genotyper table is transferred to an Excel spreadsheet for allele ratio analysis.


The criteria listed below are based on more than 19,000 QF-PCR prenatal tests, all of which were followed by karyotype analysis of cultured cells (see Note 19). For additional information, see the UK Best Practice Guidelines for the Diagnosis of Aneuploidy, available at http://www.cytogenetics.org.uk/prof_standards/ professional_standards.htm 1. Normal allele dosage ratios range between 0.8 and 1.4 (see Note 20). For normal alleles separated by more than 24 bp, ratios up to 1.5 are acceptable. Trisomy is indicated by an


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allele ratio between 1.8 and 2.4 or between 0.65 and 0.45, or by the presence of three alleles of equal areas (see Note 21). All of these results are classified as informative. 2. Two informative markers per chromosome are required for confident interpretation. This minimises the risk of misdiagnosis due to primer-site polymorphisms, SMDs and/or somatic repeat instability (see Subheadings 1.2.3, 1.2.4, and 1.2.5). Single marker abnormal results should be verified by the testing of additional markers or other techniques. 3. If both normal and abnormal marker results are obtained for a single chromosome, this may represent polymorphisms or a clinically significant partial chromosome imbalance (see Subheadings 1.2.3, 1.2.4, and 1.2.5). These results may be clarified by single marker assays, additional markers, lowering the PCR annealing temperature, and the analysis of cultured cell populations and parental samples.

4. Notes 1. To prevent sample mix-up, a minimal number of tube-to-tube transfers should be employed (three transfers are required for this protocol). Each sample transfer and analysis should be checked by another laboratory member, and the use of two identifiers per tube, such as sample number and name, aids sample tracking. 2. Contamination of a PCR by external DNA is evident by the appearance of allele peaks in the negative (no DNA) PCR control, which is a critical part of any PCR procedure and a reaction which should be set up last in a series of samples. Separation of the PCR set-up and post-PCR analysis areas will help to prevent contamination. 3. To ensure that DNA is prepared from the correct sample, it is advisable to prepare the initial sample aliquot (see Subheadings 3.1.1 and 3.1.2) one sample at a time in a class II biological containment cabinet, with only one sample in the cabinet during the procedure. As the subsequent DNA preparation (see Subheading 3.2) is carried out without further tube transfers, DNA from a number of samples can be prepared simultaneously. This DNA can then be used for subsequent PCR tests. 4. It is recommended that DNA is prepared from dissociated cells prepared from 5 to 15 mg of cleaned villi. Although only a small aliquot of this cell suspension is required, all major cell lines in both the mesoderm and cytotrophoblast

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should be represented and detected by QF-PCR. The analysis of whole villi has rarely been associated with discrepant results between QF-PCR and karyotype analysis since mosaic cell lines are confined to one region of the sample (19, 20). Analysis of cell populations that are subsequently cultured for karyotype analysis from the dissociated cell preparations should minimise discrepancies between the two techniques. 5. The QF-PCR procedure can be applied effectively to solid tissue samples (e.g. skin and cartilage). This is particularly useful for confirming a prenatal diagnosis or for a poor quality sample where chromosome analysis of cultured cells may not be possible. 6. DNA prepared using the protocols described here may also be used for other molecular prenatal tests. The protocol has benefits over the traditional phenol/chloroform-based approach in terms of reduced labour, time, and safety risks. However, as the extracted DNA may contain residual contaminants, its suitability for use in each test should be determined. 7. Although the procedure is generally successful in extracting DNA of sufficient quality for use in the multiplex PCR, in our experience, DNA extracted from blood-stained or discoloured AF fluid may contain PCR inhibitors. These can be removed by a subsequent extraction (see Note 9). 8. Deionised water lyses red blood cells and may also aid lysis of cells in villus and tissue samples. A deionised water wash is used for blood-stained/discoloured AF to remove haem that may inhibit PCR. 9. It is beneficial to adjust the volume of InstaGene Matrix to balance removal of all cell lysis products with excessive dilution of the DNA. A 300-mL volume of InstaGene Matrix is generally used for CVS and tissue samples and the larger AF cell pellets (those that cover the base of the 0.5-mL microcentrifuge tube). Only 100 mL of InstaGene Matrix is required for average and small AF pellets. It is important that the DNA extraction is not overloaded with too much of starting material. This leads to a failure by the InstaGene Matrix to chelate all metal ions and can result in inhibition of the PCR. In particular, the larger sized markers in the multiplex may fail to amplify. If inhibition is observed, a further extraction can be used to remove the contaminants; 100 mL of the DNA extract is added to 100 mL of InstaGene Matrix and treated as per the extraction protocol (see Subheading 3.2; steps 5–9). 10. As the DNA prepared here is relatively crude and cell lysis products that damage DNA may remain, the DNA should be


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stored at −20°C. However, the DNA does appear to be stable for at least three days at room temperature which allows some flexibility, such as transfer of the sample to another laboratory. 11. As well as the necessary inclusion of the negative (no DNA) PCR control for the reasons given in Note 2, the use of a DNA control trisomic for one of the chromosomes and exhibiting 2:1 or 1:2 dosage for at least one marker is recommended. Normal allele dosage is exhibited by the other two non-trisomic chromosomes. The control not only demonstrates that the amplified DNA represents allele copy number for that procedure, but it can also be used as a standard against which any spurious background bands or free-dye peaks can be compared. 12. Samples that do not generate sufficient amplified sequences for analysis due to low initial DNA concentration (usually evident by a very small original cell pellet) can be amplified with a greater number of cycles. Any change in PCR cycling conditions should always be accompanied by a trisomy control, to ensure the reaction is still quantitative. 13. Taq polymerases lacking exonuclease activity add a templateindependent deoxyadenosine triphosphate (dATP) to the 3′ end of amplified sequences (26). The 72°C incubation for 20 min ensures the addition of dATP to all amplified sequences. Without it, a single base-pair size difference in the amplified sequences can resolve as a “split peak” on the profile and hinder analysis, especially in smaller alleles. This is a particular problem here due to the small size range of the fragments generated (100–500 bp). 14. Free-dye peaks are caused by the detachment of the fluorescent molecule from the labelled primer. These molecules are resolved as broad peaks, usually up to 180 bp in size (see Note 16), and as such can be distinguished from allele peaks. The breakdown of fluorescent primers can be minimised by the use of deionised formamide stored at −20°C, and reduced exposure of labelled primers to temperatures above −20°C. Free-dye molecules can be removed, along with unincorporated primers, by standard post-PCR clean-up protocols if required. 15. Accurate transfer of samples to the wells of a 96-well plate can be difficult. The risk of error can be minimised by the use of a multichannel pipette and by the addition of loading buffer containing dextran blue, which is visible but does not interfere with the fluorescent analysis. Also, transparent pierceable sheets are available that can be sealed onto the plate, or rubber septa can be placed over wells that are not in use to further minimise pipette transfer errors.

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16. POP6 polymer can be used on the 3100 genetic analyser to improve resolution, although this requires longer run times. Resolution of free-dye molecules (see Note 14) is not linear in respect to fragment size but is influenced by both temperature and the separating matrix. A different polymer may be used to resolve free dye molecules that coincide with allele peaks. 17. Allele peak heights greater than 6,000 fluorescent units on the 3100 genetic analyser are not analysed. The CCD camera becomes saturated in this range, and peak fluorescence may be under-represented. If a sample is overloaded, injection times can be reduced to accommodate differences in DNA concentration and the corresponding amplification. This is one of the advantages of the capillary-based genetic analysers, where a repeat injection does not require repeat sample preparation. 18. Tetranucleotide alleles demonstrate few visible stutter bands (see Subheading 1.1), and only the main allele peak is labelled. However, some microsatellite markers contain a mix of both tetranucleotide and dinucleotide repeats and generate significant stutter bands. For dinucleotide alleles, the longer alleles generally exhibit more significant stutter effects than shorter alleles, due to the longer repeat. It is, therefore, necessary to recognise and label at least the first stutter peak and include it in the allele peak area measurement. 19. Although there are now a number of published studies describing the use of QF-PCR as a diagnostic test (5–14) and commercial assays are available, it is important to validate the QF-PCR strategy in the laboratory in which it is to be used. Control samples are required, and a pilot study is recommended prior to the implementation of a QF-PCR based aneuploidy diagnostic service, especially if primer sets are used that are not described in the published literature. 20. The large normal range is necessary due to the use of tetranucleotide repeats. These can result in widely spaced alleles (up to 50 bp apart) and marked preferential amplification of the smaller allele which in turn results in skewed allele dosage ratios. However, closely spaced alleles should exhibit less allele specific preferential amplification and would be expected to have dosage ratios closer to 1.0. 21. As more than one sample is usually processed, the sample identity of abnormal results should be confirmed. This can be done by a repeat QF-PCR test or by another technique such as interphase-FISH or karyotype analysis of uncultured or cultured cells. Alternatively, genotype analysis of a maternal blood sample using the same markers can be used to confirm sample identification.


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References 1. Lewin, P., Kleinfinger, P., Bazin, A., Mossafa, H., Szpiro-Tapia, S. (2000) Defining the efficiency of fluorescence in situ hybridization on uncultured amniocytes on a retrospective cohort of 27 407 prenatal diagnoses. Prenat Diagn. 20, 1–6. 2. Waters, J. J., Waters, K. S. (1999) Trends in cytogenetic prenatal diagnosis in the UK: results from UKNEQAS External Audit, 1987–1998. Prenat Diagn. 19, 1023–1026. 3. Spathas, D. H., Divane, A., Maniatis, G. M., Ferguson-Smith, M. E., Ferguson-Smith, M. A. (1984) Prenatal detection of trisomy 21 in uncultured amniocytes by fluorescence in situ hybridisation: a prospective study. Prenat Diagn. 14, 1049–1054. 4. Klinger, K., Landes, G., Shook, D., et al. (1992) Rapid detection of chromosomal aneuploidies in uncultured amniocytes by using fluorescence in-situ hybridization (FISH). Am J Hum Genet. 51, 55–65. 5. Pertl, B., Yau, S. C., Sherlock, J., Davies, A. F., Mathew, C. G., Adinolfi, M. (1994) Rapid molecular method for prenatal detection of Down’s syndrome. Lancet. 343, 1197–1198. 6. Pertl, B., Kopp, S., Kroisel, P. M., Hausler, M., Sherlock, J., Winter, R., Adinolfi, M. (1997) Quantitative fluorescence polymerase chain reaction for the rapid prenatal detection of common aneuploidies and fetal sex. Am J Obstet Gynecol. 177, 899–906. 7. Verma, L., Macdonald, F., Leedham, P., McConachie, M., Dhanjal, S., Hulten, M. (1998) Rapid and simple prenatal DNA diagnosis of Down’s syndrome. Lancet 352, 9–12. 8. Pertl, B., Kopp, S., Kroisel, P. M., Tului, L., Brambati, B., Adinolfi, M. (1999) Rapid detection of chromosome aneuploidies by quantitative fluorescence PCR: first application on 247 chorionic villus samples. J Med Genet. 36, 300–303. 9. Schmidt, W., Jenderny, J., Hecher, K., Hackeloer, B. -J., Kerber, S., Kochhan, L., Held, K. R. (2000) Detection of aneuploidy in chromosome X, Y, 13, 18 and 21 by QF-PCR in 662 selected pregnancies at risk. Mol Hum Reprod. 6, 855–860. 10. Levett, L. J., Liddle, S., Meredith, R. (2001) A large-scale evaluation of amnio-PCR for the rapid prenatal diagnosis of fetal trisomy. Ultrasound Obstet Gynecol. 17, 115–118. 11. Mann, K., Fox, S. P., Abbs, S. J., Yau, S. C., Scriven, P. N., Docherty, Z., Ogilvie, C. M. (2001) Development and implementation of a new rapid aneuploidy diagnostic service within the UK National Health Service and

12.

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implications for the future of prenatal diagnosis. Lancet. 358, 1057–1061. Mann, K., Donaghue, C., Fox, S. P., Docherty, Z., Ogilvie, C. M. (2004) Strategies for the rapid prenatal diagnosis of chromosome aneuploidy. Eur J Hum Genet. 12, 907–915. Cirigliano, V., Voglino, G., Canadas, M. P., Marongiu, A., Ejarque, M., Ordonez, E., Plaja, A., Massobrio, M., Todros, T., Fuster, C., Campogrande, M., Egozcue, J., Adinolfi, M. (2004) Rapid prenatal diagnosis of common chromosome aneuploidies by QF- PCR. Assessment on 18,000 consecutive clinical samples. Mol Hum Reprod. 10, 839–846. Ogilvie, C. M., Donaghue, C., Fox, S. P., Docherty, Z., Mann, K. (2005) Rapid prenatal diagnosis of aneuploidy using quantitative fluorescence-PCR (QF-PCR). J Histochem Cytochem. 53, 285–288. Lubin, M. B., Elashoff, J. D., Wang, S. -J., Rotter, J. I., Toyoda, H. (1991) Precise gene dosage determination by polymerase chain reaction: theory, methodology, and statistical approach. Mol Cell Probes. 5, 307–317. Donaghue, C., Roberts, A., Mann, K., Ogilvie, C. M. (2003) Development and targeted application of a rapid QF-PCR test for sex chromosome imbalance. Prenat Diagn. 3, 201–210. Stojilkovic-Mikic, T., Mann, K., Docherty, Z., Mackie Ogilvie, C. (2005) Maternal cell contamination of prenatal samples assessed by QF-PCR genotyping. Prenat Diagn. 25, 79–83. Donaghue, C., Mann, K., Docherty, Z., Ogilvie, C.M. (2005) Detection of mosaicism for primary trisomies in prenatal samples by QF-PCR and karyotype analysis. Prenat Diagn. 25, 65–72. Waters, J. J., Walsh, S., Levett, L. J., Liddle, S., Akinfenwa, Y. (2006) Complete discrepancy between abnormal fetal karyotypes predicted by QF-PCR rapid testing and karyotyped cultured cells in a first-trimester CVS. Prenat Diagn. 26, 892–897. Waters, J. J., Mann, K., Grimsley, L., Mackie Ogilvie, C., Donaghue, C., Staples, L., Hills, A., Adams, T., Wilson, C. (2007) Complete discrepancy between QF-PCR analysis of uncultured villi and karyotyping of cultured cells in the prenatal diagnosis of trisomy 21 in three CVS. Prenat Diagn. 27, 332–339. Gardner, R. J. M., Sutherland, G. R. (1996) Chromosome Abnormalities and Genetic Counseling. Oxford University Press, Oxford, UK.

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22. Mann, K., Kabba, M., Donaghue, C., Hills, A., Mackie Ogilvie, C. (2007) Analysis of a mosaic placenta to assess the cell populations in dissociated CVS and implications for QF-PCR analysis. Prenat Diagn. 27, 287–289. 23. Sharp, A. J., Locke, D. P., McGrath, S. D., Cheng, Z., Bailey, J. A., Vallente, R. U., Pertz, L. M., Clark, R. A., Schwartz, S., Segraves, R., Oseroff, V.V., Albertson, D. G., Pinkel, D., Eichler, E. E. (2005) Segmental duplications and copy-number variation in the human genome. Am J Hum Genet. 77, 78–88.

24. Andrew, S. E., Whiteside, D., Buzin, C., Greenberg, C., Spriggs, E. (2002) An intronic polymorphism of the hMLH1 gene contributes toward incomplete genetic testing for HNPCC. Genet Test. 6, 319–322. 25. Mann, K., Donaghue, C., Ogilvie, C. M. (2003) In vivo somatic microsatellite mutations identified in non-malignant human tissue. Hum Genet. 114, 110–114. 26. Clark, J. M. (1988) Novel non-templated nucleotide addition reactions catalyzed by procaryotic and eucaryotic DNA polymerases. Nucleic Acids Res. 16, 9677–9686.

Chapter 15 Use of Robotics in High-Throughput DNA Sequencing Stephen Keeney Abstract Until relatively recently, full sequencing of genes consisting of more than several exons was not considered practicable within a routine diagnostic context. As a result, many approaches to unknown mutation detection in a specific gene involved a mutation pre-screening step to limit the amount of DNA sequencing required. Protocols to pre-screen for mutations and limit the amount of DNA sequencing may not localise every base change present and/or require considerable levels of manual intervention. Advances in technology, allied with careful protocol design, now permit direct DNA sequencing to be applied to larger areas of gene sequence, allowing unequivocal mutation identification in the area of a gene being analysed. The protocol described below utilises robotic systems, allied to custom-designed PCR primers, to facilitate rapid DNA sequencing of multiple gene targets. The general approach is amenable to adaptation for use with multi-channel pipettes. Key words: Robotics, Cycle sequencing, Mutation analysis, Primer design, DNA purification

1. Introduction DNA sequencing is considered to be the gold standard for detection of point mutations associated with genetic disease. Many human genes of biological interest consist of multiple exons dispersed over many kilobases of genomic DNA. This presents a practical problem for rapid and accurate mutation detection when using the current generation of automated DNA sequencers applied to routine genetic diagnosis which are designed to sequence up to 1,000 bases per individual channel. Protocols to sequence the essential regions of a gene (considered to include the promoter region, exons, intron/exon boundaries, and the 5¢ and 3¢ untranslated regions) must take this structure into account. In practice, this means designing multiple primer sets targeted at these essential regions. For example, the relatively large FVIII gene (F8) associated


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with haemophilia A consists of 26 exons with some exons requiring the design of overlapping primer sets due to their size. For the protocol described below this translates into 37 individual PCRs to create template DNA suitable for sequencing which, if applied to the forward and reverse strands, translates into 74 cycle sequencing reactions, excluding controls. The use of robotic liquid handling systems allied with good protocol design can overcome many of the difficulties presented by these practical limitations. The principles described in this protocol can be applied to design an automation-friendly approach allowing highthroughput sequencing in the timescale required for routine diagnostic mutation analysis.

2. Materials 2.1. General Considerations

There are several fundamental elements that need to be considered when designing and implementing a high-throughput sequencing analysis protocol.

2.1.1. Choice of Robotic Platforms

There are many automated liquid handling systems in the market, and the scale and complexity of the chosen platform(s) will depend on the degree of automation to be applied, the number of ancillary steps to be undertaken in a fully automated fashion, and the required throughput. The protocol below assumes a single gene analysis, which can be accommodated in a standard 96-well microtitre plate format. 384- and 1,536-well plate formats are also available for medium to very high throughputs. Robotic platforms capable of dispensing nanolitre quantities of reagents are available, allowing cost reduction benefits to be realised which may be of particular importance in very high-throughput systems. Another key parameter controlling throughput will be the number of liquid dispensing probes the robotic platform has. This can range from a single probe, dispensing into one well at a time, up to 384-well heads, which will perform an individual dispense step across a plate in a fraction of the time. Ancillary equipment linked to the robotic platform may include a plate storage “hotel,” centrifuge, and thermal cycler. However, for many applications, performing these steps on “stand alone” equipment is perfectly adequate.

2.1.2. PCR and Sequencing Primer Design

Successful automation of direct sequencing protocols involving multiple sequencing targets requires careful consideration of primer design. A primer set designed to amplify a given gene prior to DNA sequence analysis should have as similar characteristics as possible. Particular attention should be given to ensuring that primer melting temperatures (Tm) are matched across the set.

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The only practicable way to achieve close Tm tolerances across multiple primer sets is to use primer design software. A readily available tool suitable for this purpose is Primer3, which will allow very precise control of primer parameters (1). Primers should have as high an annealing temperature as possible that will be tolerated across the regions being amplified. A good starting point is to select primers with a target Tm of 59°C with a maximum tolerated Tm difference of plus or −1°C. If this cannot be accommodated across the set then wider Tm values can be considered, but ideally these should be within 2 or 3° of the mean. Remember that careful design maximises the likelihood that the entire primer set will amplify under a common thermal cycling protocol but ultimately this will have to be determined empirically and may require some primer redesign or optimisation around a higher or lower Tm, depending on the nature of the target sequence. As well as sequence-specific primer design, automationfriendly approaches should include common tails to simplify cycle sequencing set-up. The forward primers for each PCR have a common tail to allow the use of one complementary sequencing primer in all forward sequencing reactions. All reverse primers have a second common tail for reverse strand sequencing. The benefits of this approach are obvious as it allows the preparation of only two sequencing master mixes, one containing the complementary forward primer and the other the complementary reverse primer, facilitating bidirectional sequencing without the need for multiple primer-specific master mixes. Common tails in use are M13-derived or a modified version of these (known as N13 or UniSeq primers), which can tolerate higher annealing temperatures in cycle sequencing reactions, a feature that may be useful for sequencing templates with strong secondary structure. Each primer pair will have the relevant universal primer sequences attached – the forward primer the N13F sequence and the reverse primer the N13R sequence. The relevant primer tail sequences are as follows: N13F 5¢ GTA GCG CGA CGG CCA GT 3¢ N13R 5¢ CAG GGC GCA GCG ATG AC 3¢ Example primer sequences, illustrating the use of tails, for amplification of part of F8 are shown in Table 1. For further advice on primer design, see the National Genetics Reference Laboratory (Wessex) application note Standardised Primer Optimisation and Design Specification (2). 2.1.3. Template DNA Source

DNA suitable for mutation analysis by direct cycle sequencing must be of sufficient quality to allow good PCR amplification. The method used to isolate DNA will depend on the scale of the procedure, the tissue source, and the degree of automation applied. Most robotic-based DNA extraction systems will produce relatively small amounts of DNA which may be insufficient

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Table 1 Example oligonucleotide primer sequences

Primer set

Primer sequences (N13 F and R standard tails are in italics)

Product size (bp) (size inc. N13 tails)

F8P2F F8P2R

gtagcgcgacggccagtGAGCTCACCATGGCTACATTC cagggcgcagcgatgacAATTTAAAACTATAAAGCGAGTCCTG

561 (595)

F8P1F F8P1R

gtagcgcgacggccagtGGACCTAGGCCATGGTAAAGA cagggcgcagcgatgacTGCAGAGCATTTTAAGGAACTTT

600 (634)

F8EX1F F8EX1R

gtagcgcgacggccagtTAGCAGCCTCCCTTTTGCTA cagggcgcagcgatgacCTAACCCGATGTCTGCACCT

480 (514)

F8EX2F F8EX2R

gtagcgcgacggccagtCATTACTTCCAGCTGCTTTTTG cagggcgcagcgatgacTTTGGCAGCTGCACTTTTTA

290 (333)

Example primers for mutation analysis of part of F8 illustrating the use of N13F (all primers with the suffix “F”) and N13R (suffix “R”). The standard tails are in lower case italics. For the full set of primers, see ref. (3)

for the multiple PCR amplifications required when sequencing a larger gene. For this protocol, the DNA has been purified beforehand in a bulk extraction procedure. 2.2. PCR Amplification

1. Deoxynucleotide triphosphates (dNTPs), PCR grade. Prepare stocks of each dNTP at a concentration of 10 mM in sterile water. A working stock solution can be prepared containing equal volumes of each dNTP (i.e. 2.5 mM each dNTP, 10 mM total). Use 25 mL of each dNTP and add to 900 mL of water. Store at −20°C. 2. Taq DNA Polymerase, native (5 U/mL activity), 10× Reaction buffer, 50 mM MgCl2 (Invitrogen, UK). The 10× reaction buffer and 50 mM MgCl2 are provided with the Taq. Store each reagent at −20°C (see Note 1). 3. Sterile water: This should be high-quality water. 10 mL vials designed for parenteral use are ideal. Store at room temperature. 4. Tailed PCR primers specific for the application (see examples in Table 1): These can be obtained from a variety of commercial primer suppliers. Store master primer stocks at −80°C. Primers are diluted to a concentration of 2.5 mM (see Note 2). Working stocks can be made by combining equal volumes of individual primer. If desired, these working stocks can be stored at −20°C.

2.3. Agarose Gel Electrophoresis

1. Molecular-Biology-Grade agarose powder. 2. 10× Tris Borate EDTA buffer (TBE): 108 g Tris, 55 g boric acid, 40 mL 0.5 M EDTA, pH 8.0. Make up to 1 L with


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distilled water. For use, dilute to 0.5× concentration (50 mL/ 950 mL of water). Store at room temperature. 3. Sample Loading Buffer (5×): 4.8 mL Glycerol, 0.025 g bromophenol blue, 0.2 mL 20% (w/v) SDS, 5 mL 10× TBE. Store at room temperature. 4. Ethidium bromide: 10 mg/mL in sterile water (this is considered to be a mutagen so observe appropriate safety precautions). Store at room temperature. 2.4. PCR Product Purification

2.5. Cycle Sequencing

MicroCLEAN reagent (MicroZone, UK): MicroCLEAN is a “one-tube” DNA clean-up reagent. It can be used to clean up any type of double-stranded DNA from reaction buffers, enzymes, dNTPS, and primers. There is no upper limit on the size of the target DNA fragment undergoing purification. The simplicity of this reagent makes it well suited to robotic applications; however, note that the use of MicroCLEAN involves centrifugation steps (see Note 3). 1. Applied Biosystems BigDye Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems, UK). Store at −20°C. This contains dye terminators, dNTPs, AmpliTaq DNA Polymerase FS, magnesium chloride, and buffer in a single premix. The fluorescent dyes in this kit are light sensitive. Keep in the dark as much as practicable. 2. Applied Biosystems Sequencing Buffer 5× (Applied Biosystems, UK). Store at 4°C. 3. N13 standard forward and reverse cycle sequencing primers N13F – GTAGCGCGACGGCCAGT, N13R – CAGGGCGCA GCGATGAC. These are complementary to the standardised primer tails (see Subheading 2.1.2). Dilute to a working concentration of 1 pmol/mL (see Note 4). Store at −20°C. 4. Cycle sequencing template DNA: The MicroCLEAN purified PCR products to be sequenced. Store at 4°C prior to use.

2.6. Purification and Resuspension of Cycle Sequencing Products

2.7. DNA Sequence Analysis

1. 95 and 70% Ethanol (see Note 5). 2. 125 mM EDTA: Store at room temperature. 3. Applied Biosystems Hi-Di Formamide (Applied Biosystems, UK). Store at −20°C. Applied Biosystems POP-7 Polymer (Applied Biosystems, UK). Stored at 4°C (see Note 6). Applied Biosystems 10× Running Buffer (Applied Biosystems, UK). Store at 4°C.

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3. Methods All reagents should be pre-prepared in vessels compatible with the robotic platform in use. The following protocol describes robotic preparation and amplification of F8 in a series of 37 individual PCRs covering the essential regions of the gene in a 96-well microtitre plate (3). The specific robotic platforms used are MWG Biotech/Aviso RoboAmp 4200 workstations (Aviso, Germany). This protocol uses two robots to allow for pre- and post-PCR manipulation of material being analysed. This provides high safeguards against cross-contamination but bear in mind that the protocol described below is amenable to the use of multi-channel pipettes. 3.1. Preparation of Reagents for High-Throughput DNA Sequencing Using Robotics

1. The 37 combined working stock primer sets (2.5 mM each primer) are stored at −20°C in 1.5 mL robot-compatible tubes ready to aliquot into plates prior to amplification. Defrost and mix by gentle vortexing. 2. Creating plates containing tailed primers to which a master mix can be added: Using a program on the pre-PCR robot, aliquot 6 mL of each of the 37 combined F8 primer pairs from the 1.5 mL tubes into a 96-well microtitre plate. Include an extra tube containing sterile water to act as a form of negative control (see Note 7). This creates 38 wells, which will be involved in the subsequent analysis process. Plates containing dispensed PCR primers and blank control can be pre-prepared, sealed and stored at −80°C until required. 3. Prepare a PCR master mix (see Table 2) either on the robot in a workstation position cooled to 4°C or, if prepared manually, on ice.

Table 2 PCR master mix for initial amplification of the essential regions of F8 Component

1× reaction (mL)

40× reaction master mix (mL)

10× PCR Buffer

2.5

100

Magnesium Chloride (50 mM)

0.75

30

dNTPS (10 mM)

2

80

Taq Polymerase (keep on ice)

0.1

4

Sterile water

12.65

506

Genomic DNA (100–200 ng equivalent)

1

40


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4. Take a pre-dispensed primer plate out of the freezer, defrost at room temperature and place on a workstation position cooled to 4°C. 3.2. PCR Amplification of Target Gene

Using an appropriate program, set up reactions on the pre-PCR robot platform. On run activation, the robot aliquots 19 mL of the pre-prepared master mix per well into each of the pre-dispensed 6 mL specific primer sets and mixes, creating a final reaction volume of 25 mL per well. Once the robot has completed this step, the plate can be transferred to either an on-board or stand-alone thermal cycler with a heated lid as desired. All 38 individual PCRs for the F8 gene protocol are performed in a single 96-well plate under identical conditions: 95°C for 2 min followed by 30 cycles of 94°C for 30 s, 59°C for 45 s, and 72°C for 1 min 30 s. Final extension of 72°C for 5 min (see Note 8).

3.2.1. Checking PCR Amplification

It is advisable to ensure that successful amplification of the target gene has been achieved due to the expense of subsequent processing steps. Prior to the PCR purification step, analyse 5 mL of each PCR product on an agarose check gel (see Note 9). Pour a 1.5 % gel (1.5 g agarose into 100 mL of 0.5× TBE, heat until dissolved and cool to 60°C). Take care when heating and handling hot agarose gel solutions, as they may superheat and boil over when mixed. Add 2 mL of 10 mg/mL ethidium bromide and pour. Use two 20-well, 1 mm combs in the gel and leave to set at room temperature for 45 min. Transfer the plate containing PCR amplified products (PCR Plate) to a post-PCR robot and place a second, empty plate on another workstation position. Run a robot program that dispenses 1 mL of 5× Sample Loading Buffer and 3 mL of water (both from individual 1.5 mL robot compatible tubes on the workstation) into the second plate, followed by transfer of 5 mL of each PCR product from the PCR Plate to the second plate with a mix step. Load 8 mL of the contents of this second plate onto the 1.5% agarose gel, preferably using a multichannel pipette, run at 100 V for 1 h and photograph under UV transillumination. Assuming that amplification has been successful and that the negative control lane is blank, proceed to the following step.

3.3. PCR Product (Sequencing Template) Purification Step

Prior to DNA sequencing, the PCR products (each template DNA) must be purified (see Note 3). Ideally, this is performed on a postPCR robotic workstation, but if necessary, the following steps can be performed using a multi-channel pipette. 1. Aliquot 800 mL of MicroCLEAN reagent and 800 mL of sterile water into separate 1.5-mL robot compatible tubes and place in the appropriate workstation positions on a post-PCR robot. Place the PCR Plate on the robot.

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2. Run a program that dispenses 20 mL of MicroCLEAN reagent into each well of the 38 wells of the PCR Plate containing the remaining 20 mL of each PCR amplified product, with a mixing step. 3. Take the plate off the robot, seal with a plate sealing film or silicone mat and vortex (see Note 10). Leave for 5 min at room temperature and then spin the plate at 2,700 × g for 30 min to pellet the PCR product. 4. Remove supernatant by inverting the plate onto a paper towel. 5. Remove remaining traces of the supernatant by centrifugation of the inverted plate/fresh paper towel at 40 × g for 30 s. 6. Place the PCR Plate back on the robotic workstation and dispense 20 mL of sterile water into each of the 38 wells. 7. For forward and reverse strand sequencing, instruct the robot to dispense 4 mL aliquots of each purified template DNA into two adjacent wells of a second plate, which is compatible with the automated DNA sequencer being used. This creates 76 wells that contain either template or negative controls. The robot program should then pause and wait for further instructions for automated sequencing reaction setup (see Subheading 3.4). 3.4. DNA Cycle Sequencing in a 96-Well Plate Format

1. Set up the cycle sequencing reaction in the sequencer compatible 96-well plate containing 4 mL aliquots of the forward and reverse sequencing templates (Sequencing Plate) by preparing appropriate master mixes (Table 3). 2. Place the forward and reverse cycle sequencing master mixes in separate 1.5 mL robot compatible tubes and place on the

Table 3 Cycle sequencing master mix for initial amplification of the essential regions of the F8 gene (see Note 11) Forward and Reverse cycle sequencing reactions (prepared at one-eighth AB BigDye reaction strength) Component

Per reaction (mL)

Per 40 reaction forward or reverse master mix

AB BigDye v3.1 mix

1

40 mL

AB Cycle Sequencing Buffer (5×)

2.5

100 mL

N13 Forward or Reverse primer (2 mM)

1.5

60 mL of N13 Forward or Reverse primer

Sterile water (to make the volume to 11 mL)a

6

240 mL

Final volume will be 15 mL per reaction after addition of 4 mL of template DNA

a


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post-PCR robot with the plate containing the 4 mL aliquots of the forward and reverse sequencing template (Sequencing Plate). Continue the paused robot program, which should then dispense 11 mL of the appropriate forward and reverse master mixes into the forward and reverse reactions, respectively. Seal the Sequencing Plate with a silicone mat and proceed to the cycle sequencing step. 3. Cycle sequencing: Place the Sequencing Plate in a thermal cycler and run under the following conditions: 96°C for 3 min followed by 25 cycles of 96°C for 15 s, 50°C for 10 s, 60°C for 2 min and then hold at 8°C (see Note 12). Store the Sequencing Plate at 4°C until ready to proceed to next step (see Note 13). 3.5. Purification of Cycle Sequencing Products by EDTA/ Ethanol Precipitation

1. Spin the Sequencing Plate at 40 × g for 30 s. 2. Using the post-PCR robot, return the Sequencing Plate and a vessel containing a combination of 5 mL of 125 mM EDTA and 70 mL of 95% ethanol per reaction equivalent (for this protocol, prepare a volume equivalent to 80 reactions = 6 mL). Run a program that dispenses 75 mL to each cycle sequencing reaction with a mixing step. 3. Seal the plate with a silicone seal and briefly vortex to ensure that it is thoroughly mixed. 4. Incubate plate at room temperature for 15 min (see Note 14). 5. Centrifuge plate at 2,900 × g for 30 min and proceed immediately to step 6 (see Note 15). 6. Carefully remove the silicone seal from the Sequencing Plate and invert onto a bed of paper towels to drain the supernatant. Remove remaining traces of supernatant by centrifugation of the inverted plate at 170 × g for 1 min with a paper towel underneath. 7. Remove the Sequencing Plate from the centrifuge and place on the post-PCR robot, along with a vessel containing 60 mL of 70% ethanol per reaction equivalent (use 4.8 mL for this protocol). Run a program that dispenses 60 mL to each well containing the cycle sequencing reactions with a mixing step. 8. Seal the Sequencing Plate with a silicone seal and centrifuge at 1,600 × g for 15 min and proceed immediately to step 9. 9. Remove the silicone seal and invert onto a bed of paper towels to drain the supernatant. Remove remaining traces of supernatant by centrifugation of the inverted plate at 170 × g for 1 min on a bed of paper towels and then seal the plate. The cycle sequencing products are now purified and can be stored at −20°C prior to analysis (see Note 16).

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3.6. Preparing Samples for Automated Sequence Analysis

1. Remove the plate containing the sequencing pellets from storage at −20°C and place on the post-PCR robot, along with a vessel containing 15 mL of Hi-Di formamide per reaction equivalent (use 1.2 mL for this protocol). Run a program that dispenses 15 mL to each well containing the cycle sequencing reactions (no need for a mixing step). Seal the plate with a silicone mat and proceed immediately to step 2 (see Note 17). 2. Denature the sequencing reactions on a thermal cycler with a heated lid at 94°C for 5 min. On completion of this step, immediately place the plate on ice. 3. The denatured, re-suspended, sequencing reactions are now ready for analysis. Follow the instructions relevant for loading the particular model of DNA sequence analyser in use.

4. Notes 1. Standard Taq DNA polymerases and buffers may not be suitable for templates with a high GC content. Consider using a highfidelity enzyme and/or PCR enhancing agents recommended by the enzyme supplier. 2. Specific primer concentrations may vary for the particular application. 3. For protocols where increased levels of automation are desirable, consideration should be given to the use of filter-based purification plates, which purify PCR products via a vacuum manifold built into the robotic platform. This will omit centrifugation steps but is likely to be more expensive. 4. The primers used in cycle sequencing must undergo HPLC or equivalent purification during manufacture. 5. Ethanol quality and concentration is of paramount importance to the success of the dye terminator removal step. Commercially prepared solutions are convenient and consistent; however, stocks can be prepared from AR-grade Ethanol of known starting concentration. Do not use prepared ethanol solutions more than 4 weeks old. Ethanol precipitation requires the use of centrifugation. Where increased levels of automation are required, consider using filter-based purification plates designed to purify cycle sequencing products via a vacuum mani fold built into the robotic platform. However, this approach will be considerably more expensive than ethanol precipitation. 6. POP-7 Polymer is an acrylamide-based reagent and is therefore a neurotoxin. Observe appropriate safety precautions when handling. 7. This does not allow for a true negative control since genomic DNA is in the PCR master mix. Repeat sequencing of the


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appropriate fragment exhibiting a candidate mutation, along with suitable controls, should be performed before reporting diagnostic results. 8. Thermal cycling parameters will vary according to the particular application. 9. Treat this and all subsequent steps as post-PCR and observe appropriate precautions to minimise the risk of crosscontamination. 10. MicroCLEAN is resistant to mixing, so ensure that the vortexing step is thorough or else incomplete DNA precipitation may result. 11. The BigDye Terminator v3.1 Cycle Sequencing kit is used at reduced concentration. On current generation capillary DNA sequencers, the suggested one-eighth dilution gives good signal strength. It is advisable to store aliquots of the premix to avoid multiple freeze–thaw cycles. Allow the frozen aliquots to thaw fully before use. Keep the BigDye kit on ice while setting up the reaction and do not leave in strong light for extended periods. 12. If necessary, when using the N13 tails, the 50°C primer annealing step can be raised to 55°C in an attempt to overcome any sequencing artefacts introduced by secondary structure of the template DNA. 13. Do not leave the unpurified cycle sequencing reactions for extended periods of time. Ideally, proceed to the purification and precipitation step the same day. 14. The timing of the ethanol precipitation step is important to ensure precipitation of the entire size range of cycle sequencing fragments. 15. Proceed immediately to the next step – if the plate is left to sit in the centrifuge, the unincorporated dye terminators may start to precipitate. 16. The precipitated cycle sequencing products are stable when stored at −20°C and can be left in this state for several days if required. 17. Once the cycle sequencing reactions are resuspended in formamide, they have a limited lifespan. Carry out the rest of the procedure immediately. References 1. Primer 3 (URL: http://primer3.sourceforge. net/webif.php) 2. National Genetics Reference Laboratory (Wessex) Application Note. Standardised Primer Optimisation and Design Specification

(URL: http://www.ngrl.org.uk/Wessex/ downloads_reports.htm) 3. Primers for F8 sequencing (URL: http:// hadb.org.uk/WebPages/Database/Methods/ pcr.html)

Chapter 16 Detection of Factor V Leiden and Prothrombin c.20210G>A Allele by Roche Diagnostics LightCycler® Peter C. Cooper Abstract Venous thrombosis affects one in one thousand people each year, and in many countries, it is a major cause of morbidity and death in hospitalised patients. Factor V Leiden and the prothrombin c.20210G>A transition are relatively common in the Western World, and both increase the risk of venous thrombosis. The author describes the detection of t+++hese two genetic variants on the carousel-based Roche LightCycler®. This simple method has high sensitivity for DNA, making it possible to test blood samples without the need for traditional DNA extraction and purification. Key words: Thrombophilia, Thrombosis, Polymerase Chain Reaction, LightCycler®, Factor V Leiden, Prothrombin c.20210G>A

1. Introduction Each year, at least one person in 1,000 suffers from venous thrombosis (1). Venous thromboembolism (VTE) contributes to the death of more than 50,000 people in the United States each year (1) and is the most common cause of maternal death in pregnancy in the Western world. VTE is extremely rare in childhood, unusual in adults before the age of 50 but relatively common in the elderly. Heritable and acquired factors contribute to an individual’s tendency to thrombosis. In 1993, patients with VTE were described whose plasma failed to respond normally to the anticoagulant effect of activated protein C (APC) that inactivates clotting factors Va and VIIIa. This defect was named APC-resistance (2). In 1994, the major cause of APC-resistance was found when the c.1691G>A or Factor V Leiden mutation (FVL), rs6025, was found in exon 10


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of the factor V gene (F5). This mutation, resulting from CGA in codon 506 being changed to CAA, results in p.Arg506Gln at a site necessary for cleavage of activated factor Va by APC (3). The FVL mutation is believed to have occurred in a single individual, around 21,000–34,000 years ago (4); this mutation may have arisen in the Middle East (5). The relatively high frequency of FVL led to investigation into it having possible survival advantages, and this has been demonstrated, as FVL gives some protection against bleeding and anaemia (6). In the heterozygous state, FVL has a prevalence of around 5% in Europe, with a maximum regional prevalence of 15%; there is a similar high prevalence in the Middle East, and FVL is also found in western Asia and North Africa. FVL increases risk of venous thrombosis in the region by sevenfold, whereas the homozygous state increases risk by about 80-fold; absolute risk of thrombosis depends upon age as well as other risk factors. FVL is not found in subjects native to Japan, Australia, Korea, or in the Native-American Indian populations. In 1996, a second polymorphism was found in subjects with a history of thrombosis. This is a transition in the prothrombin (F2) gene, known as the c.20210G>A allele (7). This base change (rs1799963) is immediately after the last translated codon at the 3¢ end of F2, and whilst not being translated, stabilizes F2 messenger RNA, increasing plasma levels of prothrombin (coagulation factor II) and therefore the potential to form thrombin. The PT c.20210G>A transition is present in around 1–2% of the healthy population, with a distribution similar to that of FVL. Heterozygous PT c.20210G>A confers a twofold to threefold increased tendency to VTE. The relatively high frequency of FVL and PT c.20210G>A may result in them being present in the same individual, or one being present alongside other heritable or acquired thrombotic risk factors. Such inheritance greatly increases risk of venous thrombosis. Testing for thrombophilic defects is a routine procedure. Tests generally include a phenotypic screen for FVL, and if this test is abnormal, it is followed by genetic confirmation. As there is no reliable phenotypic test for PT c.20210G>A, detection is always by genetic analysis. Thrombophilia screening may detect heritable or acquired factors, which predispose a person to thrombosis or pregnancy loss, as described in the British Committee for Standards in Haematology Guidelines for Thrombophilia Testing (8). There are many commercially available assays for FVL and PT c.20210G>A; the author’s first experience of testing for these polymorphisms was using PCR with restriction enzyme digest (9). We now use the Roche Diagnostics LightCycler® version 1, a semi-automated method, which has a reduced hands-on time, requiring only 2.5 h from thawing samples to reporting results. Laboratories carrying out these tests should participate in an external quality assessment scheme (see Note 1).

Detection of Factor V Leiden and Prothrombin c.20210G>A Allele

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In this chapter, the author uses the numbering systems for nucleotide bases and amino acids that are currently used in the majority of published literature, rather than the system recently recommended by the Human Genome Variation society. 1.1. Roche Diagnostics LightCycler ® (Carousel-Based)

The Roche Diagnostics LightCycler® versions 1 and 2 have carousels that hold up to 32 thin-walled 1.5 mm by 30 mm glass capillary tubes, for DNA amplification and analysis. The LightCycler® 1 instrument (currently version 1.5) carries out reactions in 20-ml capillaries and has three detection filters, allowing single PCR analysis and biplex, whereas the version 2 (currently version 2.0) has additional filters, and carousels for 20-ml capillaries for single and biplex analyses, and 100-ml capillaries for multiplex analysis. Fluorescencelabeled probes are used to determine the genotype of test material. The LightCycler® capillaries are loaded with reagents for PCR amplification, one or more fluorescent-labeled probes, and analyte DNA. The capillaries are capped, briefly centrifuged to move the contents to the capillary base, and then loaded into a plastic carousel that is placed on a spindle in the chamber of the LightCycler®. A heater introduces air into the chamber, and temperatures are precisely controlled for PCR and subsequent analysis of genotype. The carousel rotates the capillaries over a lens which transmits monochromatic blue light (maximum emission at 470 nm) from a light emitting diode and collects light from fluorescing probes. After performing a single fluorescence calibration, biplex and multiplex assays can be carried out on LightCycler® 1 and 2, respectively. In addition to detecting genotype by fluorescence resonance energy transfer (FRET), the instruments can determine genotype in assays using the dye Sybr Green 1, which fluoresces when bound to double-stranded DNA. Another means of detection of single nucleotide polymorphisms is through the use of SimpleProbes®, which are oligonucleotides labeled with a fluorophore that emits more light when bound to an amplicon than when in solution or removed from the amplicon by melting, simplifying the assay as only one probe is required. In contrast to the LightCycler® versions 1 and 2, the 480 RealTime PCR System utilises 96 and 384 microplate well formats. PCR block temperature is controlled by means of a Peltier effect heat pump, and amplicon detection is aided by one or more of five excitation filters and six emission filters. This review is limited to the carousel format instruments; further information on the LightCycler 480® can be found on the Roche Diagnostics website (https://www.roche-applied-science.com/sis/rtpcr/index.jsp).

1.2. Detection of Factor V Leiden and PT c.20210G>A by LightCycler ®

FVL detection is an integral part of the thrombophilia screen, and many centres detect FVL negative subjects using the APCresistance test on plasma diluted in factor V depleted plasma; in the absence of heparin, these assays are generally 100% sensitive

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to FVL (see Note 2). An abnormal APC resistance screen is followed up by confirmation of genotype by genetic analysis, and comparison of one result with the other is a valued aid to quality control. Unlike FVL, PT c.20210G>A can only be detected by genetic analysis. The author’s laboratory evaluated Roche kits for FVL and PT c.20210G>A a number of years ago and published an evaluation (10), which demonstrated that the assays have such a high sensitivity to genomic DNA that a simply prepared crude DNA extract can be tested, avoiding the need for costly and time-consuming DNA extraction and purification (see Note 1). The author recommends use of locally derived blood samples as quality control material, as this approach greatly improves evaluation of melting curves and Tm values. The great sensitivity of the LightCycler® to DNA brings with it increased risk of contamination by genomic DNA and amplified product (see Note 3). To reduce risk from contamination with amplicon from previous runs, the kits include dUTP in the amplification mix. To utilise this modification, heat-labile uracil-DNA glycolase can be added into the PCR mixture and incubated for a short time at room temperature; the enzyme is subsequently destroyed during the first DNA denaturisation step of the PCR. Uracil-DNA glycolase destroys contaminating DNA amplicon from previous LightCycler® analyses without affecting the template DNA. However, the author has not found it necessary to make use of this procedure. 1.3. Summary of Procedure

Kits for FVL and Prothrombin G20210A (c.20210G>A) are each supplied in a 32 test format from Roche Diagnostics. The kits comprise detailed instructions and four vials, which are: PCRgrade water, PCR mix, a vial of HybProbe fluorophore probes, and an abnormal (heterozygous) quality control plasmid DNA mix. The kit inserts describe two DNA preparation methodologies that are validated by Roche Diagnostics: a manual kit procedure and an automated method using MagNA Pure instrumentation. The method described below is the author’s modification, which simplifies analysis by removing the need for DNA extraction and purification (10).

1.4. Principle and Summary of the Modified Procedure

Blood is diluted 1:10 in phosphate-buffered saline and heated for 10 min at 95°C, and then centrifuged at 11,000 × g for 60 s, before using 2 mL of the supernatant in the assay. Heating precipitates haemoglobin and other proteins that impede the PCR and also releases DNA from the white blood cells. PCR reagents, probes, and sample (or blank) are added to thin-walled glass capillaries; the capillaries are capped, briefly centrifuged and placed into a carousel in the LightCycler® amplification chamber. The appropriate sequence in the F2 or F5 gene


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is amplified (222 bp for F5 and 165 bp for F2) in around 30 min. After amplification, flourophore-labeled oligonucleotide probes bind to complementary sequences in the amplified DNA. The shorter 3¢ probe is the “mutation probe” that anneals to the area of amplicon containing the nucleotide of interest; this probe is labeled with fluorescein at the 3¢ end which is necessary for detection and to prevent primer extension. The longer 5¢ probe is the “anchor probe” which binds immediately next to the mutation probe and has LightCycler-red (LC-Red ) at the 5¢ end; phosphorylation at the 3¢ end stops amplification. Fluorophores are in very close proximity when bound to the amplicon and when the mixture is suitably illuminated; FRET occurs whereby fluorescein in the mutation probe emits green light which excites LC-Red in the anchor probe to emit red light. To determine genotype, the capillaries are gradually heated and red light emission is monitored. In the absence of the mutation, the mutation probe, which is complementary to the wild-type amplicon, remains attached until a high temperature, whereas the variant amplicon loses its mutation probe at a lower temperature, as it is a poorer match. The fluorescence from LC-red at 640 nm is automatically plotted against temperature, and a melting-curve is shown on the computer screen. Genotype is more easily evaluated when the melting curve is transformed to rate of change in light emission (Fig. 1). The temperature at maximum peak height is called the Tm (melting temperature), which is when half the mutation probe is melted off the amplicon. The Tm is highly reproducible, and is used to determine genotype. The value of the Tm depends upon the degree of match of the probe with the amplicon, the CG content of the amplicon, and to a lesser extent, amplicon length.

2. Materials 1. The FVL kit and Factor II (prothrombin) G20210A kit are supplied in 32 test formats by Roche Diagnostics (Burgess Hill, UK). The kits are delivered on dry ice and then stored at −20°C. Glass capillaries are supplied as a pack of eight boxes of 96-capillary tubes with tops, and the LightCycler® capping tool is purchased separately. 2. The assays for FVL and PT c.20210G>A are designed so that reagent and sample manipulations are identical, and only the instrument set-up and interpretation of results are different. Each kit contains four vials: Vial 1: 78-ml “Yellow Cap” Forward and reverse primers, fluorescent probes, MgCl2, and Brij 35.

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Fig. 1. Results from LightCycler® screen showing fluorescence versus temperature in the top panel and rate of change in fluorescence versus temperature in the lower panel. The dashed line is from a sample that is heterozygous for FVL. In the lower panel, the peaks on the right represent the wild-type allele and peaks on the left represent the mutant allele where the mutant probe melts off at a lower temperature. When a further window is opened, each peak can be observed in isolation, and the Tm values for all samples can be displayed and printed.

Vial 2: 78-ml “Red Cap” Taq polymerase (GMP grade), dATP, dCTP, dGTP, dUTP, dTTP, Tris-buffer, MgCl2 and Brij 35. Vial 3: 50-ml “Purple Cap” is Control Template DNA (wildtype and mutant plasmid DNA). Vial 4: 1,000-ml “Clear Cap” is PCR-Grade water. 3. LightCycler® kit reagent preparation is described in Table 1. 4. Phosphate-buffered saline (Dulbecco A) tablets are purchased as a 100 tablet tub from Oxoid Ltd. (Basingstoke, UK). A tub of tablets is reserved for PCR to ensure that there is no DNA contamination by accidental hand contact. One tablet is dissolved in 100 ml sterile water of a grade that is suitable for PCR and then the solution is separated into at least two tubes which are used in sequence. 5. Automatic pipettes and PCR set-up are isolated from PCR products by being sited in a separate room from the LightCycler®.


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Table 1 Preparation of reagents Preparation of reagents for FVL and PT c.20210G>A Component

Volume per sample

Volume for 32 capillaries

Water (Clear Cap Vial 4)

14 ml

532 ml

Primers/probes (Yellow Cap Vial 1, keep in dark)

2 ml

76 ml

PCR-reagent-mix (Red Cap Vial 2)

2 ml

76 ml

Total volume

18 ml

684 ml

Immediately prior to use: Thaw reagents at room temperature and briefly centrifuge Vials 1 and 2 at 70 g to bring reagents to the base of the tubes. Mix the contents using a gentle pipetting action Prepare working reagent as described above and place in the metal cooling block (reagents are unstable in light) Undiluted reagent can be refrozen, up to five times

Sterile water, buffer, plastic tubes, and filter-tipped pipettes are labeled as for PCR use only, and kept on an area of bench that is reserved for PCR set-up.

3. Method To avoid DNA contamination, gloves must be worn when performing all aspects of the assay, replace gloves if contamination is suspected. 3.1. Blood Samples

3.2. Sample Preparation

Blood samples anticoagulated with citrate or EDTA are suitable for analysis; however, heparin is unsuitable, as it impairs the PCR. We routinely test citrated cells that have had plasma removed for phenotypic tests, as this practice minimises blood sample requirement. Blood samples are stable for many days at room temperature and can be posted to the analytical laboratory without refrigerant. Blood samples can be successfully analysed after at least six freeze–thaw cycles and after storage of blood for at least 1 month at 4°C. It is our normal practice to store blood, awaiting analysis, in the original sample bottle at −20°C; a freeze–thaw cycle increases reliability of the PCR. 1. It is our usual practice to set up an assay for 30 patients, a hete rozygous control (we do not use the Plasmid-heterozygous DNA from Kits) and a PBS blank. The author tests a heterozygous or

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homozygous sample from the previous run as a quality control. Smaller test runs are achievable, as undiluted reagents can be refrozen up to five times. 2. Prepare a worksheet and include a heterozygous sample from a previous run as quality control (QC). The last sample must always be the PBS blank. 3. Thaw samples at room temperature or in an incubator at 37°C and then mix them well. 4. Place the samples in a rack, in test sequence. Do not allow tubes to touch each other. An empty first row in the rack ensures that samples can be moved immediately after sampling to confirm they have been tested. 5. Place 500-ml Eppendorf tubes in a plastic rack, then close and label the top with a sample identity number, 1–32. 6. Just before use, open the Eppendorf tubes and add 225 ml PBS to each. 7. With careful reference to your protocol sheet, to ensure correct sample identification, add 25 ml mixed blood or PBS (for the blank) to the appropriate Eppendorf tube. Immediately after the addition, cap the Eppendorf tube and move the blood sample forward, as confirmation that it has been sampled. 8. Heat the tubes of diluted blood for 10 min (time includes heating up time) at 95°C in a dry heating block, then centrifuge for 1 min at 11,000 × g (brake allowed). The supernatant is now ready for testing for FVL or PT c.20210G>A. 3.3. Preparing Capillaries for the Assay

After thawing samples, select the appropriate assay kit and then confirm that it is for the correct mutation. Thaw the contents at room temperature: thaw the light-sensitive Vial 1 contents whilst held in the metal tub in which it is supplied. After use, undiluted reagents can be refrozen up to five times, but reagents from different kit batches must not be mixed. 1. The metal cooling block with capillary holders is stored at 4°C and transferred to the PCR set-up area immediately before use. Before the first use, Tipp-Ex correction fluid should be wiped over the embossed numbers on the metal block and capillary holders, to make them clearly visible. 2. Each capillary has a number just below the top (1–96). Ensuring that capillaries are in sequence, place the required number of glass capillaries in the numbered metal holders in the cooled capillary holder block. 3. Pipette 18 ml of PCR amplification mixture (see Table 1) into the top of each capillary. 4. Taking care to ensure correct sample order and to avoid contamination, add 2 ml test PBS-blood supernatant, QC material


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or PBS for the blank to the appropriate capillary tubes. If a small amount of denatured blood enters the capillary, this is unlikely to affect the result. To avoid mis-sampling and contamination, immediately after each addition re-cap the Eppendorf tube to show that it has been sampled and cap the capillary, using the capping device. 5. Transfer the capillaries in their holders to an angle-head micro-centrifuge and centrifuge until the RCF reaches 150 × g and then stop by applying the brake; this brings the liquid down into capillaries. 3.4. Operating the LightCycler ®

1. Set up the assay conditions on the LightCycler ® before running for the first time (Tables 2 and 3). Turn on the LightCycler® and the computer. 2. Double-click on the LightCycler® front screen icon to open up the program. 3. Click on “Run.” 4. Before loading the instrument, click on “Start Self Test” when the dialogue box instructing you to do this appears. On completion (around 2 min), click “OK” if the self test has been passed, or take appropriate action if a problem is identified. 5. Taking care to maintain test order, gently drop the capillaries into the holes of the LightCycler® rotor, ensuring that the numeric capillary label is visible. Once all are in place, gently press the capillaries down into the plastic rotor; applying gentle vertical pressure avoids breakage. 6. Check the number on the top end of the capillaries to confirm that they are sequential and that no mix up has occurred. 7. The capillaries are now ready for analysis. 8. Click on “open experiment file.” 9. Open the appropriate analysis file for FVL or PT G2010A (.exp file) as previously set up (see Tables 2 and 3). 10. Place the filled rotor into the LightCycler® chamber, ensuring the rotor slot locates with the small peg in the instrument and then close the lid. 11. Click on “Run.” 12. Enter a suitable file name, for example, fvl05nov2009a (parameter, date, run). 13. Save file (remember to enter the file name on your worksheet). 14. Click on “Clear Sample List” if patient details have previously been entered. This leaves only file details and order of testing on the screen (the author prefers this approach, rather than typing details into the LightCycler® file).

1

45

Denaturation

Amplification

None

Cooling

1

Melting curves

Melting curve 1

Quantification

None

Analysis Cycles mode

PCR stage experiment

40

95 45 80

95 55 72

95

30

0 1:00 0

0 10 5

30

20.00

20.00 20.00 0.10

20.00 20.00 20.00

20.00

Target Temperature temperature Incubation time transition rate (°C) (h:min:s) (°C/s)

Factor V Leiden kit: Settings for LightCycler 1

Table 2 LightCycler preparation for the assay of Factor V Leiden

0

0 0 0

0 0 0

0

Secondary target temperature (°C)

0.0

0.0 0.0 0.0

0.0 0.0 0.0

0.0

0

0 0 0

0 0 0

0

Step size Step delay (°C) (cycles)

Display mode

None

None None CONT

None Single None

None

Acquisition mode

1

F2

248 Cooper

1

45

Denaturation

Amplification

40

None

Cooling

1

Melting curves 95 55 45 40 70

95 55 72

95

30

1:00 30 30 2:00 0

0 10 5

30

20.00

20.00 20.00 20.00 20.00 0.10

20.00 20.00 20.00

20.00

Target Temperature temperature Incubation time transition rate (°C) (h:min:s) (°C/s)

Melting curve 1

Quantification

None

Analysis Cycles mode

PCR stage experiment

Prothrombin G20210A kit (c.20210G>A): Settings for LightCycler 1

Table 3 LightCycler preparation for the assay of prothrombin c.20210G>A

0

0 0 0 0 0

0 0 0

0

Secondary target temperature (°C)

0.0

0.0 0.0 0.0 0.0 0.0

0.0 0.0 0.0

0.0

Step size (°C)

0

0 0 0 0 0

0 0 0

0

Step delay (cycles)

Display mode

None

None None None None CONT

None Single None

None

Acquisition mode

1

F2

Detection of Factor V Leiden and Prothrombin c.20210G>A Allele 249

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15. Enter the number of capillaries to be monitored by adjusting the number in “Select Maximum No. Capillaries.” 16. Click on “Done” (this starts Analysis). 17. The amplification and analysis steps proceed automatically and are completed 40 min after pressing “run.” 18. Return the cooling block to the refrigerator immediately after an assay run, so it is always kept cool and ready for use. 3.5. Results from the LightCycler ®

1. If it is not already open, first open the appropriate file. 2. On the front screen, click on “Data Analysis.” 3. Click on the file for analysis, taking care to select the correct file. 4. Open the file. 5. Change Fluorescence Mode setting to “F2/F1.” 6. Click on “Melting Curve.” 7. The screen now shown is “Step 1: Melting Peaks.” 8. Ensure that the “Calculation Method” is selected as “polynomial” and that the “Digital Filter” is enabled. 9. Click on “Step 2: Peak Areas.” 10. Ensure that the “Weighted Filter” is selected. 11. Using the arrow keys, select the last sample (the blank) and if there is a discernable peak that indicates DNA amplification, there is contamination and the results are invalid; take appropriate action, as described in Note 3. If there is no contamination, document the maximum value on the Y-axis for the blank and state on the worksheet that the blank is “clear.” 12. Select the first test sample. 13. Now, enter the “Number of Peaks” for the sample (0, 1, or 2) which will be one for homozygous wild type or mutant or two for heterozygous genotype. The display now shows the Tm (Tm1 and/or Tm2) for these peaks. 14. Repeat for each of the samples. 15. Now, report the genotype on the worksheet. 16. FVL/PT c.20210G>A Absent – single peak at around 65°C/59°C, respectively. 17. FVL/PT c.20210G>A Homozygous – single peak at around 56°C/49°C, respectively. 18. FVL/PT c.20210G>A Heterozygous – two peaks, representing wild-type and mutant alleles. 19. The wild-type peak is generally slightly lower than the mutant peak, but if a run shows a heterozygous pattern where a peak is unusually large compared to the second peak, then


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contamination may be a cause, and it is recommended to repeat the affected test. 20. If a Tm differs from the in-house heterozygous control by 1°C or more, the test must be repeated, and if the discrepancy is confirmed, there may be an unusual mutation, which should be confirmed and identified by sequencing (see Notes 2 and 4). 21. Print out the results from the LightCycler®: first click on “Report” and then on “Print Summary Report.” 22. Check that the results you have written on the work sheet agree with the results from the printout. 23. Repeat all heterozygous PT c.20210G>A genotypes and check FVL genotypes against phenotype; all homozygous mutant phenotypes are repeated at least once. If there is very poor amplification in a sample, there is a risk that only the mutant DNA will amplify and a patient with a heterozygous genotype may be wrongly labeled as being homozygous for the mutation (see Notes 2 and 5). 24. Enter the results into the patient record computer and tick the worksheet to state that this has been done. 25. Now, look up each patient on the computer to confirm that the result in the computer agrees with the result on the work sheet, and tick the next column to confirm. 26. Gently eject the capillaries onto a layer of tissues using the capillary extractor device and dispose of the used capillaries using an approved method, such as incineration. 27. Close down the computer and turn off the LightCycler®.

4. Notes 1. Validation of the method and quality control: In a comparability study, four hundred samples were tested on the LightCycler® with PBS-blood and the results compared to restriction enzyme digest methodology; there was complete concordance for FVL: wild-type (n = 82), heterozygous (n = 100) and homozygous (n = 18) genotypes; and for prothrombin c.20210G>A: wild type (n = 135), heterozygous (n = 63) and homozygous genotypes (n = 2) (10). The LightCycler® returned the correct genotype in a study of 789 samples that were also tested by NanoChip® and restriction fragment length polymorphism (RFLP) for FVL or PT c.20210G>A mutations (11). In a study using Roche Diagnostics kits on the LightCycler® and extracted DNA, there was 100% concordance between the LightCycler®

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results and RFLP analysis in 110 samples tested for FVL and PT c.20210G>A: 2 samples were homozygous and 38 heterozygous for FVL, 8 samples were heterozygous for PT c.20210G>A and 3 of the 110 were compound heterozygotes (12) . These authors concluded that the LightCycler® methods have shorter hands-on time and are therefore highly economic, and the assays were robust, fast and accurate making them ideally applicable for testing small and large numbers of samples. Utilising a multiplex assay for the LightCycler®, samples comprising 31 wild type, 11 heterozygous, and 5 homozygous samples for FVL were tested, along with 38 wild type, 6 heterozygous, and 3 homozygous for PT c.20210G>A and the results were in full agreement with restriction enzyme analysis (13). In addition to testing abnormal and blank material to validate each assay run, external validation is necessary. In the UK, this is organized by the National External Quality Assessment Scheme for Blood Coagulation UK NEQAS (Coagulation) www.ukneqasbc.org). UK NEQAS (coagulation) currently distributes blood samples to around 95 participant centres. It is common for at least one centre to return an incorrect result, and mistakes occur due to errors in processing and transcription (14). The Haemostasis Committee of the Italian Committee for Standardisation of Laboratory Methods found significant errors in the determination of FVL and PT c.20210G>A genotypes; in a survey, 17.2% of respondents failed to detect the homozygous PT c.20210G>A genotype, and 3.4% of respondents failed to identify homozygous FVL (15). The Australasia Quality Assurance program demo nstrated that over a 5-year period when 133 DNA samples had been tested by up to 39 laboratories, 98.63% of results were correctly reported, but during this time 51% of centres reported at least one incorrect result (16). The poorest results were from a sample homozygous for FVL, where 15% of participants reported the incorrect genotype and, overall, three laboratories were responsible for 46% of all the errors. The carousel-based LightCycler® can simplify analysis by having no requirement for purified DNA. The method format, whereby numbered capillaries are sealed after adding test sample, may reduce random operator error and results are easy to read and interpret. Simplicity reduces risk of error. The only downside of the LightCycler® is that the great sensitivity to DNA makes contamination more of a hazard, but this can be successfully managed by good technique. 2. Interpretation of the FVL results: If a Tm value differs by more than 1°C from the control Tm and the mean of other samples in the run, repeat the assay on that sample, and if the discrepancy is confirmed, a further sample should be requested to confirm


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the finding. If the unusual Tm is again confirmed, then the amplified region of the gene should be sequenced to identify the cause. Where there is a phenotypic FVL screen result, check that it confirms the genetic result, and if there is a discrepancy, repeat both tests. Homozygous genotypes must be checked at least one more time before the result can be released, and a fresh sample should be requested for confirmation. The author, on rare occasions, has seen examples of bone marrow transplants and liver transplants causing discrepancy between plasma phenotype and white blood cell genotype. Antiphospholipid antibodies, high factor VIII levels, and pregnancy may all cause increased APC resistance in the original test without plasma dilution in factor V depleted plasma; however, testing plasma diluted in factor V depleted plasma greatly increases sensitivity and specificity for FVL. Interestingly, the very rare factor V Cambridge variation (c.1091G>C, p.Arg306Thr) may also increase APC resistance in plasma diluted in factor V depleted plasma (17). Arg 306 is a second site for cleavage of factor V by APC; however, APC resistance in patients with this mutation has been questioned (18). The rare silent c.1692A>C polymorphism in the Factor V gene, which is incorrectly identified as FVL by restriction enzyme digest assays, is distinguished from both wild-type Factor V and FVL when tested by LightCycler® (19). The c.1690C>T mutation, which results in the change p.Arg506X and factor V deficiency, has been reported to be misclassified as FVL by assays using restriction enzyme digest but is detected by the LightCycler®, as it has a Tm that is approximately 2°C higher than the 1690C allele (20). These authors state that the variability in Tm for wild-type and mutant FVL alleles is C, c.1689G>A, and c.1696A>G will be classified falsely as FVL by software version 4.05. In the authors’ laboratory, samples with abnormal FVL phenotype are tested by PCR and genotype compared with phenotype, and Tm values are always observed. We have not detected unusual melting temperatures in the FVL assay in over 3,000 samples tested this way. We have, however, found a small number of subjects who are heterozygous for FVL but have a phenotype suggestive of homozygous FVL. This unusual pattern is caused by the FVL gene being expressed, whereas the allelic gene encodes for factor V deficiency; this discrepant pattern has been previously described as pseudohomozygous FVL. 3. Decontamination of materials when contamination is suspected: Dispose of pipette tips and other materials that may

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have been contaminated, then wipe down surfaces and pipettes with DNAZap™ (Applied Biosystems, Warrington, UK). DNAZap™ destroys DNA and PCR products on contact. Wearing gloves, mix equal volumes of DNAZap™ solutions 1 and 2 and use immediately as this mixture is active for less than 1 min. After decontamination, rinse/wipe the apparatus with water to remove all residues. Broken capillaries can be brushed out of the carousel by means of an interdental brush (e.g. TePe Proximal™ Malmo, Sweden) and then decontaminate the carousel with DNAZap™ as described above. The LightCycler® manual describes procedures that are suitable for decontaminating the instrument. 4. Interpretation of the Prothrombin c.20210G>A results: If a Tm value differs by more than 1°C from the expected level, follow the scheme described above for FVL. The most common cause of such a defect is the c.20209C>T polymorphism that is present in around 0.37% of the African American population (21). In addition to the c.20209C>T mutation which can be detected by the LightCycler®, a novel and uncommon c.20221C>T F2 mutation was found in three subjects in a family of Lebanese/Syrian descent (22). The authors found the Tm for the mutant allele to be 54°C, which is the same as for the PT 20209T allele in our laboratory. This demonstrates that whilst the assays for FVL and PT c.20210G>A can detect unusual mutations, an unusual mutation can have the same melting curve characteristics as other genotypes. In our main Caucasian population, we have found less than 10 subjects with an unusual Tm after testing more than 12,000 blood specimens. 5. It is normal for the mutant peak to be somewhat higher than the wild-type peak, but if the discrepancy is large, poor amplification or contamination might be the cause, and the test must be repeated. Homozygous genotypes must always be confirmed by re-testing, and a fresh blood sample should be requested to confirm the homozygous genotype. References 1. Heit JA, Silverstein MD, Mohr DN, Petterson TM, Lohse CM, O’Fallon WM, Melton III LJ (2001) The epidemiology of venous thromboembolism in the community. Thromb Haemost 86, 452–463 2. Dahlbäck B, Carlsson M, Svensson PJ (1993) Familial thrombophilia due to a previously unrecognised mechanism characterised by poor anticoagulant response to activated protein C: prediction of a cofactor to activated protein C. Proc Natl Acad Sci U S A 90, 1004–1008

3. Bertina RM, Koeleman BPC, Koster T, Rosendaal FR, Dirven RJ, de Ronde H, van der Velden PA, Reitsma PH (1994) Mutation in blood coagulation factor V associated with resistance to activated protein C. Nature 369, 64–67 4. Zivelin A, Griffin JH, Xu X, Pabinger I, Samama M, Conard J, Brenner B, Eldor A, Seligsohn U (1997) A single genetic origin for a common Caucasian risk factor for venous thrombosis. Blood 89, 397–402

Detection of Factor V Leiden and Prothrombin c.20210G>A Allele 5. Zivelin A, Mor-Cohen R, Kovalsky V, Kornbrot N, Conard J, Peyvandi F, Kyrle PA, Bertina R, Peyvandi F, Emmerich J, Seligsohn U (2006) Prothrombin 20210G>A is an ancestral prothrombotic mutation that occurred in whites approximately 24 000 years ago. Blood 107, 4666–4668 6. Lindqvist PG, Zoller B, Dahlback B (2001) Improved hemoglobin status and reduced menstrual blood loss among female carriers of factor V Leiden – an evolutionary advantage? Thromb Haemost 86, 1122–1123 7. Poort SR, Rosendaal FR, Reitsma PH, Bertina RM (1996) A common variation in the 3¢-untranslated region of the prothrombin gene is associated with elevated plasma prothrombin levels and an increase in venous thrombosis. Blood 88, 3698–3703 8. Walker ID, Greaves M, Preston FE: Guideline: investigation and management of heritable thrombophilia (2001) Br J Haematol 114, 512–528 9. Beauchamp NJ, Daly ME, Cooper PC, Preston FE, Peake IR (1994) Rapid two-stage PCR for detecting factor V G1691A mutation. Lancet 334, 694–695 10. Cooper PC, Cooper SM, Smith JM, Kitchen S, Makris M (2003) Evaluation of the Roche LightCycler: a simple and rapid method for direct detection of factor V Leiden and prothrombin G2010A genotypes from blood samples without the need for DNA extraction. Blood Coagul Fibrinolysis 14, 499–503 11. Schrijver I, Lay MJ, Zehnder JL (2003) Diagnostic single nucleotide polymorphism analysis of factor V Leiden and prothrombin 20210G>A. A comparison of the Nanogen Electronic Microarray with restriction enzyme digestion and the Roche LightCycler. Am J Clin Pathol 119, 490–496 12. Nauck M, März W, Wieland H (2000) Rapid homogenous detection of factor V Leiden and prothrombin mutations on the LightCycler. Clin Biochem 33, 213–216 13. van den Bergh FAJTM, van Oeveren-Dybicz AM, Bon MAM (2000) Rapid single-tube genotyping of the Factor V Leiden and prothrombin mutations by real-time PCR using dual-color detection. Clin Chem 46, 1191–1195

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14. Preston FE, Kitchen S, Jennings I, Woods TAL (1999) A UK national quality assessment scheme (UK NEQAS) for molecular genetic testing for the diagnosis of familial thrombophilia. Thromb Haemost 82, 1556–1557 15. Tripodi A, Peyvandi F, Chantarangkul V, Menegatti M, Mannucci PM (2002) Relatively poor performance of clinical laboratories for DNA analyses in the detection of two thrombophilic mutations – a cause for concern. Thromb Haemost 88, 690–691 16. Hertzberg M, Neville S, Favaloro E, McDonald D (2005) External quality assurance of DNA testing for thrombophilia mutations. Am J Clin Pathol 123, 189–193 17. Williamson D, Brown K, Luddington R, Baglin C, Baglin T (1998) Factor V Cambridge: a new mutation (Arg306Thr) associated with resistance to activated protein C. Blood 91, 1140–1144 18. ten Cate AJ, van de Hoek YT, Reitsma PH, ten Cate H, Smits P (2001) Mutation screening for thrombophilia: two cases with factor V Cambridge without activated protein C resistance. Thromb Haemost 87, 919–920 19. Parks SB, Popovich BW, Press RD (2000) Real-time polymerase chain reaction with fluorescent hybridization probes for the detection of prevalent mutations causing common thrombophilic and iron overload phenotypes. Am J Clin Pathol 115, 439–447 20. Mahadevan MS, Benson PV (2005) Factor V null mutation affecting the Roche LightCycler factor V Leiden assay. Clin Chem 51, 1533–1535 21. Itakura H, Telen MJ, Hoppe CC, White DAE, Zehnder JL (2005) Characterization of a novel prothrombin variant, prothrombin C20209T, as a modifier of thrombotic risk among African-Americans. Thromb Haemost 3, 2357–2359 22. Wylenzek M, Geison C, Stapenhorst L, Wielckens K, Klingler KR (2001) A novel point mutation in the 3¢ region of the prothrombin gene at position 20221 in a Lebanese/Syrian family. Thromb Haemost 85, 943–944

Chapter 17 RT-PCR for the Detection of Translocations in Bone and Soft Tissue Tumours in Formalin-Fixed Paraffin-Embedded Tissues Ann Williams and D. Chas Mangham Abstract This chapter outlines the methodology for the detection of mesenchymal-tumour-specific translocations in formalin-fixed paraffin-embedded tissue (FFPET) using the reverse transcriptase-polymerase chain reaction (RT-PCR). It includes the design of appropriate primer pairs and the necessary pretreatment of the FFPET sections to give the maximum yield of analyzable RNA, in terms of both quantity and quality. Key words: Formalin-fixed paraffin-embedded tissue (FFPET), RT-PCR, Sarcoma-specific transl ocations, Acid decalcification

1. Introduction The discovery of chromosomal translocations in tumours was first made by classical karyotyping of leukaemia cells; the Philadelphia chromosome in chronic myeloid leukaemia was the first to be discovered in 1960 (1). The first sarcoma-associated balanced chromosomal translocation was discovered in Ewing’s sarcoma in 1984 (2). Since then, numerous distinctive histological types of sarcoma have been found to harbour characteristic chromosomal translocations. Table 1 gives a list of currently known sarcomas in which associated chromosomal translocations have been identified. The identification of translocations helps to accurately diagnose sarcoma subtypes when performed in conjunction with histological examination and immunohistochemical phenotyping. Accurate sarcoma subtype diagnosis provides important prognostic information and increasingly directs treatment protocols. Bimal D.M. Theophilus and Ralph Rapley (eds.), PCR Mutation Detection Protocols, Methods in Molecular Biology, vol. 688, DOI 10.1007/978-1-60761-947-5_17, © Springer Science+Business Media, LLC 2011

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Table 1 Known translocations in bone and soft-tissue neoplasms Tumour

Translocation

Fusion product

Incidence (%)

Alveolar rhabdomyosarcoma

t(2;13)(q35;q14) t(1;13)(p36;q14) t(X;2)(q13;q35) t(2;2)(q35p23)

PAX3/FOXO1(FKHR) PAX7/FOXO1(FKHR) PAX3/AFX PAX3/NCOA1

65 15

Alveolar soft part sarcoma

t(X;17)(p11;q25)

ASPL/TFE3

>95

Aneurysmal bone cyst

t(16;17)(q22;p13) t(1;17)(p34.1–34.3;p13) t(3;17)(q21;p13) t(9;17)(q22;p11-12) t(17;17)(p13;q12)

CDH11/USP6 TRAP150/USP6 ZNF9/USP6 OMD/USP6 COL1A1/USP6

Angiomatoid fibrous histiocytoma

t(2;22)(q33;q12) t(12;22)(q13;q12) t(12;16)(q13;p11)

EWS/CREB1 EWS/ATF1 FUS/ATF1

Clear cell sarcoma

t(12;22)(q13;q12) t(2;22)(q33;q12)

EWS/ATF1 EWS/CREB1

>90

Dermatofibrosarcoma protuberans/giant cell fibroblastoma

t(17;22)(q21;q13)

COL1A1/PDGFB

>90

Desmoplastic small round cell tumour

t(11;22)(p13;q12)

EWS/WT1

>95

Ewing’s sarcoma/ PNET

t(11;22)(q24;q12) t(21;22)(q22;q12) t(7;22)(p22;q12) t(17;22)(q12;q12) t(2;22)(q33;q12) 22q12 rearrangement t(2;22)(q31;q12) t(6;22)(p21;q12) t(16;21)(p11;q22) t(2;16)(q35;p11)

EWS/FLI1 EWS/ERG EWS/ETV1 EWS/ETV4(E1AF) EWS/FEV EWS/PATZ1(ZSG) EWS/SP3 EWS/POU5F1 FUS/ERG FUS/FEV

90 5