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

Methods in
Molecular Biology 1537

Gregory J. Seymour
Mary P. Cullinan
Nicholas C.K. Heng Editors

Oral Biology
Molecular Techniques
and Applications
Second Edition


Methods

in

Molecular Biology

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

For further volumes:
/>

Oral Biology
Molecular Techniques and Applications
Second Edition

Edited by

Gregory J. Seymour
Faculty of Dentistry, University of Otago, Dunedin, New Zealand

Mary P. Cullinan
Department of Oral Sciences, Faculty of Dentistry, University of Otago, Dunedin, New Zealand

Nicholas C.K. Heng
Faculty of Dentistry, Sir John Walsh Research Institute, University of Otago, Dunedin, New Zealand


Editors
Gregory J. Seymour
Faculty of Dentistry
University of Otago
Dunedin, New Zealand

Mary P. Cullinan
Department of Oral Sciences, Faculty of Dentistry
University of Otago
Dunedin, New Zealand

Nicholas C.K. Heng
Faculty of Dentistry, Sir John Walsh Research
Institute
University of Otago
Dunedin, New Zealand

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
Library of Congress Control Number: 9781493967384
© Springer Science+Business Media LLC 2017
This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is
concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction
on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation,
computer software, or by similar or dissimilar methodology now known or hereafter developed.
The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not
imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and
regulations and therefore free for general use.
The publisher, the authors and the editors are safe to assume that the advice and information in this book are believed to
be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty,
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
in Chapter 20.
Printed on acid-free paper
This Humana Press imprint is published by Springer Nature
The registered company is Springer Science+Business Media LLC
The registered company address is: 233 Spring Street, New York, NY 10013, U.S.A.


Preface
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 assessing 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 profiling of oral microbial communities, quantitative real-time PCR, and adhesion of yeast and
bacteria to oral surfaces have all been retained but substantially updated.
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 anticipated 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.
Dunedin, New Zealand
Dunedin, New Zealand

Dunedin, New Zealand

Gregory J. Seymour
Mary P. Cullinan
Nicholas C.K. Heng

v


Contents
Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
v
Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xi

Part I  Saliva and Other Oral Fluids
  1 Salivary Diagnostics Using Purified Nucleic Acids . . . . . . . . . . . . . . . . . . . . . . .
Paul D. Slowey
  2 RNA Sequencing Analysis of Salivary Extracellular RNA . . . . . . . . . . . . . . . . . .
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 . . . . . . . . . . . . . . . . .
Erdjan Salih
  4 Antioxidant Micronutrients and Oxidative Stress Biomarkers . . . . . . . . . . . . . .
Iain L.C. Chapple, Helen R. Griffiths, Mike R. Milward,
Martin R. Ling, and Melissa M. Grant
  5 NMR-Based Metabolomics of Oral Biofluids . . . . . . . . . . . . . . . . . . . . . . . . . .
Horst Joachim Schirra and Pauline J. Ford
  6 Gene Therapy of Salivary Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Bruce J. Baum, Sandra Afione, John A. Chiorini, Ana P. Cotrim,
Corinne M. Goldsmith, and Changyu Zheng

3
17

37
61

79
107

Part II  Molecular Biosciences
  7 The Oral Microbiota in Health and Disease: An Overview
of Molecular Findings . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
José F. Siqueira Jr., Mitsuo Sakamoto, and Alexandre S. Rosado
  9 Analysis of 16S rRNA Gene Amplicon Sequences
Using the QIIME Software Package . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Blair Lawley and Gerald W. Tannock
10 Adhesion of Yeast and Bacteria to Oral Surfaces . . . . . . . . . . . . . . . . . . . . . . . .
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) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Mª José Marin, Elena Figuero, David Herrera, and Mariano Sanz

vii

127

139

153
165

191


viii

Contents

12 Methods to Study Antagonistic Activities Among Oral Bacteria . . . . . . . . . . . .
Fengxia Qi and Jens Kreth
13 Natural Transformation of Oral Streptococci by Use of Synthetic
Pheromones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Gabriela Salvadori, Roger Junges, Rabia Khan, Heidi A. Åmdal,
Donald A. Morrison, and Fernanda C. Petersen
14 Markerless Genome Editing in Competent Streptococci . . . . . . . . . . . . . . . . . .
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 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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 . . . . . . . . . . . . . . . . . . . . . . . .
Trudy J. Milne
18 Genome-Wide Analysis of Periodontal and Peri-Implant Cells and Tissues . . . .
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 . . . . . . . . . . . . . .
Moritz Kebschull, Melanie Julia Fittler, Ryan T. Demmer,
and Panos N. Papapanou
20 Exploring Genome-Wide Expression Profiles Using Machine
Learning Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Moritz Kebschull and Panos N. Papapanou

203

219

233

249

279

299
307

327

347

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


Contents

24 A Method to Isolate, Purify, and Characterize Human Periodontal
Ligament Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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 . . . . . . . . . . . . .
Lynne Bingle, Felipe P. Fonseca, and Paula M. Farthing
26 Growing Adipose-Derived Stem Cells Under Serum-Free Conditions . . . . . . . .
Diogo Godoy Zanicotti and Dawn E. Coates
27 Quantitative Real-Time Gene Profiling of Human Alveolar Osteoblasts . . . . . .
Dawn E. Coates, Sobia Zafar, and Trudy J. Milne
28 Proteomic Analysis of Dental Tissue Microsamples . . . . . . . . . . . . . . . . . . . . . .
Jonathan E. Mangum, Jew C. Kon, and Michael J. Hubbard

29 Characterization, Quantification, and Visualization of Neutrophil
Extracellular Traps . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Phillipa C. White, Ilaria J. Chicca, Martin R. Ling, Helen J. Wright,
Paul R. Cooper, Mike R. Milward, and Iain L.C. Chapple

ix

413

429
439
447
461

481

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 499


Contributors
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

xi


xii

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

Contributors

xiii

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


xiv

Contributors

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


Chapter 1
Salivary Diagnostics Using Purified Nucleic Acids
Paul D. Slowey
Abstract
Saliva is an easily accessible fluid that has led to increasing interest in the development of salivary diagnostics. This chapter 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.
Key words Saliva, RNA, DNA, Nucleic acids, Stabilization, Exosomes

1  Introduction
Over the last few years, the use of saliva as a noninvasive bodily fluid
for research, forensic, and clinical testing has grown tremendously
and is now in use in many areas of the global in vitro diagnostic
(IVD) market.
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 workplace 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

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

3


4

Paul D. Slowey

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 biomarkers that are indicative of disease and maladies affecting human
beings, so saliva is the ideal sample matrix for large-scale epidemiological 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 “explosion” 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 contributed 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
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 hormones 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


Salivary Diagnostics Using Purified Nucleic Acids

5

preferable biofluid to urine, which is currently the method of
choice. This is particularly true in the case of marijuana, when testing 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 transported 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 phlebotomist to collect a sample is the key driver in this instance.
1.1  Salivary DNA
Collection

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.

1.2  Salivary RNA
Collection

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 centrifuging 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 temperatures, 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.

1.3  Exosomes

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.


6

Paul D. Slowey

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 s­ ensitivity
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 ultracentrifugation [41]; however, exosomes have also been isolated by
precipitation, microfiltration, and antibody-coated magnetic beads.

Saliva exosome studies have traditionally utilized ultracentrifugation for isolation [42–44]; however, when exosomes were isolated
by ultracentrifugation from glandular saliva and whole saliva by
Michael et al. [42], the authors concluded that viscosity and cellular 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.
1.4  Cell-Free DNA

Cell-free DNA (cfDNA) is an important component for evaluation
of oncological markers in various malignancies [49], for noninvasive 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.
Importantly, at the heart of any successfully developed saliva
diagnostic test or procedure is the need to successfully collect, stabilize, and recover the sample, so particular emphasis will be placed
on these aspects in the text to follow.

2  Materials
2.1  Salivary DNA
Collection Procedures

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, subjects expectorate (“spit”) into the Oragene device until a volume 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).


Salivary Diagnostics Using Purified Nucleic Acids

7

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, stabilizing rinse solution is taken in the mouth, “swished around,”
and then expectorated (spat) back into a collection tube provided. 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 recycled. 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 transportation 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 harvesting, two well-known technologies use simple swabs. Where
small to medium quantities of DNA are required, these devices
may be suitable.
(a) The Mawi Technologies iSWAB-DNA Isolation Kit [32, 33]
uses a series of routine swabs (iSWABs) for sample collection. 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


8

Paul D. Slowey

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 procedure is repeated with an additional three iSWABs, by alternating 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 completion, a cap is placed on the Collection Vial and the sample 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 snapping 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.
2.2  Salivary RNA
Collection Procedures

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 continued 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 mixture may be purified using a number of kits including Qiagen
RNeasy Micro or Qiagen RNeasy Mini Kits using a centrifuga-


Salivary Diagnostics Using Purified Nucleic Acids

9

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 specific 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 simultaneous 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 proteomics. In this device, saliva is collected from the pool of
saliva in the oral cavity by means of an absorbent pad connected to a stem. After 1–3 min, saliva collection is complete, signified by a color change in a Sample Volume
Adequacy Indicator (SVAI), within the device, from yellow 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 collected into two equivalent 2-mL Eppendorf tubes where it

may be stabilized. In the case of proteins, immediate stabilization 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
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 protein is required. In this RNA is required. In this case, saliva is
collected in identical fashion to the RNAPro⋅SAL™ device,
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 “sister” RNAPro⋅SAL™ technology—these applications are for
exosomes and cell-free DNA, each of which can provide
increasingly important information on disease and disease processes of relevance to diagnosis.


10

Paul D. Slowey

2.3  Exosomes

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).

2.4  Cell-Free DNA
(cfDNA)

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 cholesteryl 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 contaminating materials without loss of exosomes. A detailed description of the method comprising saliva collection, isolation of
exosomes, and quantification is detailed below.
3.1  Sample
Collection
and Stabilization
3.2  Isolation
of Exosomes

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 experiment was repeated with a second saliva pool, and similar results
were obtained. It was noted that if whole saliva is not processed at


Salivary Diagnostics Using Purified Nucleic Acids

11

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

Pure⋅SAL™ device

1.19

4.58

3.25 × 109

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.
3.3  Cell-Free DNA

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 centrifuge tube and centrifuge 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 concentrations of lambda DNA provided in the kit. Perform
triplicate readings for increased precision.
Construct a standard curve using the values from the
ten different concentrations of lambda DNA.
Measure the samples relative to the standard curve and
present in a table format.
In the experimental work performed, it was shown that
the Pure⋅SAL™ device removed 98.1–98.2 % of all DNA,
providing a total of 1.8–1.9 % of cfDNA in comparison to
the gold standard passive drool/centrifugation method
which was effective in removing 98.9–99.1 % of all DNA
and providing 0.9–1.1 % of cfDNA.


12

Paul D. Slowey

4  Notes
1.DNA from samples collected using one of the above commercial 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
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 miRNeasy 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 miRVana (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 particular 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 outlined in the manuscript [55]. In the experiments performed,
the Pure⋅SAL™ device was found to be a superior tool for harvesting 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 techniques, spin column methods, 96-well microplates, or automated methods, such as the Promega Maxwell 16 instrument
or the Qiagen QIAsymphony equipment. Whole saliva contains a significant quantity of mucinous material that can have
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 validation studies.

Salivary Diagnostics Using Purified Nucleic Acids

13

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, miRCURY 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 quantitative PCR (qPCR) as an alternate method of DNA assessment.

Acknowledgments
The author would like to acknowledge the support of Dr David T
Wong (UCLA) for his support and encouragement in preparing
this manuscript.
References
1.Punyadeera C, Slowey PD (2013) Saliva as an
emerging biofluid for clinical diagnosis and
applications of MEMS/NEMS in salivary diagnostics. Chapter 22 in “Nanobiomaterials in
Clinical Dentistry.” Elsevier, pp 453–475,
ISBN: 978-1-4557-3127-5
2.Streckfus C, Bigler L (2002) Saliva as a diagnostic fluid. Oral Dis 8:69–76
3.Slowey PD (2013) Commercial saliva collections tools. J Calif Dent 41(2):97–105
4. Malamud D, Tabak LA (eds) (1992) Saliva as a
diagnostic fluid. Vol 694, Annals NY Academy
of Science
5. Samaranayake LP (2006) Saliva tells the body’s
health. In: Stephen Moss (ed) The benefits of

chewing. pp 24–35
6. Zachary D, Mwenge L, Muyoyeta M, Shanaube
K, Schaap A, Bond V, Kosloff B, de Haas P,
Ayles HBMC (2012) Infect Dis 8(12):183
7.Testing Oral Fluid for the Presence of HIV
Antibodies February 2013. Report by the
Association of Public Health Laboratories
(APHL) />publications/Documents/ID_Feb2013_
Testing-­o f-­O ral-Fluid-for-the-Presence-of-­
HIV-Antibodies-Brief.pdf
8.See />tests/saliva
9.See />Home.aspx
10. See />Testing

11.Cone EJ, Huestis M (2007) Interpretation of
oral fluid tests for drugs of abuse. Ann NY
Acad Sci 1098:51–103
12.Bradshaw DJ, Marsh PD (1998) Analysis of
pH-driven disruption of oral microbial communities in vitro. Caries Res 32:456–462

13.Bratthall D, Hansel Petersson G (2005)
Cariogram-a multifactorial risk assessment
model for a multifactorial disease. Community
Dent Oral Epidemiol 33:256–264

14.Larmas M (1992) Saliva and dental caries:
diagnostic tests for normal dental practice. Int
Dent J 42:199–208

15.Christodoulides N, Floriano PN, Miller CS,

Ebersole JL, Mohanty S, Dharshan P, Griffin M,
Lennart A, Ballard KL, King CP Jr, Langub MC,
Kryscio RJ, Thomas MV, McDevitt JT (2007)
Lab-on-a-chip methods for point-of-­care measurements of salivary biomarkers of periodontitis. Ann NY Acad Sci 1098:411–428
16.Li Y, St John MA, Zhou X, Kim Y, Sinha U,
Jordan RC, Eisele D, Abemayor E, Elashoff D,
Park NH, Wong DT (2004) Salivary transcriptome diagnostics for oral cancer detection. Clin
Cancer Res 10:8442–8450
17.Streckfus C, Bigler L (2005) The use of soluble, salivary c-erbB-2 for the detection and
post-operative follow-up of breast cancer in
women: the results of a five-year translational
research study. Adv Dent Res 18:17–24
18.Streckfus C, Bigler L, Dellinger T, Dai X, Cox
WJ, McArthur A, Kingman A, Thigpen JT


14

Paul D. Slowey

(2001) Reliability assessment of soluble
c-erbB-2 concentrations in the saliva of healthy
women and men. Oral Surg Oral Med Oral
Pathol Oral Radiol Endod 91:174–179

19.Streckfus C, Bigler L, Dellinger T, Dai X,
Kingman A, Thigpen JT (2000) The presence
of soluble c-erbB-2 in saliva and serum among
women with breast carcinoma: a preliminary
study. Clin Cancer Res 6:2363–2370

20.Streckfus C, Bigler L, Dellinger T, Pfeifer M,
Rose A, Thigpen JT (1999) CA 15-3 and
c-erbB-2 presence in the saliva of women. Clin
Oral Investig 3:138–143
21.Streckfus C, Bigler L, Tucci M, Thigpen JT
(2000) A preliminary study of CA15-3,
c-erbB-2, epidermal growth factor receptor,
cathepsin-D, and p53 in saliva among women
with breast carcinoma. Cancer Invest
18:101–109
22.Streckfus C, Bigler L, Dellinger T, Kuhn M,
Chouinard N, Dai X (2004) The expression of
the c-erbB-2 receptor protein in glandular salivary secretions. J Oral Pathol Med
33:595–600

23.Ovchinnikov DA, Cooper MA, Pandit P,
Coman WB, Cooper-White JJ, Keith P,
Wolvetang EJ, Slowey PD, Punyadeera C
(2012) Tumor-suppressor gene promoter
hypermethylation in saliva of head and neck
cancer patients. Translat Oncol 5:321–326
24.Hu S, Zhou M, Jiang J, Wang J, Elashoff D,
Gorr S, Michie SA, Spijkervet FK, Bootsma H,
Kallenberg CG, Vissink A, Horvath S, Wong
DT (2009) Systems biology analysis of
Sjogren’s syndrome and mucosa-associated
lymphoid tissue lymphoma in parotid glands.
Arthritis Rheum 60:81–92

25.Corstjens LAM, Abrams WR, Malamud D

(2012) Detecting viruses by using salivary
diagnostics. J Am Dent Assoc 143:12S–18S
26.Shirtcliff EA, Buck RL, Laughlin MJ, Hart T,
Cole CR, Slowey PD (2015) Salivary cortisol
results obtainable within minutes of sample
collection correspond with traditional immunoassays. Clin Ther 37:505–514
27.Gallo A, Tandon M, Alevizos I, Illie GG (2012)
The majority of MicroRNAs detectable in serum
and saliva is concentrated in exosomes. PLoS
One 7, e30679,
28. See />MK-006.pdf
29.Thomas GA, Oberkanins C, Berndt A, Slowey
PD (2014) Validation of a series of genomic
StripAssays® to salivary DNA collection using
the DNA⋅SAL™ Device. Paper presented at the
American Society of Human Genetics Meeting,
San Diego, October 2014

30. See
/>saliva-dna-collection-preser vation-and-­­
isolation-kit
31. See />32. See
/>iswab-dna-collection-kit
33. El-Hahmawi B (2014) An efficient non-­invasive
sample collection technology for various population segments. Paper presented at the Qatar
Foundation Annual Research Conference, At
Doha, Qatar, Accessed 18 Nov 2014
34. See
/>isohelix-dna-buccal-swabs
35.Lee YH, Zhou H, Yan X, Zhang L, Chia D,

Wong DTW (2011) Direct saliva transcriptome
analysis. Clin Chem 57:1295–1302
36. See />
37.Patel RS, Jakymiw A, Yao B, Pauley BA,
Carcamo WC, Katz J, Cheng JQ, Chan EK
(2011) High resolution of microRNA signatures in human whole saliva. Arch Oral Biol
56:1506–1513
38. See
/>saliva-rna-collection-and-preservation-devices
39.Chiang SH, Thomas GA, Liao W, Grogan T,
Buck RL, Fuentes L, Yakob M, Laughlin MJ,
Schafer C, Nazmul-Hossain A, Wei F, Elashoff
D, Slowey PD, Wong DT (2015)
RNAPro*SAL: a device for rapid and standardized collection of saliva RNA and proteins.
Biotechniques 58:69–76
40. See
/>pure%E2%80%A2sal

41.Théry C, Zitvogel L, Amigorena S (2002)
Exosomes: composition, biogenesis and function. Nat Rev Immunol 2:569–579
42. Michael A, Bajracharya SD, Yuen PS, Zhou H,
Star RA, Illei GG, Alevizos I (2010) Exosomes
from human saliva as a source of microRNA
biomarkers. Oral Dis 16:34–38
43.Lau C, Kim Y, Chia D, Spielmann N, Eibl G,
Elashoff D, Wei F, Lin YL, Moro A, Grogan T,
Chiang S, Feinstein E, Schafer C, Farrell J,
Wong DT (2013) Role of pancreatic cancer-­
derived exosomes in salivary biomarker development. J Biol Chem 288:26888–26897
44.Lässer C, Alikhani VS, Ekström K, Eldh M,

Paredes PT, Bossios A, Sjöstran M, Gabrielsson
S, Lötvall J, Valadi H (2011) Human saliva,
plasma and breast milk exosomes contain RNA:
uptake by macrophages. J Translat Med 9:9
45.Laughlin MJ, Buck RL, Slowey PD (2014) A
simplified method for the isolation of exosomes
from saliva using a prototype saliva collection
device (RPSAL-702). Paper presented at the