Methods in molecular biology vol 1537 oral biology molecular techniques and applications 2nd edition

cannOn • Department of Oral Sciences, University of Otago School of Dentistry, Dunedin, New Zealand; Faculty of Dentistry, Sir John Walsh Research Institute, University of Otago School

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

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ISSN 1064-3745 ISSN 1940-6029 (electronic)

Methods in Molecular Biology

ISBN 978-1-4939-6683-7 ISBN 978-1-4939-6685-1 (eBook)

DOI 10.1007/978-1-4939-6685-1

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be true and accurate at the date of publication Neither the publisher nor the authors or the editors give a warranty, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made Cover illustration: Example of a bead experiment combined with in situ hybridization (ISH) analysis to study gene

expression in embryonic tissue explants The image shows the effects of BMP2 beads on ld1 gene expression in explants

of calvarial mesenchyme Photograph provided by D Rice and K Närhi The bead and ISH experiments are described

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Dunedin, New Zealand

It is widely accepted that “evidence-based dentistry” is fundamental to clinical practice and that well-controlled randomized clinical trials followed by systematic reviews and meta- analyses provide much of this evidence base However, it is still the basic biological and physical sciences that underpin advances in dentistry and form the basis for subsequent clinical trials It is equally true that the treatment of any disease should be based on an understanding of the etiology and pathogenesis of that disease, and in this context, the future of dentistry lies very much in continued research in the basic biological sciences

This second edition of Oral Biology: Molecular Techniques and Applications continues

the approach taken in the first edition and has not attempted to cover all aspects of oral biology, but rather to present a selection of cellular and molecular techniques that can be adapted to cover a range of applications and diseases The first part on saliva, for example, has been updated and expanded to include proteomic analyses by mass spectrometry and NMR-based metabolomics that can be used not only in the study of saliva but also in assess-ing other oral fluids such as gingival fluid Clearly, saliva is unique to the oral cavity but so too is gingival fluid which, in essence, is the fluid medium of the gingiva and gingival sulcus, and thus is the fluid environment where interactions between the plaque biofilm and the host take place Hence, techniques for its collection and analysis have now been included.Although it is 6 years since publication of the first edition of this book, many of the techniques described are still in widespread use and so have been retained, albeit updated,

in this second edition In the part on molecular biosciences, for example, chapters on ing of oral microbial communities, quantitative real-time PCR, and adhesion of yeast and bacteria to oral surfaces have all been retained but substantially updated

profil-Epigenetics is now a major theme in biology and is providing great insight into how we interact with our environment As DNA methylation features heavily in epigenetic studies, new chapters on tools and strategies that facilitate the analysis of genome-wide or gene- specific DNA methylation patterns have been included

As in the first edition, the last part of this second edition deals with a range of approaches that enable the behavior of cells and tissues in both health and disease to be analyzed at the molecular level The future of dentistry and of the profession lies in research, and it is antici-

pated that this second edition of Oral Biology: Molecular Techniques and Applications will

continue to be a useful resource for oral biologists at all levels, be they students, early career

or experienced veterans, and that it provides a ready reference enabling new techniques and approaches to be used in answering a range of specific scientific questions that will underpin

a deeper understanding and treatment of oral diseases

Preface

Contents

Preface v Contributors xi

Part I SalIva and Other Oral FluIdS

1 Salivary Diagnostics Using Purified Nucleic Acids 3

Paul D Slowey

2 RNA Sequencing Analysis of Salivary Extracellular RNA 17

Blanca Majem, Feng Li, Jie Sun, and David T.W Wong

3 Qualitative and Quantitative Proteome Analysis of Oral Fluids

in Health and Periodontal Disease by Mass Spectrometry 37

Erdjan Salih

4 Antioxidant Micronutrients and Oxidative Stress Biomarkers 61

Iain L.C Chapple, Helen R Griffiths, Mike R Milward,

Martin R Ling, and Melissa M Grant

5 NMR-Based Metabolomics of Oral Biofluids 79

Horst Joachim Schirra and Pauline J Ford

6 Gene Therapy of Salivary Diseases 107

Bruce J Baum, Sandra Afione, John A Chiorini, Ana P Cotrim,

Corinne M Goldsmith, and Changyu Zheng

Part II MOlecular BIOScIenceS

7 The Oral Microbiota in Health and Disease: An Overview

of Molecular Findings 127

José F Siqueira Jr and Isabela N Rôças

8 Microbial Community Profiling Using Terminal Restriction

Fragment Length Polymorphism (T-RFLP) and Denaturing Gradient

Gel Electrophoresis (DGGE) 139

José F Siqueira Jr., Mitsuo Sakamoto, and Alexandre S Rosado

9 Analysis of 16S rRNA Gene Amplicon Sequences

Using the QIIME Software Package 153

Blair Lawley and Gerald W Tannock

10 Adhesion of Yeast and Bacteria to Oral Surfaces 165

Richard D Cannon, Karl M Lyons, Kenneth Chong,

Kathryn Newsham-West, Kyoko Niimi, and Ann R Holmes

11 Quantitative Analysis of Periodontal Pathogens Using Real- Time

Polymerase Chain Reaction (PCR) 191

Mª José Marin, Elena Figuero, David Herrera, and Mariano Sanz

12 Methods to Study Antagonistic Activities Among Oral Bacteria 203

Fengxia Qi and Jens Kreth

13 Natural Transformation of Oral Streptococci by Use of Synthetic

Pheromones 219

Gabriela Salvadori, Roger Junges, Rabia Khan, Heidi A Åmdal,

Donald A Morrison, and Fernanda C Petersen

14 Markerless Genome Editing in Competent Streptococci 233

Roger Junges, Rabia Khan, Yanina Tovpeko, Heidi A Åmdal,

Fernanda C Petersen, and Donald A Morrison

15 Tools and Strategies for Analysis of Genome-Wide and Gene-Specific

DNA Methylation Patterns 249

Aniruddha Chatterjee, Euan J Rodger, Ian M Morison, Michael R Eccles,

and Peter A Stockwell

16 Generating Multiple Base-Resolution DNA Methylomes Using Reduced

Representation Bisulfite Sequencing 279

Aniruddha Chatterjee, Euan J Rodger, Peter A Stockwell,

Gwenn Le Mée, and Ian M Morison

17 A Protocol for the Determination of the Methylation Status

of Gingival Tissue DNA at Specific CpG Islands 299

Trudy J Milne

18 Genome-Wide Analysis of Periodontal and Peri-Implant Cells and Tissues 307

Moritz Kebschull, Claudia Hülsmann, Per Hoffmann,

and Panos N Papapanou

19 Differential Expression and Functional Analysis

of High- Throughput -Omics Data Using Open Source Tools 327

Moritz Kebschull, Melanie Julia Fittler, Ryan T Demmer,

and Panos N Papapanou

20 Exploring Genome-Wide Expression Profiles Using Machine

Learning Techniques 347

Moritz Kebschull and Panos N Papapanou

Part III cellS and tISSueS

21 Embryonic Explant Culture: Studying Effects of Regulatory

Molecules on Gene Expression in Craniofacial Tissues 367

Katja Närhi

22 Oral Epithelial Cell Culture Model for Studying the Pathogenesis

of Chronic Inflammatory Disease 381

Mike R Milward, Martin R Ling, Melissa M Grant,

and Iain L.C Chapple

23 Fabrication and Characterization of Decellularized Periodontal Ligament

Cell Sheet Constructs 403

Amro Farag, Cédryck Vaquette, Dietmar W Hutmacher, P Mark Bartold,

and Saso Ivanovski

24 A Method to Isolate, Purify, and Characterize Human Periodontal

Ligament Stem Cells 413

Krzysztof Mrozik, Stan Gronthos, Songtao Shi, and P Mark Bartold

25 Constructing Tissue Microarrays: Protocols and Methods Considering

Potential Advantages and Disadvantages for Downstream Use 429

Lynne Bingle, Felipe P Fonseca, and Paula M Farthing

26 Growing Adipose-Derived Stem Cells Under Serum-Free Conditions 439

Diogo Godoy Zanicotti and Dawn E Coates

27 Quantitative Real-Time Gene Profiling of Human Alveolar Osteoblasts 447

Dawn E Coates, Sobia Zafar, and Trudy J Milne

28 Proteomic Analysis of Dental Tissue Microsamples 461

Jonathan E Mangum, Jew C Kon, and Michael J Hubbard

29 Characterization, Quantification, and Visualization of Neutrophil

Extracellular Traps 481

Phillipa C White, Ilaria J Chicca, Martin R Ling, Helen J Wright,

Paul R Cooper, Mike R Milward, and Iain L.C Chapple

Index 499

Sandra aFIOne • Molecular Physiology and Therapeutics Branch, National Institute

of Dental and Craniofacial Research, National Institutes of Health (NIH), Bethesda,

MD, USA

heIdI a ÅMdal • Department of Oral Biology, Faculty of Dentistry, University of Oslo,

Oslo, Norway

P Mark BartOld • Colgate Australian Clinical Dental Research Centre, Dental School,

University of Adelaide, Adelaide, Australia

Bruce J BauM • Molecular Physiology and Therapeutics Branch, National Institute

of Dental and Craniofacial Research, National Institutes of Health (NIH), Bethesda,

MD, USA; Molecular Physiology and Therapeutics Branch, National Institute of Dental and Craniofacial Research, National Institutes of Health (NIH), Bethesda, MD, USA

lynne BIngle • Academic Unit of Oral and Maxillofacial Pathology, School of Clinical

Dentistry, University of Sheffield, Sheffield, UK

rIchard d cannOn • Department of Oral Sciences, University of Otago School of

Dentistry, Dunedin, New Zealand; Faculty of Dentistry, Sir John Walsh Research Institute, University of Otago School of Dentistry, Dunedin, New Zealand

IaIn l.c chaPPle • School of Dentistry, Institute of Clinical Sciences, College of Medical

and Dental Sciences, University of Birmingham, Birmingham, UK

anIruddha chatterJee • Department of Pathology, Dunedin School of Medicine,

University of Otago, Dunedin, New Zealand; Maurice Wilkins Centre for Molecular Biodiscovery, Auckland, New Zealand

IlarIa J chIcca • Institute of Clinical Sciences, College of Medical and Dental Sciences,

The School of Dentistry, University of Birmingham, Birmingham, UK

JOhn a chIOrInI • Molecular Physiology and Therapeutics Branch, National Institute

of Dental and Craniofacial Research, National Institutes of Health (NIH), Bethesda,

MD, USA

kenneth chOng • Department of Oral Sciences, University of Otago School of Dentistry,

Dunedin, New Zealand

dawn e cOateS • Faculty of Dentistry, Sir John Walsh Research Institute, University

of Otago, Dunedin, New Zealand

Paul r cOOPer • Institute of Clinical Sciences, College of Medical and Dental Sciences,

The School of Dentistry, University of Birmingham, Birmingham, UK

ana P cOtrIM • Molecular Physiology and Therapeutics Branch, National Institute

of Dental and Craniofacial Research, National Institutes of Health (NIH), Bethesda,

MD, USA

ryan t deMMer • Department of Epidemiology, Columbia University Mailman School

of Public Health, New York, NY, USA

MIchael r eccleS • Department of Pathology, Dunedin School of Medicine, University

of Otago, Dunedin, New Zealand; Maurice Wilkins Centre for Molecular Biodiscovery, Auckland, New Zealand

aMrO Farag • School of Dentistry and Oral Health, Regenerative Medicine Center,

Menzies Health Institute Queensland, Gold Coast, QLD, Australia

Contributors

Paula M FarthIng • Academic Unit of Oral and Maxillofacial Pathology, School

of Clinical Dentistry, University of Sheffield, Sheffield, UK

elena FIguerO • Oral Research Laboratory, Faculty of Odontology, University

Complutense, Madrid, Spain; Etiology and Therapy of Periodontal Diseases (ETEP) Research Group, University Complutense, Madrid, Spain; Department of Periodontology, Faculty of Dentistry, University Complutense of Madrid, Madrid, Spain

MelanIe JulIa FIttler • Department of Periodontology, Operative and Preventive

Dentistry, University of Bonn, Bonn, Germany

FelIPe P FOnSeca • Department of Oral Diagnosis, Faculty of Dentistry of Piracicaba,

FOP, UNICAMP, Piracicaba, São Paolo, Brazil

PaulIne J FOrd • School of Dentistry, Oral Health Centre, The University of Queensland,

Herston, QLD, Australia

cOrInne M gOldSMIth • Molecular Physiology and Therapeutics Branch, National

Institute of Dental and Craniofacial Research, National Institutes of Health (NIH), Bethesda, MD, USA

MelISSa M grant • School of Dentistry, Institute of Clinical Sciences, College of Medical

and Dental Sciences, University of Birmingham, Birmingham, UK

helen r grIFFIthS • School of Dentistry, Institute of Clinical Sciences, College of Medical

and Dental Sciences, University of Birmingham, Birmingham, UK

Stan grOnthOS • Mesenchymal Stem Cell Group, Adelaide Medical School, Faculty of

Health Sciences, University of Adelaide, Adelaide, SA, Australia

davId herrera • Etiology and Therapy of Periodontal Diseases (ETEP) Research Group,

University Complutense, Madrid, Spain; Department of Periodontology, Faculty of Dentistry, University Complutense of Madrid, Madrid, Spain

Per hOFFMann • Department of Genomics, Institute of Human Genetics, University of

Bonn, Bonn, Germany; Human Genomics Research Group, Department of Biomedicine, University of Basel, Basel, Switzerland

ann r hOlMeS • Department of Oral Sciences, University of Otago School of Dentistry,

Dunedin, New Zealand; Faculty of Dentistry, Sir John Walsh Research Institute,

University of Otago School of Dentistry, Dunedin, New Zealand

MIchael J huBBard • Department of Pharmacology and Therapeutics, University of

Melbourne, Melbourne, VIC, Australia; Department of Pediatrics, Royal Children’s Hospital, University of Melbourne, Melbourne, VIC, Australia

claudIa hülSMann • Department of Periodontology, Operative and Preventive Dentistry,

Faculty of Medicine, University of Bonn, Bonn, Germany

dIetMar w hutMacher • Queensland University of Technology, Brisbane, QLD,

Australia

SaSO IvanOvSkI • School of Dentistry and Oral Health, Regenerative Medicine Center,

Menzies Health Institute Queensland, Gold Coast, QLD, Australia; Menzies Health Institute Queensland, Griffith University, Gold Coast, QLD, Australia

rOger JungeS • Department of Oral Biology, Faculty of Dentistry, University of Oslo, Oslo,

Norway

MOrItz keBSchull • Department of Periodontology, Operative and Preventive Dentistry,

Faculty of Medicine, University of Bonn, Bonn, Germany; Division of Periodontics, Section of Oral, Diagnostic and Rehabilitation Sciences, Columbia University College of Dental Medicine, New York, NY, USA

raBIa khan • Department of Oral Biology, Faculty of Dentistry, University of Oslo, Oslo,

Norway

Jew c kOn • Department of Pharmacology and Therapeutics, University of Melbourne,

Melbourne, VIC, Australia; Department of Pediatrics, Royal Childern’s Hospital, University of Melbourne, Melbourne, VIC, Australia

JenS kreth • Oregon Health and Science University, Portland, OR, USA

BlaIr lawley • Department of Microbiology and Immunology, University of Otago,

Dunedin, New Zealand

Feng lI • Division of Oral Biology and Oral Medicine, School of Dentistry, University of

California Los Angeles (UCLA), Los Angeles, CA, USA

MartIn r lIng • School of Dentistry, Institute of Clinical Sciences, College of Medical

and Dental Sciences, University of Birmingham, Birmingham, UK

karl M lyOnS • Faculty of Dentistry, Sir John Walsh Research Institute, University of

Otago School of Dentistry, Dunedin, New Zealand; Department of Oral Rehabilitation, University of Otago School of Dentistry, Dunedin, New Zealand

Blanca MaJeM • Biomedical Research Unit in Gynecology, Vall Hebron Research Institute

(VHIR) and University Hospital, University Autonoma of Barcelona (UAB),

Barcelona, Spain

JOnathan e ManguM • Department of Pharmacology and Therapeutics, University of

Melbourne, Melbourne, VIC, Australia

Mª JOSé MarIn • Oral Research Laboratory, Faculty of Odontology, University

Complutense, Madrid, Spain

gwenn le Mée • Department of Pathology, Dunedin School of Medicine, University of

Otago, Dunedin, New Zealand

trudy J MIlne • Faculty of Dentistry, Sir John Walsh Research Institute, University of

Otago, Dunedin, New Zealand

MIke r MIlward • School of Dentistry, Institute of Clinical Sciences, College of Medical

and Dental Sciences, University of Birmingham, Birmingham, UK

Ian M MOrISOn • Department of Pathology, Dunedin School of Medicine, University of

Otago, Dunedin, New Zealand

dOnald a MOrrISOn • Department of Biological Sciences, College of Liberal Arts and

Sciences, University of Illinois at Chicago, Chicago, IL, USA

krzySztOF MrOzIk • Colgate Australian Dental Research Centre, Dental School,

University of Adelaide, Adelaide, SA, Australia

katJa närhI • Institute for Molecular Medicine Finland, University of Helsinki, Helsinki,

Finland

kathryn newShaM-weSt • Faculty of Dentistry, Sir John Walsh Research Institute,

University of Otago School of Dentistry, Dunedin, New Zealand; Department of Oral Rehabilitation, University of Otago School of Dentistry, Dunedin, New Zealand

kyOkO nIIMI • Department of Oral Sciences, University of Otago School of Dentistry,

Dunedin, New Zealand

PanOS n PaPaPanOu • Division of Periodontics, Section of Oral, Diagnostic and

Rehabilitation Sciences, Columbia University College of Dental Medicine, New York, NY, USA

Fernanda c PeterSen • Department of Oral Biology, Faculty of Dentistry, University of

Oslo, Oslo, Norway

FengxIa QI • University of Oklahoma Health Sciences Center BRC364, Oklahoma City,

OK, USA

ISaBela n rôçaS • Department of Endodontics and Molecular Microbiology, Estácio de Sá

University, Rio de Janeiro, RJ, Brazil

euan J rOdger • Department of Pathology, Dunedin School of Medicine, University

of Otago, Dunedin, New Zealand

alexandre S rOSadO • Institute of Microbiology Prof Paulo de Góes, Federal University

of Rio de Janeiro, Rio de Janeiro, Brazil

MItSuO SakaMOtO • Microbe Division/Japan Collection of Microorganisms, RIKEN

BioResource Center, Wako, Saitama, Japan

erdJan SalIh • Department of Periodontology, Henry M Goldman School of Dental

Medicine, Boston University, Boston, MA, USA

gaBrIela SalvadOrI • Department of Oral Biology, Faculty of Dentistry, University of Oslo,

Oslo, Norway

MarIanO Sanz • Etiology and Therapy of Periodontal Diseases (ETEP) Research Group,

University Complutense, Madrid, Spain; Department of Periodontology, Faculty of Dentistry, University Complutense of Madrid, Madrid, Spain

hOrSt JOachIM SchIrra • Centre for Advanced Imaging, The University of Queensland,

Brisbane, QLD, Australia

SOngtaO ShI • Department of Anatomy and Cell BiologySchool of Dental Medicine,

University of Pennsylvania, Philadelphia, PA, USA

JOSé F SIQueIra Jr • Department of Endodontics and Molecular Microbiology, Estácio de

Sá University, Rio de Janeiro, RJ, Brazil; Faculty of Dentistry, Estácio de Sá University, Rio de Janeiro, Brazil

Paul d SlOwey • Oasis Diagnostics ® Corporation, Vancouver, WA, USA

Peter a StOckwell • Department of Biochemistry, University of Otago, Dunedin,

New Zealand

JIe Sun • Medical School of Shenzhen University, Shenzhen, Guangdong, China

gerald w tannOck • Department of Microbiology and Immunology, University of Otago,

Dunedin, New Zealand

yanIna tOvPekO • Department of Biological Sciences, College of Liberal Arts and Sciences,

University of Illinois at Chicago, Chicago, IL, USA

cédryck vaQuette • Queensland University of Technology, Brisbane, QLD, Australia

PhIllIPa c whIte • Institute of Clinical Sciences, College of Medical and Dental Sciences,

The School of Dentistry, University of Birmingham, Birmingham, UK

davId t.w wOng • Division of Oral Biology and Oral Medicine, School of Dentistry,

University of California Los Angeles (UCLA), Los Angeles, CA, USA; Johnson

Comprehensive Cancer Center, University of California Los Angeles (UCLA), Los Angeles, CA, USA; Molecular Biology Institute, University of California Los Angeles (UCLA), Los Angeles, CA, USA; Head & Neck Surgery/Otolaryngology, Henry Samuel School of Engineering and Applied Science, University of California Los Angeles

(UCLA), Los Angeles, CA, USA

helen J wrIght • Institute of Clinical Sciences, College of Medical and Dental Sciences,

The School of Dentistry, University of Birmingham, Birmingham, UK

SOBIa zaFar • Faculty of Dentistry, Sir John Walsh Research Institute, University of Otago,

Dunedin, New Zealand

dIOgO gOdOy zanIcOttI • Faculty of Dentistry, Sir John Walsh Research Institute,

University of Otago, Dunedin, New Zealand

changyu zheng • Molecular Physiology and Therapeutics Branch, National Institute of

Dental and Craniofacial Research, National Institutes of Health (NIH), Bethesda, MD, USA

Part I Saliva and Other Oral Fluids

Gregory J Seymour et al (eds.), Oral Biology: Molecular Techniques and Applications, Methods in Molecular Biology, vol 1537,

DOI 10.1007/978-1-4939-6685-1_1, © Springer Science+Business Media LLC 2017

diagnos-Key words Saliva, RNA, DNA, Nucleic acids, Stabilization, Exosomes

The number of applications for saliva is growing exponentially

as evidenced by the increasing number of available tools for saliva acquisition and subsequent testing either immediately at the point

of care or under controlled laboratory conditions

Saliva is now used in tests for adverse responses to multiple therapeutics, genomics for cystic fibrosis, fragile X syndrome in autism, disorders of the salivary glands, cancers (including breast, head and neck, and oral cancers), abused drug testing in the work-place and other environments, as well as certain systemic diseases including HIV, hepatitis C, and Sjögren’s syndrome

The success of any test, whether for research or diagnostic purposes, relies on the successful harvesting of the specimen from

a subject in a standardized, repeatable fashion and careful handling

of the sample throughout the collection and downstream testing process This rule applies to all specimen types, but care should

especially be taken with respect to processing and stabilizing saliva samples to ensure optimum results.

The following text describes some of the newer tools and procedures for collection, stabilization, and storage of oral fluid matrices that aid in the successful use of saliva as a test specimen This chapter focuses particularly on nucleic acid components for downstream molecular diagnostic (MDx) testing, since this is probably the area where saliva is likely to have the greatest impact

in improving healthcare for the general population For more detailed information on current salivary diagnostics and available tools, the reader is referred to several review articles on the subject [1–5]

Dr Lawrence Tabak (Deputy Director of the NIH and former head of the National Institute for Dental and Craniofacial Research, NIDCR) characterized saliva as a “mirror of the body” and is therefore reflective of disease and disease processes going on in the human body This precious biofluid contains many of the biomark-ers that are indicative of disease and maladies affecting human beings, so saliva is the ideal sample matrix for large-scale epidemio-logical studies, population screening, and diagnosis of multiple diseases and conditions

Saliva is cost-effective, noninvasive, easy to transport, amenable

to simple disposal, and highly attractive in certain cultures (and religions), which find the use of blood an unacceptable option More importantly, saliva contains many of the indicators of disease found in blood, urine, and tissue samples

Typically, levels of biomarkers in saliva are 10–1500 times lower than in blood, but with the advent of newer, more sensitive detection technologies, the analysis of salivary biomarkers has become a much more attractive option When patient preference

to eliminate the use of needles is considered as an additive factor, the “compelling story” for saliva grows significantly stronger These are some of the major reasons that there has been an “explo-sion” in research and development in salivary diagnostics, in the last few years, resulting in the development of a plethora of tools and tests using this unique bodily fluid

A series of technological developments, which have also tributed to the growing importance of saliva as a diagnostic medium, include several high-throughput technologies such as next-generation sequencing, proteomics, mass spectrometry, genome wide association studies (GWAS), and genotyping, which allow large numbers of samples to be tested in a short time Saliva has already been shown to be a readily adaptable specimen for use

con-in these high-impact technologies

Saliva is now in routine use for the diagnosis of HIV in the privacy of one’s home [6 7] and for the detection of multiple hor-mones as part of a “general wellness” program, sold direct to the consumer [8–10] Saliva has also been used to detect drugs of abuse [11] and in certain situations has been shown to be a

preferable biofluid to urine, which is currently the method of choice This is particularly true in the case of marijuana, when test-ing for “impairment” and whether a particular individual is fit to drive a vehicle or perform dangerous tasks.

Multiple diseases have also been detected using saliva, including caries risk [12–14]; periodontitis [15]; oral [16], breast [17–22], and head and neck cancers [23]; and salivary gland disorders [24] Point of care tests are now also in development looking at viruses, bacteria [25], and difficult to measure hormones using saliva [26]

Perhaps the area where saliva has gained the most traction is for the collection of nucleic acids (DNA and RNA) The noninvasive nature of saliva means that samples of DNA or RNA can be collected

at a remote site, sometimes without professional input, and ported to a laboratory where on-site testing is performed and the results reported back to the physician, who in turn can provide rapid feedback to the subject or patient The elimination of the phleboto-mist to collect a sample is the key driver in this instance

trans-There are a number of tools available for genomic DNA collection from saliva and more are currently in development These are based upon the collection of whole saliva, or in some cases buccal epithelial cells, harvested by a rinse solution or mouthwash system

Since the discovery of RNA in saliva [16], there has been a rapid uptake in transcriptomic analysis using saliva specimens A group

of RNAs termed “core” RNAs have been found to be present in both whole saliva and saliva supernatant and verified through experimental work [16]

The “gold standard” for salivary RNA collection termed

“direct saliva transcriptome analysis” (DSTA) [35] has been well used routinely for collection and isolation of RNA (miRNA and mRNA) from patients with multiple diseases The DSTA method involves processing “salivary supernatant” obtained by centrifug-

ing saliva collected by the passive drool technique at 2600 × g for

15 min at 4 °C followed by aspiration from the pellet The salivary supernatant so obtained is stored ready for use at cool tempera-tures, without stabilizing agents, until use mRNAs can be isolated

by one of a number of commercial kits, but in the study by Lee

et al [35], mRNAs were isolated using the MagMAX Viral RNA Isolation kit (Applied Biosystems) The integrity of the mRNAs harvested was confirmed using a series of reference genes This method remains the gold standard for comparative purposes.The discovery [27] that small microvesicles, exosomes found in saliva, contain highly important salivary micro-RNAs (miRNAs) and messenger RNAs (mRNAs) has spawned the development of a series

of tools to capture and interrogate microvesicles, exosomes, and cell-free DNA (and RNA) and miRNAs for transcriptomic analysis

A report by Gallo et al in 2012 [27] confirming that miRNAs

in serum and saliva exist primarily inside exosomes, and that using the exosomal fractions of these bodily fluids increases the sensitivity

of miRNA detection, has focused a lot of attention on various microvesicles, including exosomes

Only recently tools for the analysis and quantification of exosomes in blood have become available, and work has begun on the evaluation of saliva as a readily available source of exosomes, and early work in this area is highly promising

The established standard for exosome isolation involves centrifugation [41]; however, exosomes have also been isolated by precipitation, microfiltration, and antibody-coated magnetic beads Saliva exosome studies have traditionally utilized ultracentrifuga-tion for isolation [42–44]; however, when exosomes were isolated

ultra-by ultracentrifugation from glandular saliva and whole saliva ultra-by Michael et al [42], the authors concluded that viscosity and cel-lular contamination in whole saliva make it a less than ideal medium for exosomal isolation, so a purified saliva specimen may be a more advantageous specimen to use

Cell-free DNA (cfDNA) is an important component for evaluation

of oncological markers in various malignancies [49], for sive prenatal testing (NIPT, [50]), and for other diseases including rheumatoid disease, trauma, myocardial infarction, and fever and inflammatory disease [49, 51–54] Methods for the isolation of cfDNA again typically include blood, amniotic fluid, and other invasive bodily fluids While isolation of cfDNA has been carried out using saliva, the process involves centrifugation of a whole saliva specimen collected by the passive drool technique

noninva-Importantly, at the heart of any successfully developed saliva diagnostic test or procedure is the need to successfully collect, sta-bilize, and recover the sample, so particular emphasis will be placed

on these aspects in the text to follow

2 Materials

A number of commercial tools are now available for the collection

of genomic DNA from saliva specimens (see Note 1)

1 The Oragene device from DNA Genotek (Ottawa, Canada) is the market-leading technology [28] To collect a sample, sub-jects expectorate (“spit”) into the Oragene device until a vol-ume of 2 mL of saliva has been collected A cap on the Oragene device containing proprietary stabilizing buffers is closed, and this causes a stabilizing buffer to flow into the saliva sample, resulting in a laboratory ready sample with long-term shelf life

(1 year) (see Note 2)

1.4 Cell-Free DNA

2.1 Salivary DNA

Collection Procedures

2 The DNA⋅SAL™ device (Oasis Diagnostics®, Vancouver, USA)

is a raking/scraping tool that collects cells from the inside of the oral cavity (buccal mucosa) [23, 29] The collection head

of the DNA⋅SAL™ tool is rubbed gently on the inside of the cheeks for 30 s, resulting in the accumulation of cells on the body of the DNA⋅SAL™ device In addition, cells are abraded

by the mild raking action and remain “free-flowing” in the saliva in the pool formed in the mouth In order to harvest these cells and saliva, a small amount (2.5 mL) of a safe, stabi-lizing rinse solution is taken in the mouth, “swished around,” and then expectorated (spat) back into a collection tube pro-vided The detachable head of the DNA⋅SAL™ device is then removed into the collection tube, to increase the yield of DNA The sample obtained is stable for up to 30 days at room temperature

3 Norgen Biotek (Ontario, Canada) has a device called the Saliva DNA Collection and Preservation Device [30] The principles of this device are similar to the Oragene system In this case, the subject expectorates into a Collection Funnel connected to a Collection Tube until a 2-mL sample of saliva has been collected (marked by a line on the Collection Funnel) The Collection Funnel is removed and may be recy-cled A preservation agent is added to the saliva sample by means of an ampoule, and then the contents of the tube are mixed by shaking and are now ready for analysis or transporta-tion to a laboratory for downstream testing The Norgen sample is stable for up to 2 years

4 The DNAgard® Saliva device from Biomatrica is a relatively new entrant into the field [31] Once again, the Biomatrica device is modeled on similar principles to the Oragene and Norgen DNA devices Subjects expectorate into a tube through

a removable funnel until a “fill mark” is reached The contents

of a dropper bottle are then added to the saliva sample and the mixture inverted 5–7 times to stabilize the sample for up to

30 months at room temperature

5 In addition to methods using passive drool and buccal cell vesting, two well-known technologies use simple swabs Where small to medium quantities of DNA are required, these devices may be suitable

har-(a) The Mawi Technologies iSWAB-DNA Isolation Kit [32, 33] uses a series of routine swabs (iSWABs) for sample collec-tion One of the “iSWABs” is placed in the mouth and rubbed against the inside of the cheek covering the whole cheek while rotating the iSWAB The iSWAB is then placed into a Collection Vial with a narrow neck and screwed down in a corkscrew- like motion until the iSWAB reaches

the bottom of the Collection Vial containing a proprietary buffer solution In order to mix the sample with the liquid

in the Collection Vial, the iSWAB is moved up and down inside the Collection Vial 10–15 times The iSWAB is then removed from the Collection Vial, and the entire proce-dure is repeated with an additional three iSWABs, by alter-nating between the left and right cheek In each case, the iSWAB samples are introduced into the same Collection Vial in order to enrich the sample with DNA Upon com-pletion, a cap is placed on the Collection Vial and the sam-ple stored or analyzed Sample stability is several months at ambient temperature

(b) The Isohelix DNA Buccal Swab kit [34] is described by the manufacturer as “using a unique swab matrix design to efficiently collect buccal cell samples.” Two different swab types are available, and in each case, samples are collected

by rubbing one of the swab types (designated SK-1 and SK-2) firmly against the inside of the cheek or underneath the lower or upper lip for 1 min The head of the swab is then placed into a small Collection Tube, then the swab head removed from the shaft of the device, either by snap-ping the shaft at a notch etched into the side of the shaft (SK-1) or by sliding a plastic cover over the swab head and detaching the swab head by exerting pressure to dislodge the swab head (SK-2) Details of sample stability are not provided

The number of salivary RNA collection methods is fewer than for its counterpart, DNA; however, three or four technologies are worthy of mention:

1 For the Oragene RNA device from DNA Genotek (Ottawa, Ontario, Canada) [36, 37], subjects are asked to place a small amount of table sugar in the palm of their hands then touch the top of their tongue to the sugar, in order to stimulate greater saliva flow The sugar and pooled saliva in the mouth are left there for 10–15 s without swallowing The saliva that pools in the oral cavity is then expectorated into the Oragene container, a plastic Collection Tube Expectoration is contin-ued until a line on the Oragene device is reached (2.0 mL) The sample is then capped and tightened causing a buffer in the cap of the Oragene device to be released into the saliva sample causing immediate stabilization of the sample The mixture of sample and buffer reagent is then shaken vigorously

to mix the sample, which is reported to have a stability of 60 days at ambient temperature The crude Oragene RNA mix-ture may be purified using a number of kits including Qiagen RNeasy Micro or Qiagen RNeasy Mini Kits using a centrifuga-

2.2 Salivary RNA

Collection Procedures

tion followed by pelleting step to obtain purified RNA for

downstream analysis (see Note 3)

2 Norgen Biotek (Canada) offers “Saliva RNA Collection and Purification Devices” [38] based upon identical principles to the Saliva DNA Collection Devices branded by the company

(see Subheading 2.1, item 3) The only significant difference in

the collection procedure is the addition of an RNA stabilizing reagent instead of a DNA stabilizing agent Norgen offers spe-cific kits for isolation of RNA from saliva samples based upon a spin column technique

3 Two devices are available from Oasis Diagnostics® (Vancouver, WA) for transcriptomic workup:

(a) The RNAPro⋅SAL™ device [39] is a system for the taneous harvesting of two “cell-free” samples of saliva that may be used for both RNA and proteins or combined to provide a higher yield of saliva for transcriptomics or pro-teomics In this device, saliva is collected from the pool of saliva in the oral cavity by means of an absorbent pad con-nected to a stem After 1–3 min, saliva collection is com-plete, signified by a color change in a Sample Volume Adequacy Indicator (SVAI), within the device, from yel-low to bright blue The saturated absorbent pad is squeezed through a compression tube and then through a narrow bore filter containing a proprietary filtration medium The sample is subsequently bifurcated (split into two) and col-lected into two equivalent 2-mL Eppendorf tubes where it may be stabilized In the case of proteins, immediate stabi-lization is necessary, and this is facilitated using a protein stabilizing agent provided with the device In the case of RNA, the purified saliva is stable for up to 14 days but may

simul-be stabilized as required by means of “off the shelf” RNA stabilizing reagents The total yield of purified saliva is 1.0 mL

(b) The Pure⋅SAL™ device [40] may be a better option if tein is required In this RNA is required In this case, saliva is collected in identical fashion to the RNAPro⋅SAL™ device,

pro-but a single sample of saliva is collected by squeezing the

saliva sample obtained through a compression tube into which has been inserted a proprietary separation medium

A minimum of 1.0 mL of cell-free saliva is collected into a single 2-mL Eppendorf tube and stabilized as above

Two important applications have been reported for the Pure⋅SAL™ device particularly, which equally apply to the “sis-ter” RNAPro⋅SAL™ technology—these applications are for exosomes and cell-free DNA, each of which can provide increasingly important information on disease and disease pro-cesses of relevance to diagnosis

1 Pure⋅SAL™ Oral Specimen Collection Device (Catalog Number PRSAL-401).

2 Precipitating reagent (ExoQuick-TC, System Biosciences, Mountain View, CA)

3 EXOCET lysis buffer (System Biosciences)

1 Pure⋅SAL™ Oral Specimen Collection Device (Catalog Number PRSAL-401)

2 Falcon tubes

3 Roche High Pure PCR Template Preparation Kit

4 Quant-iT™ PicoGreen® dsDNA Assay Kit (Life Technologies)

3 Methods

Recently, the Pure⋅SAL™ device has been compared to whole saliva and validated for the collection of exosomes [45], quantified using precipitating reagents (ExoQuick-TC Kits) from System Biosciences [46] Isolated exosomes were quantitated by a choles-teryl ester transfer protein (CETP) assay (EXOCET, System Biosciences) validated for the purification and quantification of exosomes [47, 48] It was found that using the Pure⋅SAL™ device simplified collection significantly eliminated non-exosomal con-taminating materials without loss of exosomes A detailed descrip-tion of the method comprising saliva collection, isolation of exosomes, and quantification is detailed below

Collect a saliva specimen by one of the methods described above in Subheading 2.2

1 Combine 1.7 g of collected sample with 340 μL of ExoQuick- TC

and mix by inversion (see Note 6)

2 Incubate overnight at 4 °C

3 Centrifuge sample at 16,000 × g for 5 min.

4 Resuspend resultant pellet in EXOCET lysis buffer (85 μL per tube) and incubate at 37 °C for 5 min

5 Centrifuge at 2000 × g for 5 min.

6 Use resultant supernatant for analysis

Results from the experiments are shown in Table 1 The iment was repeated with a second saliva pool, and similar results were obtained It was noted that if whole saliva is not processed at

sufficient centrifuge speeds, non-exosomal materials remaining in the exosome pellet will interfere with quantitation of exosomes by the cholesteryl ester transfer protein (CETP) assay.

1 Sample collection.

I Pure⋅SAL™: collect a saliva specimen as described above in Subheading 2.2

II Whole Saliva:

(a) Collect saliva by the passive drool technique into a 50-mL Falcon tube

(b) Centrifuge at 3000 × g for 20 min.

(c) Take the supernatant and transfer to another

centri-fuge tube and centricentri-fuge at 16,000 × g for 5 min.

2 Store all samples (I) and (II) at −80 °C prior to DNA isolation

3 DNA isolation

(a) Isolate DNA with the Roche High Pure PCR Template Preparation Kit by using 700 μL saliva aliquots per isolation

4 DNA quantification using PicoGreen

(a) Measure DNA quantity using the Quant-iT™ PicoGreen®

dsDNA Assay Kit (see Note 7)

●

● Prepare a standard curve using ten different trations of lambda DNA provided in the kit Perform triplicate readings for increased precision

3.3 Cell-Free DNA

Table 1

Comparison of the quantity of salivary exosomes collected by the Pure ⋅SAL™ device and whole

saliva followed by centrifugation

Process for sample isolation Number of exosomes per mL DNA ( μg/mL) Protein (mg/mL)

Whole saliva—centrifuged 16,000 × g 3.10 × 109 1.47 4.75

4 Notes

1 DNA from samples collected using one of the above cial tools may be isolated using one of a significant number of DNA isolation kits provided by a number of manufacturers The number of possibilities available is too numerous to cover

commer-in this manuscript; however, a number of manufacturers have developed specific saliva kits or validated certain kits to work well for saliva specimens The list includes Qiagen Corporation (www.Qiagen.com), DNA Genotek (www.DNAGenotek.com), Norgen Biotek (www.NorgenBiotek.com), Biomatrica (www.Biomatrica.com), Oasis Diagnostics® (www.4saliva.com), Life Technologies (www.ThermoFisher.com), and others

2 DNA Genotek received FDA 510(k) clearance for the use of Oragene in conjunction with a test for warfarin sensitivity developed by the company GenMark Diagnostics, so the device

may be used clinically for this single application.

3 For RNA isolation, there are fewer kits available that have been specifically optimized for saliva specimens The Qiagen miR-Neasy kit has been used successfully for the isolation of purified RNA for transcriptome work, RNA sequencing, and other applications, as has the QIAzol lysis reagent from the same company Other methods that have been used include organic extraction methods (TRIzol LS), spin filter-based methods (QIAamp Viral (Qiagen)), NucleoSpin (Clontech), and miR-Vana (Life Technologies) and combined method of organic extraction and spin filter clean up (miRNeasy micro (Qiagen)) and Quick-RNA MicroPrep (Zymo Research)

4 In reference to Subheading 1.4, the performance of one ticular device (the Pure⋅SAL™ device) has been evaluated side- by- side with the “gold standard” method (passive drool/centrifugation) for cell-free DNA according to protocols out-lined in the manuscript [55] In the experiments performed, the Pure⋅SAL™ device was found to be a superior tool for har-vesting cfDNA

5 In Subheading 2.1, care should be taken to investigate options for DNA purification based upon the specific application required These may include simple ethanol precipitation tech-niques, spin column methods, 96-well microplates, or auto-mated methods, such as the Promega Maxwell 16 instrument

or the Qiagen QIAsymphony equipment Whole saliva tains a significant quantity of mucinous material that can have

con-an impact on the quality of DNA obtained It is recommended that investigators contact the individual manufacturers for details of any methods and how they may be applied to DNA isolation from saliva, prior to the commencement of any vali-dation studies

6 The method used in this chapter for isolation of exosomes is only one of a number of exosomal isolation kits now available These include the Exo-spin kit from Cell Guidance Systems, Total Exosome Isolation Reagent from Thermo Fisher, miR-CURY from Exiqon, PureExo Exosome Isolation kit from PureExo, and ExoCap Capture Kit from JSR Biosciences Investigators are encouraged to validate the best method for exosome isolation in their own laboratory.

7 The authors also carried out DNA quantification by tive PCR (qPCR) as an alternate method of DNA assessment

quantita-Acknowledgments

The author would like to acknowledge the support of Dr David T Wong (UCLA) for his support and encouragement in preparing this manuscript

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Chapter 2

RNA Sequencing Analysis of Salivary Extracellular RNA

Blanca Majem, Feng Li, Jie Sun, and David T.W Wong

Abstract

Salivary biomarkers for disease detection, diagnostic and prognostic assessments have become increasingly well established in recent years In this chapter we explain the current leading technology that has been used to characterize salivary non-coding RNAs (ncRNAs) from the extracellular RNA (exRNA) fraction: HiSeq from Illumina® platform for RNA sequencing Therefore, the chapter is divided into two main sec- tions regarding the type of the library constructed (small and long ncRNA libraries), from saliva collection, RNA extraction and quantification to cDNA library generation and corresponding QCs Using these invaluable technical tools, one can identify thousands of ncRNA species in saliva These methods indicate that salivary exRNA provides an efficient medium for biomarker discovery of oral and systemic diseases.

Key words Saliva, exRNAs, Small and long ncRNA profiling, Biomarkers, RNA sequencing

1 Introduction

Extracellular RNA (exRNA) in human saliva is an emerging field for noninvasive diagnostic applications The discovery of saliva- derived mRNA in normal and oral cancer patients [1–3] and other forensic applications [4 5] opened up a new field for noninvasive molecular diagnosis Our laboratory has extensively studied microarray- based gene profiling followed by real-time quantitative- PCR (RT-qPCR) for saliva mRNA detection We have identified certain macromole-cules associated with salivary mRNA that were protecting against ribonucleases [6] Salivary RNA was found in complexes with lipids, proteins, lipoproteins, and phospholipids as well [7 8] Apoptotic bodies [9] or other vesicular structures in saliva also play a protec-tion role Therefore, RNA in the saliva may not be as fragile as it was previously assumed to be Despite the numerous studies based on characterizing and finding mRNA diagnostic biomarkers in saliva, the introduction of deep sequencing technologies [10, 11] has revealed a new landscape of salivary exRNA [12]: micro-RNAs (miRNAs), piwi-interacting-RNAs (piRNAs), circular-RNAs (cir-cRNAs), and other noncoding RNAs (ncRNAs) To date, only a few

studies characterizing ncRNAs in saliva have used RNA-sequencing (RNA-Seq) technologies [13] In this chapter, we present the detailed methodology for RNA extraction, cDNA library construc-tion and quality controls (QCs), and data analysis of sequencing data Although a variety of platforms are available for RNA-seq, the Illumina® platform is the most used nowadays Increasing knowl-edge on salivary composition thanks to this platform will make a difference in understanding the biology of the diagnostic biomark-ers found in saliva for local and systemic diseases.

The purpose of this chapter is to provide robust and reliable methods for isolating and profiling of salivary exRNA, dividing it in two main sections regarding the type of the library (small and long ncRNA libraries) constructed We also describe a protocol for RNA extraction after saliva collection, including detailed explanations of DNase treatment, RNA precipitation for sample concentration and specific QCs for the extracted RNA The commercial kits for RNA extraction allow high RNA yield, but the eluted RNA usually is contained in big volumes and therefore low concentrations, which

is not recommended for subsequent steps, since library preparation starts with little volumes and requires a high concentration sample Either way, the lower limit of detection of the QCs makes the RNA precipitation crucial for sample concentration, resulting in high reproducibility among samples and accurate RNA and cDNA quan-tification, which at the same time is translated into good quality of raw read data after sequencing Thus, our protocols are a guide for RNA-seq of salivary exRNA, but some concepts and methodology may also be applied to other types of body fluids

2 Materials

1 50 mL sterile tube

2 Laboratory vortex

3 Refrigerated benchtop centrifuge with 50 mL tube adapters

4 SUPERase-In RNase inhibitor, Cat# AM2694 Ambion

4 Absolute ethanol.

5 Nuclease-free water

1 QuanTi™ RiboGreen RNA assay kit

2 96-Well half area microplate (black solid plate), Cat# 3694 Corning

3 Agilent RNA 6000 Pico kit

1 NEBNext® Multiplex Small RNA library Prep Set for Illumina®

2 Exiqon Spike-in miRNA kit v2, Cat# 208041 Exiqon

3 8-Tube PCR strip

4 Thermal Cycler PCR machine

5 6 % Novex® TBE PAGE gel, 1.0 mM 10-well

6 SYBR® Gold Nucleic Acid Gel Stain (Life Technologies, Inc

#S-11494)

7 Gel broker tubes, Cat#, 3388-100 SeqMatic

8 Corning®, Costar®, Spin-X® Centrifuge Tube Filters (Cellulose Acetate Filters)

9 3 M Sodium Acetate, pH 5.5

10 100 and 80 % ethanol (freshly prepared)

11 QIAquick PCR purification kit

1 NEBNext Ultra Directional RNA Library Prep kit for Illumina®

2 ERCC spike-in, Cat# 4456740 Ambion

3 NEBNext Singleplex or NEBNext Multiplex Oligos for Illumina®

4 Actinomycin D (Sigma# A1410, dissolved in ide [DMSO] to 5 μg/μL)

5 8-tube PCR strip

6 80 % thanol (freshly prepared)

7 Thermal Cycler PCR machine

8 DynaMag™-2 Magnet

9 Agencourt® AMPure® XP Beads (Beckman Coulter, Inc

#A63881)

1 Qubit® dsDNA BR assay kit

2 96-well half area microplate (black solid plate), Cat# 3694 Corning

3 Agilent High Sensitivity DNA Kit

2.2.3 RNA Quantification

and QCs

2.2.4 cDNA Library

Preparation

Small ncRNA Library

Long ncRNA Library

2.2.5 cDNA Library

Quantification and QCs

1 EB buffer.

2 Tween™ 20 Surfact-Amps™ detergent solution

3 HiSeq2000 Illumina system

1 Cutadapt

2 Bowtie mapping 16 s rRNA/Microbiome

3 Bowtie mapping Human Genome

1 Ask subjects to refrain from eating, drinking, smoking, or oral hygiene procedures for at least 1 h prior to collection

2 Instruct subjects to rinse the mouth thoroughly with water and to void the mouth of saliva The subject should be seated comfortably with eyes open and head tilted slightly forward For unstimulated saliva collection subjects should rest for

5 min and minimize orofacial movements

3 To collect un-stimulated saliva (see Note 1) allow, saliva to accumulate in the floor of the mouth and ask the subject to spit into a preweighed or graduated test tube every 60 s Collection for 5 min usually yields sufficient saliva (~5 mL) for analysis

4 Following collection, centrifuge saliva samples at 2600 × g for

15 min at 4 °C Saliva supernatant will then be separated from the cellular phase

5 Add SUPERase-In RNase inhibitor (at a ratio of 1 μL/mL) to

1 mL of cell-free saliva (CFS) supernatant for preserving exRNA degradation

6 Store aliquots of 1 mL CFS at −80 °C for further analysis

1 Thaw 4 aliquots of 1 mL of saliva, resting the tubes on ice and

not for more than half an hour (see Note 2)

2 Split the sample in 500 μL of CFS and centrifuge for 5 min at

10,000 × g (see Note 3) Collect the supernatant to proceed with step 2, and discard the pellet fraction.

3 Split 0.5 mL of cell free saliva (CFS) in two tubes—250 μL in each

4 Add 750 μL of Qiazol to 250 μL of CFS Vortex for 30 s and incubate 5 min at RT

5 Add 200 μL chloroform and mix by vortex for 30 s, and then incubate 5 min at RT.

6 Centrifuge the sample at 12,000 × g for 15 min at 4 °C.

7 Carefully collect 600 μL (at least) of upper aqueous phase and transfer to the new tubes

8 Add 900 μL (1.5 Vol) of 100 % ethanol and mix thoroughly by pipetting up and down several times Do not centrifuge Continue without delay to the next step

9 Pipette 700 μL of the sample into an RNeasy MinElute spin

column Centrifuge at 9300 × g for 30 s at RT Discard the

flow-through Repeat this step using the remaining sample

10 Pipette 700 μL buffer RWT into the RNeasy MinElute spin

column and centrifuge at 9300 × g for 30 s to wash Discard

the tube with flow-through and place the column in a new

2 mL collection tube

11 Pipette 700 μL Buffer RPE onto the RNeasy MinElute spin

column Close the lid gently and centrifuge at 9300 × g for 30 s

to wash the column Discard the flow-through

12 Pipette 300 μL Buffer RPE onto the RNeasy MinElute spin

column Close the lid gently and centrifuge at 9300 × g for 30 s

to wash the column Discard the flow-through

13 Pipette 500 μL of 80 % ethanol onto the RNeasy MinElute

spin column (see Note 4) Close the lid and centrifuge at

≥9300 × g for 2 min to wash membrane Discard the collection

tube with the flow-through

14 Place the RNeasy MinElute spin column into a new 2 mL lection tube Centrifuge at full speed for 5 min to dry the membrane Discard the collection tube

15 Place the column in a new 1.5-mL tube Add 30 μL preheated water (~50 °C) directly to the center of the membrane Close the lid and incubate for 1–2 min at RT, and then centrifuge for

1 min at full speed

16 Maintain the column in the same tube Add 30 μL more (see

Note 5) of preheated water directly to the center of the

mem-brane Close the lid and incubate for 1–2 min at RT (see Note

6), and then centrifuge for 1 min at full speed Proceed directly

to DNase treatment and RNA precipitation step (see Note 7)

1 Mix the next components to perform off-column DNase ment to the eluted RNA of 8 samples at the same time:

treat-– 2 μL TURBO DNase—18 μL (for 8 samples)

– 11 μL Buffer—99 μL (for 8 samples)

– 27 μL H2O (nuclease-free)—243 μL (for 8 samples)

3.2.2 DNase Treatment

and RNA Precipitation

2 Add 40 μL DNase Mix/sample (100 μL final volume = 60 μL RNA + 40 μL DNase Mix).

3 Leave it for 15 min at RT Continue with step 4 for RNA

precipitation.

4 Add 10 μL (0.1 Vol) of sodium acetate 3 M pH5.5

5 Add 1 μL (5 μg) of glycogen (Glycogen is at 5 μg/μL concentration)

1 Prepare serial dilution of rRNA standards (12.5–200 ng/mL)

2 Make 70 μL aliquots of each standard and stock at −80 °C for future use

3 Make 5 μL aliquots of fluorescent dye (Component A in RiboGreen kit) and stock them at −80 °C (see Note 9)

4 Take one set of standards and one Fluorescent Dye aliquot from freezer and thaw them at room temperature (RT) in the

dark (important for the Dye) (see Note 10)

5 Prepare RNA sample dilutions at 1/30 in 1× TE buffer: Mix

1 μL of RNA/sample and 29 μL of 1× TE buffer for each sample

6 Prepare enough working solution (WS) for all the experiment

at a ratio of 1:200 dilution of Fluorescent Dye: 1× TE into a

15 mL tube (in the darkness)

7 Plate 15 μL of the standards (multichannel micropipette is ommended for reproducibility) in triplicate, and 15 μL of diluted samples in duplicate

8 Add 15 μL of the WS into each well (standard and samples) and incubate the plate for 15 min at RT in the dark (with lid)

9 Read the plate at 480–520 nm in a spectrophotometer (see

gel-– Quant-iT Ribogreen RNA assay: salivary RNA tion normally ranges from 50 to 80 ng/mL saliva If total RNA amount is <5 ng it is not recommended to proceed with the library construction

– RNA 6000 Pico Chip, Bioanalyzer: detection of intact ribosomal RNA peak indicates residual cell contamination (eukaryotic: 18S (1869 nt), 28S rRNA (5070 nt); pro-karyotic: 16S (1542 nt), 23S rRNA (2906 nt)) and excludes the sample for further analysis.

Prepare the Exiqon Spike-in miRNA kit v2: Dissolve the CURY LNA™ Array Spike-in microRNA Kit v2 in 30 μL/vial of nuclease-free water (supplied) upon receipt Vortex to thoroughly dissolve the lyophilized RNA, pulse briefly in a microfuge, and leave the suspension on ice for 30 min to dissolve Aliquot the dissolved spike-in miRNAs and store at –80 °C until use and avoid repeated cycles of freeze/thawing

1 Mix the following components in a sterile nuclease-free PCR tube:

(Green) 3 ′ SR adaptor for Illumina 1 μL

2 Incubate in a preheated thermal cycler for 2 min at

70 °C Transfer tube to ice

3 Add the following components:

RNA+ 3 ′ SR adaptor mix 7 μL (Green) 3 ′ ligation reaction buffer (2×) 10 μL (Green) 3 ′ ligation enzyme mix 3 μL

4 Incubate for 1 h at 25 °C in a thermal cycler

1 Add the following components to the ligation mixture from

step 4 and mix well:

3′ Ligation reaction mix from step 1 20 μL Nuclease-free water 4.5 μL (Pink) SR RT primer for Illumina 1 μL Total volume now should be 25.5 μL

2 Heat samples for 5 min at 75 °C Transfer to 37 °C for 15 min, followed by 15 min at 25 °C

3.2.4 cDNA Library

Preparation

Small ncRNA Library

Hybridize the Reverse

Transcription Primer

1 With 5 min remaining, resuspend the (yellow) 5′ SR adaptor in

120 μL of nuclease-free water and store at −80 °C (This step is only necessary when the kit is first opened)

2 Aliquot 1.1 μL × N of the (yellow) 5′ SR Adaptor into a

sepa-rate, 200 μL nuclease-free PCR tube, with N equal to the

number of samples being processed for the current experiment

3 Incubate the adaptor in the thermal cycler at 70 °C for 2 min and then immediately place the tube on ice Keep the tube on ice and use the denatured adaptor within 30 min of denaturation

4 Add the following components to the ligation mixture from

step 6 and mix well:

Reaction mix from step 2 25.5 μL (Yellow) 5 ′ SR Adaptor for Illumina

(Yellow) 5 ′ ligation reaction buffer (10×) 1 μL (Yellow) 5 ′ ligation enzyme mix 2.5 μL

5 Incubate for 1 h at 25 °C in a thermal cycler

Mix the following components in a sterile, nuclease-free tube:

Adaptor ligated RNA from step 3 30 μL (Red) first strand synthesis reaction buffer 8 μL (Red) murine RNase inhibitor 1 μL (Red) ProtoScript II reverse transcriptase 1 μL

Incubate for 60 min at 50 °C

Immediately proceed to PCR amplification

Safe Stopping Point: If you do not plan to proceed immediately to PCR amplification, then heat inactivate the RT reaction at 70 °C for

15 min Samples can be safely stored at –15 to –25 °C.

Add the following components to the RT reaction mix from step

4 and mix well:

Perform Reverse

Transcription

Perform PCR Amplification

RT reaction mix from step 4 40 μL (Blue) LongAmp Taq 2× Master Mix 50 μL (Blue) SR Primer for Illumina 2.5 μL

1 Add 500 μL Buffer PB to the PCR reaction and mix

2 Apply the sample to the QIAquick column and centrifuge at

15,600 × g for 30–60 s.

3 Add 750 μL Buffer PE to the QIAquick column and

centri-fuge at 15,600 × g for 30–60 s.

4 Centrifuge the column with the lid of the spin column open

for 5 min at 15,600 × g (see Note 12)

5 Place each QIAquick column in a clean 1.5 mL fuge tube

6 To elute amplified DNA add 26 μL Nuclease-free Water Let

the column stand for 1 min, and then centrifuge at 15,600 × g

for 1 min

Prepare 500 mL Running Buffer (100 mL 5× Running Buffer + 400 mL Water), leave 100 mL Running Buffer with 10 μL SYBR Gold stain for later use

1 Mix the purified PCR product (25 μL) with 10 μL of Gel Loading Dye, Blue (6×)

2 Load 5 μL of Quick-Load pBR322 DNA-MspI Digest in a well on the 6 % PAGE 10-well gel

3 Load two wells with 17 μL each of mixed amplified cDNA and loading dye on the 6 % PAGE 10-well gel

4 Run the gel for ~1.5 h at 90 V Do not let the blue dye exit the gel.

5 Remove the gel from the apparatus and stain the gel with SYBR Gold nucleic acid gel stain in a clean container for 2–3 min and view the gel on a UV transilluminator The 140 and 150 nt bands correspond to adapter-ligated constructs derived from the 21 and 30 nt RNA fragments, respectively For miRNAs, isolate the bands corresponding to ~140 bp For piRNAs, iso-

late the band corresponding to ~150 bp (see Fig 1)

6 Place the two gel slices from the same sample in a Gel Broker tube

(SeqMatic) with a 2 mL tube, then centrifuge at 14,000 × g for

1 min, and then soak in 400 μL DNA Gel Elution buffer (1×)

7 Rotate in eppendorf shaker for at least 2 h at RT

8 Transfer the eluate and the gel debris to SpinX column with

1 cm diameter Whatman filter

9 Centrifuge the filter for 2 min at >15,600 × g.

10 Recover eluate and add 1 μL linear acrylamide, 40 μL 3 M sodium acetate pH 5.5, 500 μL of 100 % ethanol, and 500 μL

of isopropanol Vortex well

11 Precipitate at –20 °C for at least 4 h or −80 °C at least 1.5 h

12 Spin >15,600 × g for 30 min at 4 °C.

13 Remove the supernatant, taking care not to disturb the pellet

14 Wash the pellet with 500 μL 80 % ethanol

Fig 1 Transilluminator view of miRNA and piRNA bands The lanes S1 to S4

cor-respond to 4 different small ncRNA libraries Each library has been run per cate and bands were cut below 140 bp and above 300 bp miRNAs isolated bands correspond to ~140 bp piRNAs isolated bands correspond to ~150 bp

15 Spin >15,600 × g for 10 min at 4 °C.

16 Air-dry pellet for up to 10 min at RT to remove residual ethanol

17 Resuspend pellet in 12 μL EB Buffer (2 of 12 μL will be used for cDNA library quantification in Subheading 3.2.5)

Prepare the ERCC spike-in: Dissolve in nuclease-free water the lyophilized product making 1:100 dilution stocks Vortex to thor-oughly dissolve the lyophilized RNA, pulse briefly in a microfuge, and leave the suspension on ice for 30 min to dissolve Aliquot (1–2 μL) the dissolved spike-in RNAs and store at –80 °C until use and avoid repeated cycles of freeze/thawing

Total saliva RNA + ERCC spike-in (4.5 μL of

1 Incubate the samples at 94 °C for 2 min

2 Transfer the tube on ice

3 Proceed to First Strand cDNA Synthesis

Dilute Actinomycin D stock solution (5 μg/μL) to 0.1 μg/μL

in nuclease-free water for immediate use

The fragmented and primed mRNA 10 μL (Pink) murine RNase inhibitor 0.5 μL Actinomycin D (0.1 μg/μL) 5 μL (Pink) ProtoScript II reverse transcriptase 1 μL

Long ncRNA Library

Preparation of First Strand

The First Strand Synthesis reaction mixes 20 μL

1 Mix thoroughly by gentle pipetting

2 Incubate in thermal cycler for 1 h at 16 °C, with heated lid set

at ≥40 °C

1 Vortex AMPure XP beads to resuspend

2 Add 200 μL (2.5×) of resuspended AMPure XP beads to the second strand synthesis reaction (≈80 μL) Mix well on a vor-tex mixer or by pipetting up and down at least 10 times

3 Incubate for 5 min at RT

4 Quickly spin the tube in a microcentrifuge to collect any ple on the sides of a tube Place the tube on an appropriate magnetic rack (DynaMag™-2 Magnet) to separate beads from supernatant After the solution is clear (about 5 min), carefully remove and discard the supernatant Be careful not to disturb the beads that contain DNA targets

5 Add 200 μL of freshly prepared 80 % ethanol to the tube while

in the magnetic rack Incubate at RT for 30 s, and then fully remove and discard the supernatant

6 Repeat step 5 once for a total of 2 washing steps.

7 Air-dry the beads for 10 min while the tube is on the magnetic rack with lid open (recommend hood)

8 Elute the DNA target from the beads into 60 μL nuclease-free water Mix well on a vortex mixer or by pipetting up and down Quickly spin the tube and then place it in the magnetic rack until the solution is clear

9 Remove 55.5 μL of the supernatant and transfer to a clean nuclease-free PCR tube

The purified double- stranded cDNA 55.5 μL (Green) NEBNext end repair reaction buffer (10×) 6.5 μL (Green) NEBNext end prep enzyme mix 3 μL

30 min at 65 °C.

Hold at 4 °C

Proceed immediately to Adaptor Ligation

Dilute the NEBNext Adaptor for Illumina (15 μM) to 1.5 μM with a 10-fold dilution (1:9) with nuclease-free water for immedi-ate use

(Red) Blunt/TA Ligase Master Mix 15 μL (Red) Diluted NEBNext adaptor 1 μL Nuclease-free water 2.5 μL

Incubate 15 min at 20 °C in a thermal cycler

The adaptor is provided in NEBNext Singleplex or NEBNext Multiplex Oligos for Illumina

1 To the ligation reaction (83.5 μL), add 16.5 μL nuclease-free water to bring the reaction volume to 100 μL

2 Add 100 μL (1.0×) resuspended AMPure XP beads and mix well

on a vortex mixer or by pipetting up and down at least 10 times

3 Incubate for 5 min at RT

4 Quickly spin the tube in a microcentrifuge and place the tube

on an appropriate magnetic rack to separate beads from natant After the solution is clear (about 5 min), discard the supernatant that contains unwanted fragments (Caution: do not disturb the beads)

5 Add 200 μL of freshly prepared 80 % ethanol to the tube while

in the magnetic rack Incubate at RT for 30 s, and then fully remove and discard the supernatant

6 Repeat step 5 once for a total of two washing steps.

7 Briefly spin the tube, and put the tube back in the magnetic rack

8 Completely remove the residual ethanol, and air-dry beads for

10 min while the tube is on the magnetic rack with the lid open (recommend hood)

9 Elute DNA target from the beads with 50 μL nuclease-free water Mix well on a vortex mixer or by pipetting up and down, and put the tube in the magnetic rack until the solution is clear

10 Transfer the 50 μL supernatant to a clean PCR tube Discard the beads

Perform Adaptor Ligation

Purify the Ligation

Reaction Using AMPure

XP Beads

11 To the 50 μL supernatant, add 50 μL (1.0×) of the pended AMPure XP beads and mix well on a vortex or by pipetting up and down at least 10 times.

12 Incubate for 5 min at RT

13 Quickly spin the tube in a microcentrifuge and place the tube

on an appropriate magnetic rack to separate beads from the supernatant After the solution is clear (about 5 min), discard the supernatant that contains unwanted fragments (Caution:

do not discard the beads)

14 Add 200 μL of freshly prepared 80 % ethanol to the tube while

in the magnetic rack Incubate at RT for 30 s, and then fully remove and discard the supernatant

15 Repeat step 14 once for a total of two washing steps.

16 Briefly spin the tube, and put the tube back in the magnetic rack

17 Completely remove the residual ethanol, and air-dry beads for

10 min while the tube is on the magnetic rack with the lid open (recommend hood)

18 Elute DNA target from the bead with 25 μL nuclease-free water Mix well on a vortex mixer or by pipetting up and down, and put the tube in the magnetic rack until the solution is clear

19 Without disturbing the bead pellet, transfer 20 μL of the supernatant to a clean PCR tube and proceed to PCR enrich-

ment (see Note 13)

Optional stopping point: at this point cDNA library can be

stored at −20 °C

The Universal PCR primer and Index (X) Primer are contained in the NEBNext SinglePlex or NEBNext Multiplex Oligos for Illumina

(Blue) NEBNext USER enzyme 3 μL (Blue) NEBNext High-Fidelity PCR Master Mix, 2× 25 μL (Blue) Universal PCR Primer (25 μM) 1 μL (Blue) Index (X) Primer (25 μM) 1 μL