Atlas of Oral Microbiology: From Healthy Microflora to Disease [2 ed.] 9789811578984, 9789811578991

Table of contents :
Preface
Contents
Editors and Contributors
Chapter 1: Basic Biology of Oral Microbes
1.1 Cytological Basis of Microorganisms
1.2 Microbial Morphology
1.2.1 Microbial Size
1.2.2 Microbial Morphology
1.3 Microbial Cell Structure
1.3.1 Basic Bacterial Structures
1.3.1.1 Cell Wall
1.3.1.2 Cell Wall-Deficient Bacteria (Bacterial L Form)
1.3.1.3 Cell Membrane
1.3.1.4 Cytoplasm
1.3.1.5 Nuclear Material
1.3.2 Special Bacterial Structures
1.3.2.1 Capsule
1.3.2.2 Flagellum
1.3.2.3 Pilus
1.3.2.4 Spore
1.3.3 Basic Virus Structure
1.4 Microbial Physiology
1.4.1 Binary Fission Reproduction
1.4.2 Bacterial Growth
1.5 Microbial Genetics
1.5.1 Heredity
1.5.2 Variation
1.5.3 Genetic Material of Bacteria
1.5.4 Mechanism of Microbial Variation
1.5.5 Gene Transfer and Recombination
1.5.5.1 Transformation
1.5.5.2 Conjugation
1.5.5.3 Transduction
References
Chapter 2: Techniques for Oral Microbiology
2.1 Smear and Stain Techniques
2.1.1 Kopeloff´s Modification of Gram Stain
2.1.1.1 Mechanism
2.1.1.2 Preparation of Gram Stain Reagents (Commercially Available)
2.1.1.3 Staining Procedure
2.1.1.4 Reporting Results
2.1.1.5 Precautions
2.1.2 Staining Bacterial Spores
2.1.2.1 Gram Stain
2.1.2.2 Fuchsin-Methylene Blue Stain
2.1.3 Staining the Bacterial Capsule
2.1.3.1 Murs Stain
2.1.3.2 Wright´s Stain
2.1.4 Flagellar Stain
2.1.4.1 Preparation of Staining Solutions
2.1.4.2 Smear Preparation
2.1.4.3 Staining Protocol
2.1.5 Staining Procedure for Special Fungal Structures
2.1.5.1 Crystal Violet Stain
2.1.5.2 Lactophenol Cotton Blue Stain
2.1.6 Giemsa Stain for Mycoplasma
2.1.7 Negative Congo Red Staining of Plaque Bacteria
2.1.8 Protozoan Smears
2.1.8.1 Wet Smear for Fresh Samples
2.1.8.2 Giemsa Staining
2.2 Isolation, Incubation, and Identification Techniques
2.2.1 Collection and Transportation of Samples
2.2.1.1 Sample Collection
2.2.1.2 Sample Transportation
2.2.2 Suspension and Dilution of Samples
2.2.2.1 Sample Suspension in Solution
2.2.2.2 Sample Dilution
2.2.3 Inoculation and Incubation of Samples
2.2.3.1 Choice of Medium
2.2.3.2 Sample Inoculation
2.2.3.3 Sample Incubation
2.2.4 Growth Characteristics and Identification
2.2.4.1 Growth Characteristics
2.2.4.2 Biochemical Tests
2.2.5 Instruments for Microbiological Identification
2.2.5.1 Spiral Plater
2.2.5.2 Microbiology Analyzer
2.2.5.3 Microbial Identification System
2.3 Microscopy Techniques
2.3.1 Stereomicroscopy
2.3.2 Scanning Electron Microscopy
2.3.2.1 Mechanism
2.3.2.2 Operating Procedure
2.3.2.3 Sample Observation
2.3.3 Transmission Electron Microscopy
2.3.3.1 Preparation of Bacterial Samples
2.3.3.2 Section
2.3.3.3 Uranium Staining and Observation
2.3.4 Confocal Laser Scanning Microscopy
2.3.4.1 Principles of CLSM
2.3.4.2 Application in Dental Plaque Research
2.4 Oral Microecology Techniques
2.4.1 Methods for Measuring Microbial Growth Curves
2.4.1.1 Protocol
2.4.1.2 Methods
2.4.2 Methods for Measuring Colony Forming Units
2.4.2.1 Protocol
2.4.2.2 Methods
2.4.3 Measurement of Adhesion Strength and Rate of Adhesion Inhibition
2.4.3.1 Measurement of Adhesion Strength
2.4.3.2 Measuring the Rate of Adhesion Inhibition
2.4.4 Techniques for the Detection of Plaque Biofilm
2.4.4.1 Biofilm Formation Assay
2.4.4.2 Biofilm Detection and Analysis
2.5 Oral Microbiome Techniques
2.5.1 Denaturing Gradient Gel Electrophoresis and Temperature Gradient Gel Electrophoresis
2.5.2 Sanger Sequencing
2.5.3 Next-Generation Sequencing
2.5.3.1 Pyrosequencing
2.5.3.2 Illumina Sequencing
References
Chapter 3: Supragingival Microbes
3.1 Gram-Positive Bacteria
3.1.1 Actinomyces
3.1.1.1 Actinomyces israelii
3.1.1.2 Actinomyces naeslundii
3.1.1.3 Actinomyces odontolyticus
3.1.1.4 Actinomyces viscosus
3.1.2 Bifidobacterium
3.1.2.1 Bifidobacterium dentium
3.1.2.2 Bifidobacterium breve
3.1.3 Lactobacillus
3.1.3.1 Lactobacillus acidophilus
3.1.3.2 Lactobacillus casei
3.1.3.3 Lactobacillus fermentum
3.1.4 Rothia
3.1.5 Staphylococcus
3.1.6 Streptococcus
3.1.6.1 Streptococcus salivarius
3.1.6.2 Streptococcus sanguinis
3.1.6.3 Streptococcus gordonii
3.1.6.4 Streptococcus mutans
3.1.6.5 Streptococcus sobrinus
3.1.6.6 Streptococcus oralis
3.1.6.7 α-Hemolytic Streptococcus
3.1.6.8 β-Hemolytic Streptococcus
3.2 Gram-Negative Bacteria
3.2.1 Leptotrichia
3.2.2 Veillonella
3.2.2.1 Veillonella parvula subsp. parvula
References
Chapter 4: Subgingival Microbes
4.1 Gram-Positive Bacteria
4.1.1 Enterococcus
4.1.2 Eubacterium
4.1.3 Peptostreptococcus
4.1.4 Propionibacterium
4.2 Gram-Negative Bacteria
4.2.1 Bacteroides
4.2.1.1 Bacteroides fragilis
4.2.1.2 Bacteroides thetaiotaomicron
4.2.2 Capnocytophaga
4.2.2.1 Capnocytophaga gingivalis
4.2.2.2 Capnocytophaga sputigena
4.2.3 Eikenella
4.2.4 Fusobacterium
4.2.4.1 Fusobacterium nucleatum
4.2.4.2 Fusobacterium necrophorum
4.2.4.3 Fusobacterium varium
4.2.5 Helicobacter
4.2.6 Aggregatibacter
4.2.7 Prevotella
4.2.7.1 Prevotella intermedia
4.2.7.2 Prevotella nigrescens
4.2.7.3 Prevotella melaninogenica
4.2.7.4 Prevotella corporis
4.2.7.5 Prevotella loescheii
4.2.8 Porphyromonas
4.2.8.1 Porphyromonas gingivalis
4.2.8.2 Porphyromonas endodontalis
4.2.9 Treponema
References
Chapter 5: Oral Mucosal Microbes
5.1 Gram-Positive Bacteria
5.1.1 Staphylococcus
5.1.2 Streptococcus
5.1.2.1 Streptococcus mitis
5.1.2.2 Streptococcus pyogenes
5.1.2.3 Streptococcus pneumoniae
5.1.2.4 Streptococcus vestibularis
5.2 Gram-Negative Bacteria
5.2.1 Escherichia
5.2.2 Haemophilus
5.2.3 Moraxella
5.2.4 Neisseria
5.3 Mycoplasma
5.4 Fungi
5.4.1 Saccharomyces
5.4.1.1 Candida albicans
5.5 Virus
References
Chapter 6: New Oral Microbial Isolations
6.1 Gram-Positive Bacteria
6.1.1 Leuconostoc lactis
6.1.2 Corynebacterium argentoratense
6.2 Gram-Negative Bacteria
6.2.1 Stenotrophomonas maltophilia
6.2.2 Chryseobacterium indologenes
6.2.3 Elizabethkingia anophelis
6.2.4 Klebsiella pneumoniae
6.2.5 Escherichia coli
6.2.6 Campylobacter jejuni
6.2.7 Veillonella atypica
6.3 Fungi
6.3.1 Candida tropicalis
6.3.2 Candida glabrata
6.3.3 Candida parapsilosis
6.3.4 Candida krusei
6.3.5 Aspergillus fumigatus
References
Chapter 7: The Oral Microbiome Bank of China
7.1 Introduction
7.2 Results
7.2.1 Overview of the Online Database
7.2.1.1 Web-Accessible Functions
7.3 Discussion
7.4 Materials and Methods
7.4.1 Collection and Transport of Samples
7.4.2 Dispersion and Dilution of Samples
7.4.3 Inoculation and Incubation of Samples
7.4.4 Smear and Stain
7.4.5 Growth Characteristics and Identification
7.4.6 Biochemical Tests
7.4.7 Molecular Method
7.4.8 DNA Extraction and Sequencing
7.4.9 16S rRNA Alignment
7.4.10 Database and Web Design
7.4.11 Curation
7.4.12 Service and Function
References
Chapter 8: Invasion of Oral Microbiota into the Gut
8.1 Introduction
8.2 Results
8.2.1 The Oral Microbiota of the HOMA Mouse Model
8.2.2 Biogeography of the Host Gut-Selected Oral Microbiota
8.2.3 Ecological Invasion by Oral Microbiota in the Gut
8.2.4 Porphyromonas Competed for Colonization with the Small Intestinal Microbiota
8.3 Discussion
8.4 Materials and Methods
8.4.1 Sample Collection from Humans
8.4.2 Animal Husbandry
8.4.3 Establishment of the HOMA and HMA Mouse Models
8.4.4 Cohousing Experiment
8.4.5 16S rRNA Gene Sequencing
8.4.6 Bioinformatics and Statistical Analysis
References
Chapter 9: Mycobiome Dysbiosis in Oral Lichen Planus
9.1 Introduction
9.2 Results
9.2.1 Participant Demographics and Sequence Data
9.2.2 Lower Saliva Biodiversity of the Mycobiome and Higher Biodiversity of the Bacteriome in OLP
9.2.3 Taxonomic Differences Among Healthy Individuals and OLP Patients
9.2.4 Inversion of Myco-Bacteriome Co-occurrence Patterns from Antagonization to Co-prosperity
9.2.5 Distinct Network Topology Between OLP and Healthy Individuals
9.2.6 Fungal Disturbance Promotes OLP Exacerbation
9.3 Discussion
9.4 Materials and Methods
9.4.1 Subject Recruitment and Sample Collection
9.4.2 Cytokine Assay
9.4.3 DNA Extraction
9.4.4 Illumina Sequencing
9.4.5 Data Preprocessing, OTU Clustering, and Taxonomic Classification
9.4.6 Statistical Analysis
9.5 Data Availability
9.6 Ethics Statement
References
Chapter 10: Intestinal Microbiota and Osteoporosis
10.1 Introduction
10.1.1 The Intestinal Microbiota and Its Regulators
10.1.2 The Intestinal Microbiota Regulates Bone Metabolism
10.1.2.1 Involvement of the Intestinal Microbiota in Bone Metabolism
10.1.2.2 Mechanisms by Which the Gut Microbiota Regulates Bone Metabolism
10.1.3 Relationship Between the Intestinal Microbiota and PMO
10.1.3.1 PMO Animal Models
10.1.3.2 PMO Development Depends on the Intestinal Microbiota and Host Genetic Background
10.1.3.3 Probiotics Prevent Bone Loss in PMO Murine Models
10.1.4 Host and Microbiota Interactions in the Pathogenesis and Treatment of PMO
10.1.4.1 Intestinal Microbial Diversity in PMO Is Regulated by Estrogen and Probiotics
10.1.4.2 Intestinal Epithelial Barrier Function in PMO Is Regulated by Estrogen and Probiotics
10.1.4.3 Host Immune Responses in PMO Are Regulated by Estrogen and the Intestinal Microbiota
10.1.4.4 The Intestinal Microbiota and Estrogen Orchestrate Calcium Absorption
10.1.4.5 The Gut Microbiota Produces Estrogen-Like Metabolites with Regulatory Effects on Bone Metabolism
10.1.5 Conclusion
References
Index

Citation preview

Xuedong Zhou Yuqing Li  Editors

Atlas of Oral Microbiology: From Healthy Microflora to Disease Second Edition

Atlas of Oral Microbiology: From Healthy Microflora to Disease

Xuedong Zhou • Yuqing Li Editors

Atlas of Oral Microbiology: From Healthy Microflora to Disease Second Edition

Editors Xuedong Zhou State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology Sichuan University Chengdu, Sichuan, China

Yuqing Li State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology Sichuan University Chengdu, Sichuan, China

Associate Editors Xian Peng State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology Sichuan University Chengdu, Sichuan, China

Biao Ren State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology Sichuan University Chengdu, Sichuan, China

ISBN 978-981-15-7898-4 ISBN 978-981-15-7899-1 https://doi.org/10.1007/978-981-15-7899-1

(eBook)

Jointly published with Zhejiang University Press # Zhejiang University Press 2020 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, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Singapore Pte Ltd. The registered company address is: 152 Beach Road, #21-01/04 Gateway East, Singapore 189721, Singapore

Preface

The human body is home to a diverse range of microorganisms, including a variety of bacteria, archaea, fungi, and viruses, estimated to outnumber human cells in a healthy adult by tenfold. The importance of characterizing human microbiota for understanding health and disease is highlighted by the recent launch of the Human Microbiome Project (HMP) by the National Institutes of Health. Changes in colonization sites or imbalance of normal microflora may lead to the occurrence of disease. The mouth harbors a diverse, abundant, and complex microbial community, and oral microorganisms are often closely related to many infectious diseases. This book is keeping up with the advanced edge of the international research field of microbiology. For the first time, it innovatively gives us a complete description of the oral microbial systems according to different oral ecosystems. This book collects a large number of oral microbial pictures, including color pictures, colonies photos, and electron microscopy photos. It is by far the most abundant oral microbiology atlas, containing the largest number of pictures. In the meantime, it also describes in detail a variety of experimental techniques, including microbiological isolation, culture, and identification. It is a monograph with strong practical function. This book is the first professional and comprehensive color atlas of oral microbiology and it has important academic and application value. The editors and writers have long been engaged in teaching and research work in oral microbiology and oral microecology; they had completed the first color atlas of the oral microbial morphology and ultrastructure, which was written in Chinese. They all have solid theoretical foundation and rich experience in oral microbiological research. This book deserves a broad audience, and it will meet the needs of researchers, clinicians, teachers, and students major in biology, stomatology, basic medicine, or clinical medicine. It can also be used to facilitate teaching and international academic exchanges. Chengdu, Sichuan, China

Xuedong Zhou Yuqing Li

v

Contents

1

Basic Biology of Oral Microbes . . . . . . . . . . . . . . . . . . . . . . . Yuqing Li, Xian Peng, Xuedong Zhou, Biao Ren, Liying Xiao, Yan Li, Mingyun Li, and Qiang Guo

1

2

Techniques for Oral Microbiology . . . . . . . . . . . . . . . . . . . . . Xian Peng, Biao Ren, Yuqing Li, Xuedong Zhou, Jing Xie, Chenchen Zhou, Demao Zhang, Xin Zheng, and Xinxuan Zhou

25

3

Supragingival Microbes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Xuedong Zhou, Yuqing Li, Xian Peng, Biao Ren, Jiyao Li, Xin Xu, Jinzhi He, and Lei Cheng

81

4

Subgingival Microbes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 145 Quan Yuan, Chenchen Zhou, Jing Xie, Demao Zhang, Liwei Zheng, Yuqing Li, Biao Ren, Xian Peng, and Xuedong Zhou

5

Oral Mucosal Microbes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211 Biao Ren, Lei Cheng, Xian Peng, Yuqing Li, Yan Li, Yujie Zhou, Chengguang Zhu, and Xi Chen

6

New Oral Microbial Isolations . . . . . . . . . . . . . . . . . . . . . . . . 253 Yuqing Li, Xian Peng, Biao Ren, Boyu Tang, Tao Gong, Zhengyi Li, and Xuedong Zhou

7

The Oral Microbiome Bank of China . . . . . . . . . . . . . . . . . . . 287 Xian Peng, Xuedong Zhou, Xin Xu, Yuqing Li, Yan Li, Jiyao Li, Xiaoquan Su, Shi Huang, Jian Xu, and Ga Liao

8

Invasion of Oral Microbiota into the Gut . . . . . . . . . . . . . . . . 301 Bolei Li, Yang Ge, Lei Cheng, Benhua Zeng, Jinzhao Yu, Xian Peng, Jianhua Zhao, Wenxia Li, Biao Ren, Mingyun Li, Hong Wei, and Xuedong Zhou

9

Mycobiome Dysbiosis in Oral Lichen Planus . . . . . . . . . . . . . 315 Yan Li, Kun Wang, Bo Zhang, Qichao Tu, Yufei Yao, Bomiao Cui, Biao Ren, Jinzhi He, Xin Shen, Joy D. VanNostrand, Jizhong Zhou, Wenyuan Shi, Liying Xiao, Changqing Lu, and Xuedong Zhou

vii

viii

10

Contents

Intestinal Microbiota and Osteoporosis . . . . . . . . . . . . . . . . . 333 Xin Xu, Xiaoyue Jia, Longyi Mo, Chengcheng Liu, Liwei Zheng, Quan Yuan, and Xuedong Zhou

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 359

Editors and Contributors

Associate Editors Xian Peng State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, Sichuan, China Biao Ren State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, Sichuan, China

Contributors Lei Cheng State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China Xi Chen State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Bomiao Cui State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Yang Ge State Key Laboratory of Oral Diseases, Sichuan University, Chengdu, China National Clinical Research Center for Oral Diseases, Sichuan University, Chengdu, China Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China Tao Gong State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

ix

x

Qiang Guo State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Jinzhi He State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Shi Huang Single-Cell Center, CAS Key Laboratory of Biofuels and Shandong Key Laboratory of Energy Genetics, Qingdao Institute of BioEnergy and Bioprocess Technology, Chinese Academy of Sciences, Qingdao, Shandong, China Xiaoyue Jia State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Department of Operative Dentistry and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China Ga Liao State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Bolei Li State Key Laboratory of Oral Diseases, Sichuan University, Chengdu, China National Clinical Research Center for Oral Diseases, Sichuan University, Chengdu, China Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China Jiyao Li State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China Mingyun Li State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Chengcheng Liu State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Department of Periodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China Wenxia Li Department of Laboratory Animal Science, College of Basic Medical Sciences, Third Military Medical University, Chongqing, China Yan Li State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

Editors and Contributors

Editors and Contributors

xi

Yuqing Li State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Zhengyi Li State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Changqing Lu Department of Anatomy, West China School of Basic Medical and Forensic Medicine, Sichuan University, Chengdu, China Longyi Mo State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Xin Shen State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Wenyuan Shi The Forsyth Institute, Cambridge, MA, USA Xiaoquan Su Single-Cell Center, CAS Key Laboratory of Biofuels and Shandong Key Laboratory of Energy Genetics, Qingdao Institute of BioEnergy and Bioprocess Technology, Chinese Academy of Sciences, Qingdao, Shandong, China Boyu Tang State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Qichao Tu Institute of Marine Science and Technology, Shandong University, Qingdao, China Institute for Environmental Genomics, Department of Microbiology and Plant Biology, University of Oklahoma, Norman, OK, USA Joy D. VanNostrand Institute for Environmental Genomics, Department of Microbiology and Plant Biology, University of Oklahoma, Norman, OK, USA Kun Wang State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Hong Wei Department of Laboratory Animal Science, College of Basic Medical Sciences, Third Military Medical University, Chongqing, China Liying Xiao State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Jing Xie State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

xii

Jian Xu Single-Cell Center, CAS Key Laboratory of Biofuels and Shandong Key Laboratory of Energy Genetics, Qingdao Institute of BioEnergy and Bioprocess Technology, Chinese Academy of Sciences, Qingdao, Shandong, China Xin Xu State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Department of Operative Dentistry and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China Yufei Yao State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Quan Yuan Department of Dental Implantology, West China Hospital of Stomatology, Sichuan University, Chengdu, China State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Jinzhao Yu State Key Laboratory of Oral Diseases, Sichuan University, Chengdu, China National Clinical Research Center for Oral Diseases, Sichuan University, Chengdu, China Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China Benhua Zeng Department of Laboratory Animal Science, College of Basic Medical Sciences, Third Military Medical University, Chongqing, China Bo Zhang State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Demao Zhang State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Jianhua Zhao Shanghai Majorbio Bio-pharm Technology Co., Ltd, Shanghai, China Liwei Zheng Department of Pediatric Dentistry, West China Hospital of Stomatology, Sichuan University, Chengdu, China State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Xin Zheng State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Chenchen Zhou State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

Editors and Contributors

Editors and Contributors

xiii

Jizhong Zhou Institute for Environmental Genomics, Department of Microbiology and Plant Biology, University of Oklahoma, Norman, OK, USA Xinxuan Zhou State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Xuedong Zhou State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China Yujie Zhou State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Chengguang Zhu State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

1

Basic Biology of Oral Microbes Yuqing Li, Xian Peng, Xuedong Zhou, Biao Ren, Liying Xiao, Yan Li, Mingyun Li, and Qiang Guo

Abstract

morphology, microbial cell structure, microbial physiology, and microbial genetics.

The cell is the fundamental unit of all living organisms. Various subunit structures and chemical substances found on and inside the cell make complex cellular functions possible. Understanding the morphology and structures of microorganisms is of great significance to study their physiological activities and pathogenicity, to identify different microorganisms, and to diagnose and prevent diseases. The morphological structure, physiological metabolism, pathogenicity, and drug resistance of microorganisms are determined by genes. The heredity of microorganism guarantees the stability of species, while the variation produces the variety or new species, which is beneficial to the evolution of species. The main contents of this chapter include cytological basis of microorganisms, microbial

1.1

Y. Li (*) · X. Peng · B. Ren · L. Xiao · Y. Li · M. Li · Q. Guo State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China e-mail: [email protected]

1.2

X. Zhou State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China

Keywords

Microbial morphology · Microbial cell structure · Microbial physiology · Microbial genetics

Cytological Basis of Microorganisms

The cell is the fundamental unit of all living organisms. Various subunit structures and chemical substances found on and inside the cell make complex cellular functions possible. Microbes can be divided into three major groups according to their morphological structure, degree of differentiation, and chemical composition: eukaryote, prokaryote, and acellular microorganisms (Figs. 1.1, 1.2, 1.3, and 1.4).

Microbial Morphology

Microorganisms, also known as microbes, are tiny organisms that are only visible under an optical microscope or an electron microscope. They are small in size and simple in structure. Microbes reproduce quickly, can tolerate a wide range of environmental conditions, are widely distributed and highly variable, and tend to congregate.

# Zhejiang University Press 2020 X. Zhou, Y. Li (eds.), Atlas of Oral Microbiology: From Healthy Microflora to Disease, https://doi.org/10.1007/978-981-15-7899-1_1

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Fig. 1.1 Eukaryotic microbes (Saccharomyces, SEM) The eukaryotic cell has a high degree of nuclear differentiation; it has a nuclear membrane, nucleoli, and chromosomes. There is a complete complement of structured organelles in the cytoplasm, and cellular reproduction takes place by mitosis. Examples include fungi and algae

Fig. 1.2 Prokaryotic microbes (bacteria)

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Fig. 1.3 Prokaryotic microbes (Mycoplasma) The prokaryotic cell has a primitive nucleoplasm and cell membrane; it has no nuclear membrane, nucleolus, or organelles. Prokaryotes include bacteria such as Mycoplasma, Chlamydia, and Rickettsia

Fig. 1.4 Acellular microorganisms (herpes simplex virus) Acellular microorganisms are the smallest microorganism with no typical cell structure and no enzymatic energyproduction system. They consist merely of a nucleic acid genome (DNA/RNA) and a protein coat (the capsid). Acellular microorganisms can only reproduce inside a living cell. Examples include viruses and subviral agents

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Fig. 1.5 Microbial size 1. Staphylococcus 1000 nm 2. Rickettsia 450 nm 3. Chlamydia 390 nm 4. Tobacco mosaic virus 300  15 nm 5. Escherichia coli bacteriophages, head 65  95 nm, tail 12  100 nm 6. Adenovirus 70 nm 7. Poxvirus 300  230 nm 8. Influenza virus 100 nm 9. Poliovirus 30 nm 10. Japanese encephalitis virus 40 nm 11. Molecule of egg protein 10 nm

1.2.1

Microbial Size

As many types of microbes exist, they vary widely in their size. Generally, the units used to measure microbes are μm and nm. Most cocci are 1 μm in diameter. Bacilli can be further divided into coccobacillus, brevibacterium, and long bacilli and measure approximately 1–10 μm in length and 0.3–1 μm in width. Spirochetes measure approximately 6–20 μm in length and 0.1–0.2 μm in width. Fungi are several times larger than bacteria. Most viruses are smaller than 150 nm and only visible under the electron microscope. The same microbe can change in size depending on their environment or age (Fig. 1.5).

1.2.2

Microbial Morphology

Different types of microbes have different, but characteristic shapes. Under suitable conditions, the shape and size of microbes are relatively stable. It is important to know the morphological

structure of microbes, as it provides us with a better understanding of microbial physiology, pathogenic mechanisms, and antigenic features and allows us to identify them by species. Also, knowledge of microbial morphology can be helpful in diagnosing disease and in preventing microbial infections. 1. Bacteria Bacteria are complex and highly variable microbes. They come in four basic shapes: spherical (cocci), rod-shaped (bacilli), arc-shaped (vibrio), and spiral (spirochete) (Fig. 1.6). 2. Fungi Fungi are divided into unicellular and multicellular according to the number of cells that make up the organism. Unicellular fungi, such as Saccharomyces and other yeast-like fungi, are usually round or oval. Multicellular fungi have hyphae and spores. The hypha and spores of different fungi are shaped differently (Figs. 1.7 and 1.8). 3. Virus Many viruses are spherical or almost spherical; some are rod-shaped (often seen in plant viruses), filamentous (e.g., freshly isolated influenza virus), bullet-shaped (e.g.,

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Fig. 1.6 Basic shape of bacteria

Fig. 1.7 Fungal spores

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Fig. 1.8 Fungal hyphae

Fig. 1.9 Morphology and structure of viruses 1. Poxvirus, 2. paramyxovirus, 3. orthomyxovirus, 4. coronavirus, 5. Togaviridae, 6. adenovirus, 7. bullet-shaped

virus, 8. herpes virus, 9. T2 bacteriophage, 10. reovirus, 11. papovavirus, 12. picornavirus, 13. picodnavirus, 14. tobacco mosaic virus

rabies virus), brick-shaped (e.g., poxvirus), and tadpole-shaped (e.g., bacteriophage) (Fig. 1.9).

1.3.1.1 Cell Wall The cell wall is the outermost structure of the bacterial cell and is located outside the cell membrane. It is transparent, tough, and flexible. The average thickness ranges from 15 to 30 nm. It mainly consists of peptidoglycan, which is also called mucopeptide, glycopeptides, or murein. Bacteria are classified into Gram-positive and Gram-negative based on the appearance of the cells after Gram stain. The peptidoglycan of Gram-positive bacteria is composed of a glycan backbone, tetrapeptide side chains, and a pentapeptide cross-linking bridge (Fig. 1.11). The peptidoglycan of Gram-negative bacteria is composed of a glycan backbone and a tetrapeptide side chain (Fig. 1.12). Gram-positive and Gram-negative bacteria have unique structures other than peptidoglycan in their cell walls (Figs. 1.13 and 1.14). Other substances, such as compound polysaccharide, surface protein, protein M and G of

1.3

Microbial Cell Structure

Although different microbes possess different cellular structures, there are certain commonalities within groups of microbes.

1.3.1

Basic Bacterial Structures

The architecture of bacterial cells consists of basic and special structures. Basic structures include the cell wall, cell membrane, cytoplasm, nuclear material, ribosome, plasmid, etc. Special structures, which are only found in some bacteria, include the flagellum, pilus, capsule, spore, etc. (Fig. 1.10).

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Fig. 1.10 Schematic representation of bacterial cell structure

Fig. 1.11 Schematic representation of Staphylococcus aureus cell wall peptidoglycan M, N-acetylmuramic acid; G, N-acetylglucosamine; O, β-1,4 glycosidic bond; a, L-alanine; b, D-aspartic acid; c, L-lysine; d, D-aspartic acid; x, glycine

Streptococcus, protein A of Staphylococcus aureus, etc., are found on the outer layer of the cell wall of some Gram-positive bacteria.

1.3.1.2

Cell Wall-Deficient Bacteria (Bacterial L Form) Cell wall-deficient bacteria are strains of bacteria that lack cell walls. The peptidoglycan that makes up the cell wall can be destroyed or inhibited by

physical, chemical, or biological factors. When Gram-positive bacteria lack a cell wall, the cytoplasm is surrounded by the cell membrane, and the entire structure is known as a protoplast. When Gram-negative bacteria do not have a cell wall, the cytoplasm is protected by the outer membrane, and the entire structure is called a spheroplast. Bacteria that have lost their cell wall are still capable of growing and dividing as

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Fig. 1.12 Schematic representation of Escherichia coli cell wall peptidoglycan M, N-acetylmuramic acid; G, N-acetylglucosamine; Ala, Alanine; Glu, glutamic acid; DAP, diaminopimelic acid

Fig. 1.13 Schematic of Gram-positive bacteria cell wall Gram-positive bacteria have a thick (20–80 nm) cell wall composed of 15–50 layers of peptidoglycan, many teichoic acids, and some teichuroic acid. Teichoic acids are unique to Gram-positive bacterial cell walls and constitute a class of important antigens related to the serotype classification of certain bacterial species. Teichoic acid accounts for about 50% of the dry weight of the cell

wall. It is a polymer consisting of ribitol or glycerin residues that are bound by phosphodiester bonds into a long-chain, which is then anchored in peptidoglycan. Teichoic acids are classified into cell wall teichoic acids and membrane teichoic acids (also known as lipoteichoicacid, LTA) according to the cellular structure to which they are anchored

cell wall-deficient bacteria. Examples of these were first isolated in 1935 by Emmy Klieneberger-Nobel, who named them “L-forms” after the Lister Institute in London where she was working at the time. L-form

bacteria give rise to a variety of cell morphologies and sizes and can be spherical, rod-shaped, filiform, etc [1]. The rate of growth and division of L-form bacteria is slow. They also form distinctive bacterial colonies when plated on agar. Some

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Fig. 1.14 Schematic representation of the cell wall of Gram-negative bacteria Gram-negative bacteria have a comparatively thin cell wall, approximately 10–15 nm in thickness. It is made up of 1–2 layers of peptidoglycan and other complex structures. On the outside of the peptidoglycan is the outer membrane, which is the main component of the

Gram-negative bacteria cell wall and accounts for approximately 80% of the dry weight of the cell wall. The outer membrane is composed of three layers: lipoprotein, lipid bilayer, and lipopolysaccharide (LPS) ordered from the interior to the exterior. OMP outer membrane protein, PP porin, BP nutrient binding protein, CP carrier protein, M N-acetylmuramic acid, G N-acetylglucosamine

L-form strains have a tendency to revert to the normal phenotype when the conditions that were used to produce the cell wall deficiency are reduced. L-from bacteria are difficult to stain or stain unevenly. In a Gram stain test, L-form bacteria always show up as Gram-negative, due to the lack of a cell wall.

The cell membrane of some bacteria can form invaginations into the cytoplasm called mesosomes.

1.3.1.3 Cell Membrane The cell membrane is a selectively permeable biological membrane found inside the cell wall and surrounding the cytoplasm. It is made of a lipid bilayer. The cell membrane is compact and flexible and measures approximately 7.5 nm in thickness. It accounts for 10–30% of the bacterial cell dry weight. The structure of the bacterial cell membrane resembles that of eukaryotic cell membranes, except it is deficient in cholesterol. The lipid bilayer is embedded with carrier proteins and zymoprotein, which possess specific functions.

1.3.1.4 Cytoplasm The cytoplasm is the gel-like substance enclosed within the cell membrane, which is made up of water, protein, lipid, nucleic acid, inorganic salts, etc. Most metabolic activities take place within the cytoplasm, and subcellular structures, such as ribosomes, plasmids, cytoplasmic granules, and others, are located in the cytoplasm. Ribosomes are found in cytoplasm. They are approximately 15–20 nm in diameter and are composed of a small (30S) and a large (50S) subunit. The association between subunits requires the presence of Mg2+. Ribosomes are made up of 30% ribosomal proteins and 70% ribosomal RNA. Plasmids are small, circular, double-stranded DNA molecules and are extrachromosomal genetic material. They can replicate

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Fig. 1.15 Capsule of S. pneumoniae (Murs staining method) Bacterial cells are stained red and the capsule around the cell appears as blue transparent circles

independently of chromosomal DNA and transmit genes encoding drug resistance, bacteriocins, and toxins and more from one bacterium to another via conjugation and transduction. Cytoplasmic granule is a general term referring to many types of cytoplasmic inclusion granules. They are an intracytoplasmic (inside the cytoplasm of a cell) form of storing nutrients and energy and include molecules such as polysaccharides, lipids, phosphate, etc. They are not essential or permanent structures in cells. Cytoplasmic granules are also known as metachromatic granules because they may stain into different colors than other bacterial cell structures.

1.3.1.5 Nuclear Material The bacterial nuclear material is also called the nucleoid. It is a piece of double-stranded DNA devoid of nuclear membrane, nucleolus, or histones and is the bacterial equivalent of chromatin. The function of the nucleoid is similar to that of the nucleus in eukaryotic cells and encodes genes necessary for activities and traits such as growth, metabolism, reproduction, heredity, mutation, etc.

1.3.2

Special Bacterial Structures

1.3.2.1 Capsule The capsule is a layer of slime that lies outside the bacterial cell wall. It is secreted by bacteria and diffuses into the surrounding medium. Based on its appearance when examined by light microscope, the bacterial capsule is classified into two types: micro-capsule, which is less than 0.2 μm in thickness and escapes optical detection, and capsule or large capsule, which is over 0.2 μm in thickness, binds tightly to the cell wall, and presents an obvious boundary under optical microscope. The capsule shows up as negatively stained when ordinary staining techniques are used. It appears as a clear halo around the bacterium when stained samples are examined by light microscope. Using special staining, the capsule can be stained differently from the bacterial cell (Fig. 1.15). Most bacterial capsules are composed of polysaccharides, but a few capsules are made of polypeptides. Capsular polysaccharides are highly hydrated molecules in which water accounts for more than 95% of the composition. They bind to

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Fig. 1.16 Schematic of Escherichia coli flagellum (1) Basal body: Located at the base of the flagellum. The basal body, embedded in the cell wall and cell membrane, is the output device. It acts as an engine to provide energy for locomotion. The nearby switch determines the direction of rotation (2) Hook: This structure points directly away from the cell

Fig. 1.17 Examples of bacterial flagellar arrangement

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and has a sharp bend (about 90 ) from which filaments protrude (3) Filament: This filiform structure protrudes from the bacterial cell. It is a hollow tube made of the protein flagellin. It acts like a ship or plane’s propeller to move the bacterial cell

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Fig. 1.18 Periplasmic flagella (flagella staining) The bacterial cell is stained red, and the flagellum is stained light red around the bacterial cell

phospholipids or lipid A on the cell surface through covalent bonds. The capsule is considered as an important virulence factor because it protects bacteria from engulfment by eukaryotic immune cells and desiccation and helps bacteria adhere to surfaces.

1.3.2.2 Flagellum The flagellum is a lash-like appendage that protrudes from the cell body and usually measures 5–20 μm in length and 10–30 nm in diameter. It is the locomotive organelle of motile bacteria such as Selenomonas and Wolinella succinogenes. The flagellum is composed of three parts: basal body, hook, and filament (Fig. 1.16). Different bacteria can have anywhere from one or two flagella to hundreds of flagella (Fig. 1.17). It only can be observed directly by electronic microscope or by light microscope after special staining (Fig. 1.18). The flagellum is involved in the pathogenesis of some diseases and is antigenic (e.g., antigen H). Examples of flagellate bacteria include Vibrio cholerae and Campylobacter jejuni, which use multiple flagella to propel themselves through the mucus lining of

the small intestine to reach the epithelium and produce toxin. Flagella can be classified as monotrichous, amphitrichous, lophotrichous, and peritrichous according to their number and location.

1.3.2.3 Pilus The pilus is a hair-like structure associated with bacterial adhesion and related to bacterial colonization and infection. Pili are primarily composed of oligomeric pilin proteins, which arrange helically to form a cylinder. New pilin protein molecules insert into the base of pilus. Pili are antigenic, and genes encoding pili can be located in the bacterial chromosome or in plasmids. Pili are not locomotive structures. They are classified into ordinary pilus or sex pilus according to their morphology, distribution, and function. The pilus is found on the surface of many Gram-positive bacteria and some Gram-negative bacteria. It is thinner and shorter than the flagellum. Ordinary pili are 0.3–1.0 μm in length and about 7 nm in diameter and are distributed all over the bacterial cell surface. Sex pili can be found in a handful of Gram-negative bacteria.

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Fig. 1.19 Schematic representation of the bacterial spore

These pili are longer and thicker than ordinary pili, and each bacterial cell can have from 1 to 4 sex pili.

1.3.2.4 Spore The spore is a small round or oval body that forms in bacteria due to cytoplasmic dehydration under unfavorable conditions (Fig. 1.19). It is surrounded by multiple membrane layers and has low permeability. Only Gram-positive bacteria can form spores, including species such as Bacillus subtilis, Clostridium tetani (Fig. 1.20), etc. The spore contains a complete karyoplasm and enzymatic system and can maintain all the essential activities for the bacteria to remain alive. The multiple membrane layers of the spore are, from the exterior to the interior, as follows: spore coating, spore shell, outer membrane, cortex, cell wall of spore, and inner membrane, which surrounds the nucleus of the spore. Spores are difficult to stain due to their thick cell wall. Special staining is required to stain the spore and distinguish it from the bacterial cell (Fig. 1.20). The size, morphology, and location of the spore differ between bacterial species and can be used to help identify bacteria (Fig. 1.21). For example, the Clostridium tetani spore is round and larger than the transverse diameter of the

bacterial cell, forming a drumstick-like structure, as the spore is located at the tip of the bacterial cell (Fig. 1.20).

1.3.3

Basic Virus Structure

Viruses are a kind of acellular microbe consisting mainly of nucleic acid and proteins. Some viruses are composed of a small amount of lipids and polysaccharides. The basic structure of virus is made up by the viral core, viral capsid, as well as a membrane envelope in some viruses (Fig. 1.22). The size, morphology, and structure of viruses play important roles in viral taxonomy and in diagnosing viral infections. (1) Viral core: namely, the nucleic acid component, which makes up the genome of the virus. The viral core provides genetic information that determines pathogenicity, antigenicity, proliferation, heredity, variation, etc. The chemical components of the viral core are DNA or RNA, based on whether the virus is classified as a DNA virus or a RNA virus. Nucleic acid can be single or double stranded. The relative molecular mass of the viral core is 2–160  106.

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Fig. 1.20 Spore stain of Clostridium tetani (fuchsinmethylene blue stain)

Fig. 1.21 Size, morphology, and location of bacterial spores

(2) Viral capsid: it is a protein shell that surrounds and protects the nucleic acid of the virus. The viral capsid is sometimes associated with the viral nucleic acid, and this structure is known as the nucleocapsid. In virions without an envelope, the nucleocapsid makes up the entirety of the virus. The viral capsid is composed of repeated protein subunits known as capsomeres, which is made of one or more proteins known as the chemical subunit or structural subunit. (3) Envelope: this is the one or two layers of membrane that surround the capsid of some viruses, which is a structure unique to the class of viruses known as enveloped viruses. The envelope is formed during the maturation

process when certain viruses bud out from the cell membrane. Therefore, the envelope can be composed of the host cell membrane and/or the nuclear membrane. The surface of some viral envelopes carries protein protrusions called as peplomers or spikes.

1.4

Microbial Physiology

Bacterial cells are synthesis machines that multiply themselves. The growth and division of bacteria include approximately 2000 different types of biochemical reactions that mediate energy conversion or enzymatic biosynthesis.

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collected information, it is possible to generate a growth curve using culture time as the horizontal axis and the logarithmic number of viable cells in the culture as the vertical axis (Fig. 1.26). Growth curves can generally be divided into four major sections.

Fig. 1.22 Basic structure of a virus

1.4.1

Binary Fission Reproduction

Binary fission is a process by which many prokaryotes reproduce from a single cell into two new cells (Fig. 1.23). Bacteria reproduce asexually by binary fission. Cocci can divide from different planes to form different arrangements. Bacilli divide along their horizontal axis; however, some bacterial species such as Mycobacterium tuberculosis occasionally spilt by branching. During cell division, the cell volume increases and a diaphragm is generated where the cell division is to take place. Then, a single cell will divide into two cells (Figs. 1.24 and 1.25). Under suitable conditions, the majority of bacteria divide quickly, about 20–30 min for one division. However, the growth of some bacterial species can be relatively slow. For example, Mycobacterium tuberculosis takes 18–20 h to complete one round of division.

1.4.2

Bacterial Growth

Mastering the fundamentals of bacterial growth allows the researcher to change culture conditions artificially, adjust the bacterial phases of growth and reproduction, and use beneficial bacteria more efficiently. Bacterial growth involves inoculating a certain number of bacteria into a suitable liquid medium and checking the number of viable cells at different time intervals. With the

1. Lag phase: During this process, the bacteria are adapting to their new environment. The volume of bacterial cells increases and their metabolism is active, but cell division is slow and reproduction is minimal. 2. Logarithmic phase: In this period, bacteria grow rapidly, and they divide and reproduce at a constant speed. The number of bacteria increases exponentially and the number of viable cells increases logarithmically. 3. Stationary phase: The bacterial growth rate gradually decreases, and the number of dead bacteria increases. The number of newly produced bacteria is approximately equal to the number of dying bacteria, and the number of viable cells remains relatively stable. 4. Decline phase: Bacterial growth rate slows and stops, and the number of dead bacteria is higher than that of viable cells. Cells become polymorphic, showing morphologies such as cell deformation, swelling, or autolysis.

1.5

Microbial Genetics

Like the other living organisms, microorganisms have heredity and variable characteristics. Heredity keeps microbial genetic traits relatively stable to ensure the reproduction of the species, while variations produce changes in the microorganisms that are useful for microbial survival and evolution and ultimately lead to the generation of new species.

1.5.1

Heredity

Heredity is the similarity in biological traits between offspring and its parent. Cells can be considered as a chemical plant in which

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Fig. 1.23 Binary fission in bacilli

information can be stored and transformed into a useful product. Enzymes are the molecular machines that catalyze specific chemical reactions. The information is stored DNA, which exists in the cell as two long molecular coiled chains (DNA double helix). The intracellular genetic information is replicated, transcribed, and translated by enzymes, which then leads to protein synthesis (Fig. 1.27).

1.5.2

Variation

Variation refers to the differences between offspring and its parent under certain conditions, including variations in morphology and structure, virulence, drug resistance, and so on. The variability of microorganisms is divided into genetic variation and non-genetic variation. The Fig. 1.24 Synthesis of Gram-positive bacterial cell wall During cell division, the volume of the cell increases, and a former is due to changes in bacterial gene strucnew cell wall is formed. New cell wall materials are added ture. The new characteristics can be stably transto the pre-existing cell wall to maintain structural integrity mitted to future generations, which is why this

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Fig. 1.25 Division of Gram-positive bacteria (SEM) Cell division in Streptococcus gordonii, showing a clear division

Fig. 1.26 The growth curve of Escherichia coli

type of variation is called genotype variation, as the change is mostly irreversible (Fig. 1.28). The latter is caused by the influence of certain environmental conditions that do not change the genetic structure of bacteria. The change is not transmitted to the offspring and is therefore called a phenotypic variation. Genetic variation is rarely influenced by external environmental factors. Therefore, genetic variation tends to occur in individual bacterial cells, while phenotypic variation tends to occur in a bacterial flora due to the

effect of environmental factors. These variations can revert with the removal of the stimulating environmental factors.

1.5.3

Genetic Material of Bacteria

Nucleic acids are the basis of organismal heredity. Two types of nucleic acid exist: DNA and RNA. DNA is the genetic material in prokaryotic and eukaryotic organisms, while the genetic

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Fig. 1.27 The processes of replication, transcription, translation of intracellular genetic information, and protein synthesis

Fig. 1.28 DNA mutation

material in viruses is DNA and RNA. The genetic material found in microorganisms includes chromosomes, plasmids, bacteriophages, and transposable elements. Bacteriophages are viruses that infect bacteria, fungi, actinomycetes, mycoplasma, and spirochetes. They inject their genetic material into the infected host cell and can induce bacterial cell lysis under certain conditions.

Bacteriophages, known as phage for short, can only reproduce in specific host strains and have high specificity for their host. The specificity is related to phage cell binding molecules and the structure and complementarity of the host bacterial strain’s surface receptor molecules. Since phages are small, they can only be observed under electron microscope. They can be divided into three basic morphologies:

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sequence. These changes are stable and heritable. Mutations can generally be classified as gene mutations and chromosomal aberrations. The spontaneity and randomness of microbial mutations can be tested by using fluctuation test or replica plating (Figs. 1.31 and 1.32).

1.5.5

Fig. 1.29 The structural model of a phage The phage head is icosahedral is approximately 80  100 nm in size. It is consisted of the capsid protein which surrounds the internal nucleic acid. The tail is a tubular structure composed of protein, including the tail whiskers, tail collar, tail tube, tail sheath, tail fibers, and tail baseplate

tadpole-shaped, spherical, and rod-shaped. Most phages are tadpole-shaped and are made up of a head and a tail (Fig. 1.29). The relationship between the phage life cycle and its host bacteria is shown in Fig. 1.30. Phages that infect bacteria can produce two outcomes. The first, observed in virulent phage, involves phage multiplication resulting in the production of many progeny phages, bacterial cell lysis, and cell death. This cycle is known as the lytic cycle. The second outcome is lysogeny, observed in temperate phages. It involves the integration of phage nucleic acid with the bacterial chromosome, resulting in the formation of a prophage. The phage genetic material is reproduced when the bacterial cell divides.

1.5.4

Mechanism of Microbial Variation

Genetic variation in microorganisms has its basis in mutations caused by changes in their genetic

Gene Transfer and Recombination

Gene transfer is a process by which exogenous genetic material from a donor cell is transferred to the receptor cell. However, simply the process of transferring genetic material is not enough, as the recipient cell must be able to accommodate exogenous genes. Integration between the transferred gene and the DNA of the recipient cell is a process known as recombination, whereby the recipient cell acquires certain characteristics of the donor strain. Gene transfer and recombination in bacteria can take place through processes such as transformation, conjugation, transduction, cell fusion, and lysogenic conversion.

1.5.5.1 Transformation Transformation takes place when donor bacteria DNA is cleaved and free DNA fragments are directly taken up by receptors. As a result, recipient cells acquire certain genetic traits from the donor cell. This phenomenon was confirmed in Streptococcus pneumoniae, Staphylococcus, and Haemophilus influenzae [8–10]. Griffith first showed that bacterial transformation takes place by infecting mice with Streptococcus pneumoniae in 1928 [2]. An outline of the experiment is shown in Fig. 1.33. 1.5.5.2 Conjugation Conjugation [3] is the method by which bacteria physically connect with one another through their pilus to transfer genetic material (mainly plasmid DNA). Plasmid transfer from the donor to the recipient cell results in that the recipient cell acquires some of the genetic traits of the donor cell. Plasmids that can be transferred through conjugation are called conjugative plasmids, which include the F, R, Col, and virulence

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Fig. 1.30 Lysogenic and lytic cycles of lysogenic phage

Fig. 1.31 Fluctuation test The fluctuation test shows that mutations pre-exist in a population of bacteria in the absence of selection. This was tested by Luria and Delbruck [4] using naturally existing phage-resistant mutations in bacterial populations. A given concentration (103/ml) of Escherichia coli sensitive to specific phages were inoculated in two equal volumes (10 ml) of broth medium. One inoculum was concentrated in a large test tube, and the other was evenly distributed into 50 small test tubes. After 24–36 h incubation under

the same conditions, the bacterial cultures from the large and small tubes were plated onto phage-containing plates, and the number of colonies was measured. The results typically showed that of 50 phage-coated plates inoculated with bacteria from the large test tube, the fluctuation in the colony number was small (3–7). On the other hand, when the 50 small tubes were plated onto 50 plates, the number of the colonies tended to fluctuate significantly, from zero colonies to several hundreds

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Fig. 1.32 Replica plating Replica plating (Lederberg et al. 1952) [5] involves plating antibiotic-susceptible strains on agar plates in the absence of antibiotics until scattered single colonies grow. Using a block covered with sterile velvet, gently press the velvet onto the surface of the agar plates so that bacterial colonies are imprinted onto the sticky velvet surface. Then, press to the velvet surface onto an agar plate containing antibiotics. After an appropriate culture time, bacteria susceptible to

the antibiotics are completely suppressed, but drugresistant colonies will be visible on the plate. We can find colonies corresponding to drug-resistant colonies on the original plates lacking antibiotics. The drug-resistant colonies can then be transplanted to culture broth containing the appropriate antibiotics to observe bacterial growth. Although the bacterial colonies on the original agar plate have never been in contact with antibiotics, they are nonetheless resistant to antibiotic drugs

plasmids. The F plasmid encodes the pilus and controls pilus formation and whether or not the pilus enables conjugation (Fig. 1.34). Plasmids that cannot be transferred between bacteria through a pilus are called non-conjugative plasmids. The non-conjugative plasmid ColE1 is relatively low in molecular mass and does not encode the necessary gene required for it to be transferred from one cell to another. However, if there is another conjugative plasmid in the host cell, ColE1 can tag along and be transferred from one cell to another. For example, the F plasmid which encodes the genes necessary for pilus formation can help ColE1 transfer from one cell to another (Fig. 1.35). Conjugation is widespread in Gram-negative bacteria and can be observed in almost members of the Enterobacteriaceae. Some Gram-positive bacteria have been reported to conjugate (e.g.,

Streptococcus, Bacillus subtilis), and the phenomenon has also been observed in Streptomyces.

1.5.5.3 Transduction Transduction uses a temperate phage as the vehicle by which DNA from a donor cell is transferred into a recipient cell [6, 7]. Transduction can transfer larger fragments of DNA than transformation. According to the genes involved in transduction, the process can be divided into generalized transduction and restricted transduction (Figs. 1.36 and 1.37). Generalized transduction can transfer plasmids. The transduction of plasmid R in Staphylococcus aureus is a very important clinical feature. In generalized transduction, the packaged DNA can be from any part of the donor strain chromosome. When phages exit the lysogenic

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Fig. 1.33 Streptococcus pneumoniae transformation in mice Streptococcus pneumoniae with a polysaccharide capsule belong to the virulent type III strain. These colonies are smooth (S) in appearance. Streptococcus pneumoniae without the polysaccharide capsule belong to the avirulent type II strain and appear as rough (R) type colonies. In the classic Griffith experiment, type II-R bacteria and type III-S bacteria were injected into mice. Mice that received the type II-R bacteria survived and those that received the type III-S bacteria died. The type III-S bacteria were isolated from the blood of the dead mice and were heated

until they were no longer active. These dead type III-S bacteria were injected into mice, and the mice survived. However, when dead type III-S bacteria and live type II-R bacteria were both injected into the same mice, the mice died, and type III-S bacteria were isolated from their blood. This experiment showed that the live type II R-type bacteria were able to obtain genetic material from dead type III-S bacteria that transformed them from an avirulent strain to a virulent one. It also suggests that the genetic material encoded the capsule virulence factor from type III-S bacteria

Fig. 1.34 The transfer and replication of the F plasmid during conjugation Bacteria possessing the F plasmid and F-pili are the male equivalent strain (F+), while bacteria lacking the F plasmid and F-pili are the female equivalent strain (F ). When the F+ and F strains are present in the same environment, the F-pilus from the F+ bacterial cell conjugates to the F surface receptor. The F-pilus gradually shortens so that the two cells are pulled close to one another and a channel

is formed. Plasmid DNA from the F+ bacteria break at the origin of transfer (oriT), and the 5 ‘end extends through the channel into the F bacteria. Single-stranded DNA in both bacterial cells replicate by rolling circle replication, and each cell forms a complete F plasmid. Therefore, the donor cell has not lost its F plasmid after the transfer and the recipient cell becomes F+ strain bacteria after receiving the F plasmid

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Fig. 1.35 Non-conjugative plasmid (ColE1) induced to transfer by F plasmid The ColE1 plasmid can be induced to transfer from a donor to a recipient cell. This process requires two genes encoded on the ColE1 plasmid: the specific site bom on ColE1 DNA (also known as the nic locus) and the nuclease

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encoded by the mobA gene. When both F and ColE1 plasmids are present in the same bacterial cell, the mobA gene is transcribed and its product creates a single strand break at the bom locus to form a gap. As a result, the ColE1 plasmid goes from a supercoiled plasmid to an open loop

Fig. 1.36 Generalized mode of transduction

phase, the prophage will excise itself from the bacterial chromosome and enter lytic phase. In the late stages of the lytic phase, phage DNA is replicated in large quantities, and errors can occur during phage assembly. Approximately one in 105–107 phages will contain bacterial DNA fragments that have been mistakenly packaged into the phage head; these results in the formation of a transducting phage. Transducting phages can infect another host bacterium and inject the DNA fragment it is carrying into the recipient cell,

thereby transferring DNA from one bacterial cell to another. Restricted transduction, or specific transduction, describes the process in which transduction is restricted to specific genes from the chromosome of the donor strain. If phage λ is transferred into Escherichia coli K12 while it is in the lysogenic phase, the phage DNA is integrated into a specific site in the Escherichia coli chromosome, between the galactose (gal) and biotin (bio) genes.

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Fig. 1.37 Restricted mode of transduction

References 1. Joseleau-Petit D, Liebart JC, Ayala JA, D’Ari R. Unstable Escherichia coli L forms revisited: growth requires peptidoglycan synthesis. J Bacteriol. 2007;189(18):6512–20. 2. Griffith F. The significance of pneumococcal types. J Hyg (Lond). 1928;27(2):113–59. 3. Lederberg J, Tatum EL. Gene recombination in E. coli. Nature. 1946;158(4016):558. 4. Luria SE, Delbruck M. Mutations of bacteria from virus sensitivity to virus resistance. Genetics. 1943;28(6):491–511. 5. Lederberg J, Lederberg EM. Replica plating and indirect selection of bacterial mutants. J Bacteriol. 1952;63 (3):399–406.

6. Morse ML, Lederberg EM, Lederberg J. Transduction in Escherichia coli K-12. Genetics. 1956;41 (1):142–56. 7. Morse ML, Lederberg EM, Lederberg J. Transductional heterogenotes in Escherichia coli. Genetics. 1956;41(5):758–79. 8. Straume D, Stamsås GA, Håvarstein LS. Natural transformation and genome evolution in Streptococcus pneumoniae. Infect Genet Evol. 2015;33:371–80. 9. Marraffini LA, Sontheimer EJ. CRISPR interference limits horizontal gene transfer in staphylococci by targeting DNA. Science. 2008;322(5909):1843–5. 10. Poje G, Redfield RJ. Transformation of Haemophilus influenzae. Methods Mol Med. 2003;71:57–70.

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Techniques for Oral Microbiology Xian Peng, Biao Ren, Yuqing Li, Xuedong Zhou, Jing Xie, Chenchen Zhou, Demao Zhang, Xin Zheng, and Xinxuan Zhou

Abstract

In recent years, with the rapid development of molecular biology, microecology, systems biology, and other disciplines, oral microbiology has been transformed from the traditional microbiology which is based on isolation and culture to a modern oral microbiology which is interdisciplinary and multi-scale. The continuous development of microscope technologies has made it possible for us to understand the cell structure, biofilm structure, and ecological interactions of oral microorganisms. With the application of 16S rDNA gene sequencing technology, metagenomics technology, and pyrosequencing technology being gradually improved, it has laid a foundation for elucidating the composition of oral microbiome and the correlation between oral microbes and human diseases. The main contents of this chapter include smear and X. Peng (*) · B. Ren · Y. Li · J. Xie · C. Zhou · D. Zhang · X. Zheng · X. Zhou State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China e-mail: [email protected] X. Zhou State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

stain techniques, isolation, incubation and identification techniques, microscopy techniques, oral microecology techniques, and oral microbiome techniques. Keywords

Smear and stain · Isolation and identification · Microscopy techniques · Oral microecology techniques · Oral microbiome techniques

2.1

Smear and Stain Techniques

Smear test and slide stain are basic techniques for microbial identification and are primarily used for morphological observation. The combination of smear and stain is widely used in oral microbiology research to differentiate between spirochetes, bacteria, fungi, and protozoa, as well as to identify specific cellular structures including spore, capsule, flagellum, and others. Currently, some of the commonly used techniques for cellular morphology examination are direct smear test and stained smear test, including Gram stain and Congo red staining. The procedure for smear test is shown in Fig. 2.1. Specific stains can be used to observe the specific microbial structures, including the bacterial spore, capsule, flagellum, fungal hypha, chlamydospore, etc.

Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China # Zhejiang University Press 2020 X. Zhou, Y. Li (eds.), Atlas of Oral Microbiology: From Healthy Microflora to Disease, https://doi.org/10.1007/978-981-15-7899-1_2

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Fig. 2.1 Procedure of smear test

2.1.1

Kopeloff’s Modification of Gram Stain

Gram stain is a commonly used method for the identification of bacteria and fungi. After Gram staining, bacterial species can be generally differentiated into Gram-positive and Gramnegative groups, and most clinical anaerobic microbes can be differentiated by their typical cellular morphology. In addition, mycoplasma with their ring shape can also be identified. Kopeloff’s modification of Gram stain is recommended by the Virginia Polytechnic Institute (VPI) for better visualization and differentiation of microbes.

2.1.1.1 Mechanism Many theories have been proposed to explain the mechanism of Gram staining, including isoelectric point theory, chemical theory, and permeation theory. Among them, permeation theory has gained wide acceptance. It is believed that crystal violet placed on the microbe smear slides during staining interacts with aqueous iodine to form insoluble precipitates within the cell. During decolorization, alcohol and acetone can dissolve the lipid in the outer membrane and leach the precipitated dye-iodine complex out of the microbial cell. Gram-positive microbes have a thick cell wall with highly cross-linked peptidoglycans and

relatively fewer lipids. Thus, Gram-positive microbes cannot be easily decolorized and become deeply stained in purple. In contrast, Gram-negative microbes have a thin membrane with high lipid content and little peptidoglycan. The precipitated dye-iodine complex can therefore be easily dissolved and leave the cell. As a result, the cell is stained red from the counterstain.

2.1.1.2

Preparation of Gram Stain Reagents (Commercially Available) 1. Crystal violet Liquid A (add 10 g crystal violet to 100 0 ml distilled water and mix well) Liquid B (add 50 g NaHCO3 to 1000 ml distilled water and mix well) 2. Gram’s iodine Dissolve 4 g NaOH into 25 ml distilled water and add 20 g iodine and 1 g KI. After everything is completely dissolved, add 975 ml distilled water, mix well, and store the liquid in a brown bottle. 3. Decolorizers Add 300 ml acetone to 700 ml 95% alcohol and mix well. 4. Fuchsin counterstain

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Add 5–10 g basic fuchsin to 100 ml 95% alcohol to form a supersaturated liquid. Mix 10 ml of the supersaturated liquid with 90 ml 5% carbolic acid solution followed by the addition of 900 ml water. Mix well and filter thorough filter paper. (Safranin solution can also be used in place of fuchsin solution for counterstain: add 20 g safranin to 100 ml 95% alcohol and adjust the volume to 1000 ml with distilled water.)

2.1.1.3 Staining Procedure Gram stain procedure is shown in Fig. 2.2. 1. Add crystal violet Liquid A onto the fixed smear slide. After 5 s, add Liquid B and gently shake the slide to mix Liquids A and B for 30 s. Gently rinse off the crystal violet with tap water (Fig. 2.2a). 2. Flood the smear slide with Gram’s iodine for 30 s. Gently rinse off the iodine with tap water (Fig. 2.2b). 3. Add decolorizer to the smear while holding the slide at an angle to allow the decolorizer to drain. Gently shake the slide for 5–10 s until crystal violet leaches out. Gently rinse off the decolorizer with tap water (Fig. 2.2c). 4. Flood the smear with fuchsin or safranin counterstain for 5–10 s. Gently rinse off any excess fuchsin or safranin with tap water. Drain and air-dry the slide before observation (Fig. 2.2d).

Fig. 2.2 Gram stain procedure

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2.1.1.4 Reporting Results Examine the slide under a light microscope using oil immersion. Gram-positive bacteria appear purple (Figs. 2.3 and 2.4), while Gram-negative bacteria appear pinkish red (Figs. 2.5 and 2.6). 2.1.1.5 Precautions 1. Smear only a monolayer of cells on the slide. 2. Avoid over-heating of the smear slide. 3. Slightly prolong the time of decolorization if the smear layer is thick. 4. Old cultures of Gram-positive bacteria may appear as Gram-negative bacteria.

2.1.2

Staining Bacterial Spores

Gram stain and fuchsin-methylene blue are commonly used to observe bacterial spores.

2.1.2.1 Gram Stain The Gram stain procedure is the same as described above. The result of Gram stain is shown in Fig. 2.7. 2.1.2.2 Fuchsin-Methylene Blue Stain Reagent preparation: 1. Fuchsin solution: Add 5–10 g basic fuchsin to 100 ml 95% alcohol to form a supersaturated liquid. Mix 10 ml of the supersaturated liquid with 90 ml 5% carbolic acid solution followed by 900 ml water. 2. Methylene blue solution: Liquid A: add 0.3 g methylene blue to 30 ml 95% alcohol Liquid B: add 0.01 g KOH to 100 ml distilled water Mix Liquids A and B 3. Decolorizer: 95% alcohol Stain procedure: Flood the smear slide with fuchsin solution. Heat the slide on the alcohol burner to steam for 3–5 min (avoid drying out the smear slide and add fuchsin solution when necessary). Cool off the slide and rinse off excess fuchsin under tap water. Flood the slide with decolorizer for 1 min and rinse. Stain the slide

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Fig. 2.3 Gram-positive bacteria (Actinomyces naeslundii)

Fig. 2.4 Gram-positive yeast (Candida albicans)

with methylene blue solution for 0.5 min and rinse. Observe the slide under microscope using oil immersion. Results of spore staining are illustrated in Fig. 2.8.

2.1.3

Staining the Bacterial Capsule

The bacterial capsule can be stained using a variety of methods, including the Murs stain, ink

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Fig. 2.5 Gram-negative bacteria (Eikenella corrodens)

Fig. 2.6 Gram-negative bacteria (mycoplasmal pneumonia)

methylene blue stain, Hiss copper sulfate stain, Ott safranin stain, etc. Encapsulated bacteria that are found in the oral cavity include Streptococcus pneumoniae, Porphyromonas gingivalis, and others.

2.1.3.1 Murs Stain Preparation of staining solutions: 1. Staining solution (carbol fuchsin solution): add 1 g basic fuchsin to 10 ml 95% ethanol

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Fig. 2.7 Bacillus subtilis (Gram stain) Under light microscope with oil immersion, bacterial sporex appear colorless with strong refraction. They are located toward the center (central spore) or ends of the cell (terminal spore) with round or ovoid shape

Fig. 2.8 Spore stain with Clostridium tetani (fuchsinmethylene blue stain) The cell appears blue and the spore appears red. A round spore is located at the end of the cell, giving the cell a drumstick appearance

solution, and then add 40 ml 5% aqueous carbolic acid solution and mix. 2. Mordant staining solution: mix 100 ml 2% tannic acid, 250 ml 10% potassalumite

(K2SO4
12H2O), and 100 ml 7% saturated mercuric chloride. 3. Counterstaining solution (alkaline methylene blue solution): add 0.3 g methylene blue to

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Fig. 2.9 Capsule of S. pneumoniae (Murs staining method) Bacterial cells are stained red, and the capsule around the cell shows up as blue transparent circles

30 ml 95% ethanol solution, and then add 100 ml 0.01% potassium hydroxide solution. 4. Destaining solution: 95% ethanol. Staining protocol: Prepare a smear sample using physiological saline and fixed. Heated carbol fuchsin solution is used to stain the smear for 1 min. Flush with water after destaining using 95% ethanol. Stain with mordant staining solution for 0.5 min, and destain with 95% ethanol for 0.5 min. Stain with alkaline methylene blue solution for 0.5 min, and then flush the stain solution with water. Examine microscopically after air-drying. Typical results are shown in Fig. 2.9.

2.1.3.2 Wright’s Stain Bacterial capsules can also be stained with the Wright’s staining method. Furthermore, the Wright’s staining method can be used to stain Rickettsia and spirochetes, which appear purple after staining. Preparation of staining solutions: 1. Wright stain solution: Weigh out 0.1 g dry Wright’s stain (same as eosin methylene blue stain). Grind eosin Y using mortar and pestle until it becomes a fine powder. Add 10 ml

methanol to the mortar to dissolve the eosin Y until fully dissolved. Add 20 ml methanol, mix, and allow to stand for a moment. Decant the liquid into a clean storage bottle, add methanol to the mortar, mix, and repeat several times until all of the stain is dissolved. Sixty milliliters of methanol should be used for 0.1 g stain. Shake and seal the storage bottle, and store it in the dark at room temperature. 2. Phosphate buffer (pH 6.4–6.8): 30 ml 1% KH2PO4, 73.5 ml M/15 KH2PO4, 20 ml 1% Na2HPO4, 26.5 ml M/15 Na2PO4, complete with H2O to1000 ml). Staining protocol: Prepare a smear using the conventional method and allow to air dry. Place 4–6 drops of Wright’s stain onto the smear and allow to staining for 1 min. Add an equal amount of phosphate buffer (pH 6.4–6.8). Shake gently to allow the phosphate buffer and the staining solution to mix evenly. Let the stain stand for 6–8 min and flush with water. The smear should appear pink (if the smear appears bluish violet, it can be destained using by 20% hydrochloric acid methanol solution). A typical result is shown in Fig. 2.10.

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Fig. 2.10 Capsules of Histoplasma capsulatum (Wright’s stain) Bacterial cytoplasm is stained light blue, and the capsules surrounding the cells are colorless transparent circles

2.1.4

Flagellar Stain

Flagellar stain is used to observe whether bacteria have flagella, as well as the number and location of flagella. It is a helpful tool for bacterial identification. Flagellar staining is of great importance in identifying motile oral bacteria. For example, Selenomonas can have up to 16 flagella, while straight Campylobacter only has one polar flagellum.

Preparation of Staining Solutions Liquid A: 20 ml 2% tannic acid solution dissolved in heated water bath, 20 ml 20% potassium solution (heat to dissolve), 50 ml 5% aqueous solution of carbolic acid; mix all three solutions. Liquid B: alkaline fuchsin ethanol saturated liquid (same as in Gram stain solution). Mix 30 ml Liquid A with 10 ml Liquid B and filter. Keep the filtrate for 6–10 h at room temperature for optimum results.

farthest away from the original streak and resuspend in a small tube in sterile distilled water. Keep the small tube at room temperature for 20–30 min, or keep it in a 37  C incubator for 4–5 min to allow the culture to become fully resuspended. Remove a loop of the bacterial suspension with a disposable sterile loop, and place it gently on a clean slide. Tilt the slide gently or push the bacterial suspension with the loop. Then allow the smear to dry naturally at room temperature or in the 37  C incubator.

2.1.4.1

2.1.4.2 Smear Preparation Flagellar staining requires correct preparation of the smear. Pick a small, young agar culture

2.1.4.3 Staining Protocol Drop the Liquid A and Liquid B mixture onto a dry smear and allow to stain for 1–2 min. Flush the slide with water. When the smear is dry, observe the result under an oil immersion lens (Fig. 2.11).

2.1.5

Staining Procedure for Special Fungal Structures

Gram stain and lactophenol cotton blue stain are used to visualize certain fungal structures that are significant for identification. These include the germinal tube, hyphae, and spores and are

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Fig. 2.11 Periplasmic flagella (flagella staining) The bacterial cell is stained red, and the light red flagella can be seen around the bacterial cell

especially important in clinical analyses of oral mucosal diseases. For example, in the identification of thrush, angular stomatitis closely related to Candida albicans. The checking specimen is originated from albuginea or focal secretion of oral mucosa. Gram staining and acid phenol cotton blue stains are often used to verify the structures in fungi such as germ tubes, hyphae, and spores, especially for oral mucosal diseases such as thrush and angular cheilitis-related Candida albicans. Most specimens are taken from the oral mucosa lesions albuginea and secretions.

2.1.5.1 Crystal Violet Stain Crystal violet staining solution is prepared in the same way as Liquid A used in Gram stain. Take a small quantity of culture and mix with physiological saline to prepare a smear. Stain the smear with crystal violet solution. Observe under oil immersion lens (Figs. 2.12 and 2.13). 2.1.5.2 Lactophenol Cotton Blue Stain Preparation of staining solution: Dissolve 20 g carbolic acid (solid), 20 ml lactic acid, and 40 ml glycerol into 20 ml distilled water (heat as gently as possible). Add 0.05 g cotton blue, shake until well-mixed, and filter before storing.

Staining protocol: Place a drop of lactatophenol cotton blue staining solution onto a clean slide and mix the fungal culture or clinical sample with the staining solution. Place a coverslip on top and heat gently. Press the coverslip gently to remove any bubbles. Observe the slide under an oil immersion lens (Figs. 2.14 and 2.15).

2.1.6

Giemsa Stain for Mycoplasma

Giemsa stain is used to observe Mycoplasma (Fig. 2.16).

2.1.7

Negative Congo Red Staining of Plaque Bacteria

Due to the simplicity of this method in producing high-quality stained smears, the Congo red staining method has been widely used in the examination of periodontitis and other oral clinical specimens. 1. Sample: Saliva, plaque, other oral specimens. 2. Staining solution: 2% Congo red solution.

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Fig. 2.12 Germinal tube and blastospore of Candida albicans (crystal violet stain) The germinal tube and blastospore appear purple when Candida albicans is incubated in 0.5–1 ml human serum or in sheep serum for 2–4 h at 37  C and stained by the crystal violet staining method

Fig. 2.13 Pseudohypha chlamydospore of Candida albicans (crystal violet stain) Pseudohypha chlamydospores of Candida albicans appear purple when stained with crystal violet. The chlamydospores are big, spherical, and thick-walled, and they are located at the tip or side wall of unevenly stained pseudohypha

3. Staining procedure: Place a drop of the 2% Congo red solution on a clean slide. Mix the specimen and the staining solution and spread the mixture thinly with a slide. After the smear dries naturally, smoke it over a bottle of

concentrated hydrochloric acid until the red smear turns blue. 4. Results: The blue smear is examined under light microscope under oil immersion lens (Fig. 2.17).

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Fig. 2.14 Germinal tube and blastospore of Candida albicans (lactatophenol cotton blue stain) Germinal tube and blastospores are stained bright blue after Candida albicans are incubated in 0.5–1 ml human serum or in sheep serum for 2–4 h at 37  C and stained with lactatophenol cotton blue stain

Fig. 2.15 Pseudohypha chlamydospore of Candida albicans (lactophenol cotton blue stain) Pseudohypha chlamydospores of Candida albicans are bright blue; the big, spherical, thick-walled chlamydospore is at the tip or side wall of the pseudohypha

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Fig. 2.16 Mycoplasma (Giemsa stain) Typical ring-shaped mycoplasma cells stain purple

Fig. 2.17 Negative Congo red stain of plaque bacteria The background is blue while the bacteria remain unstained (bright white with different shapes), creating what is called a negative stain. (a) coccoid cells; (b) short bacilli; (c) fusiform bacilli; (d) long bacilli; (e) filamentous bacilli; (f) curved rods; (g) spirochetes

Listgarten classification is generally used in negative Congo red staining and dark-field microscopy to classify the observed plaque bacteria. The method involves selecting an evenly spread field, counting 200 bacterial cells, and reporting the percentage of different species according to their morphology.

1. Coccoid cells: Cell diameter from 0.5 to 1.0 μm, including several kinds of coccobacilli. 2. Straight rods: Cells measure approximately 0.5–1.5 μm in width, 1.0–1.9 μm in length. Some types of mycobacteria are included.

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3. Filaments: Cells measure 0.5–1.5 μm in width and the ratio of length to width is greater than 6. Most bacteria are shaped as irregular long filamentous cells. 4. Fusiform cells: Cells measure approximately 0.3–1.0 μm in diameter, 10 μm in length, and show tapered ends. 5. Curved rods: Cells are similar in dimension to straight rods, but are curved or crescentshaped. 6. Spirochetes: Cells have a spiral shape and measure 0.2–0.5 μm in width and 10–20 μm in length.

2.1.8

Protozoan Smears

Entamoeba gingivalis and Trichomonas tenax are the main protozoans found in the oral cavity. Wet smears can be prepared using fresh specimens to observe the morphology and mobility of the protozoans. Giemsa stain can be used on a fixed specimen to observe the cellular structure.

2.1.8.1 Wet Smear for Fresh Samples Mix gingival margin plaque samples or subgingival plaque with saline to prepare fresh wet smears. Examine the shape and mobility of the protozoans under the microscope immediately. Trophozoites of Trichomonas tenax are lively pear-shaped parasites which move faster than Trichomonas vaginalis. Examination of fresh smears should preferably be performed at room temperatures above 20  C and should be completed within 30 min after smear preparation in order to avoid any influence from the lower temperature of the room or the time spent on the slide on the motility of the protozoan. 2.1.8.2 Giemsa Staining Place a drop of saline solution onto a clean slide and mix it with the clinical sample (e.g., subgingival plaque). Stain the air-dried specimen with Giemsa stain solution, and examine it with a light microscope under oil immersion.

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2.2

Isolation, Incubation, and Identification Techniques

The isolation and identification of oral microorganism can be difficult. Because oral microorganism are great in number and composed of diverse species, new genera and species are constantly being discovered, while the classification of some previously discovered species change with time. Therefore, it is currently impossible to isolate and identify all the microbes in an oral specimen. Phenotypic identification of organisms is the most basic and important part of microbiology. Classical methods for bacteria identification require observation of the phenotypic characteristics of a pure bacterial culture, including characteristic colonies (size, color, shape, etc.), cell characteristics (size, shape, arrangement, and stain), special structures (with or without spores, capsule, pili, and flagellum), culture characteristics (sensitivity to oxygen, optimum growth temperature, pH, requirements for nutrients and growth factors, etc.), metabolites, etc. Bergey’s Manual of Systematic Bacteriology is an authoritative reference book for bacterial isolation.

2.2.1

Collection and Transportation of Samples

2.2.1.1 Sample Collection 1. Collection of saliva samples: Saliva samples include stimulated salivary and unstimulated salivary samples. Stimulated salivary samples are collected when subjects are chewing paraffin or a rubber block, while unstimulated salivary samples are secreted naturally by the subjects. More saliva is collected by stimulation, but at the same time, it can influence the mucosa, oral plaque, and the oral microflora. Saliva samples for microbiological examination should preferably be fresh unstimulated salivary samples. The best time for collection is early in the morning upon waking and before the teeth are brushed, or alternatively,

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samples can be collected between meals (around 10:00 or 16:00). Subjects should gently rinse their mouths with warm water to remove any food residue. Anywhere from 0.5 to 1 ml of naturally secreted saliva is collected into a sample tube, or saliva samples can be directly taken from the oral cavity using sterile pipette tips. 2. Collection of plaque samples: As plaque microbes are complex in their composition, plaque samples should be collected in different ways depending on specific clinical requirements and purposes. If necessary, plaque indicators should be used to show dental plaque. Plaque samples can be found on adjacent surfaces and fissures in occlusal surfaces. Before sample collection, subjects should gargle with warm water to remove any food residue. Then, sterile gauze or a yarn ball should be used to absorb saliva, while plaque samples are collected. A sterile probe is commonly used in plaque collection from occlusal surface fissures. Plaque found on adjacent surfaces can be collected with sterile probe, dental floss, or fine orthodontic wire. Sterile curettes can be used to collect root surface plaque samples. Plaque samples on the gum or in the gingival margin can be collected using a spoon scaler. Subgingival plaque is divided into attached plaque and unattached plaque. Collection using the MooreOO bacteria taker or using a sterile paper point is currently the most widely used and the easiest way to collect subgingival plaque. Plaque samples from infected root canals are usually collected with sterile paper point. 3. Collection of other samples of infected tissue: In samples of infected tissue such as lip carbuncle, aerobic bacteria and facultative anaerobes such as Staphylococcus aureus are the main pathogens and are generally collected with sterile cotton swabs. Samples of purulent fluid from a periodontal abscess can be collected with sterile syringes. Tissues in the alveolar socket are generally collected as samples of dry socket developed after tooth extraction.

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4. Collection of oral mucosal diseases samples: White membranous materials are usually collected with curettes or cotton swabs. To collect the quantitative samples, filter papers of a specific size are used.

2.2.1.2 Sample Transportation Fungal species, aerobic bacteria, and general facultative anaerobic bacteria can be collected by cotton swab sampling and transported in sterile tubes. For most anaerobic bacteria or microaerophilic bacteria, samples must be sent to the laboratory as soon as possible and maintained in an anaerobic environment. With the exception of pus or saliva samples that can be inserted directly into the sterile rubber stopper of the syringe needle used to collect the sample for transportation, most anaerobic samples must be placed in pre-reduced culture media and for transportation to the laboratory immediately after acquisition. This is to minimize the death of oxygen-sensitive bacteria during transportation. Chairside inoculation and anaerobic transportation can help improve the detection rate of obligate anaerobic bacteria. Transportation in pre-reduced medium requires that samples be placed immediately in a small covered tube with containing pre-reduced liquid transportation medium. In order to reduce the infiltration of oxygen during transportation, sterile liquid paraffin can be placed on the pre-reduced medium to isolate it from air. For clinical specimens that cannot be examined in a timely manner or that must be transported over a long distance, anaerobic bags (commercially available) or pre-reduced liquid medium in a spiral tube sealed with liquid paraffin can be used for transportation.

2.2.2

Suspension and Dilution of Samples

Clinical oral infections are mixed infections involving many different species of bacteria within a concentrated plaque mass. In order to obtain pure cultures of individual bacteria, oral

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Fig. 2.19 Sample dispersion (ultrasonic generator)

difficult to avoid oxygen infiltration during ultrasonic dispersion, which can lead to the death of anaerobic bacteria in the sample. Fig. 2.18 Sample dispersion (vortex generator)

clinical specimens usually require processing and dilution after inoculation.

2.2.2.1 Sample Suspension in Solution Generally, two methods are used for sample suspension: spiral vortex oscillation (Fig. 2.18) and ultrasonic dispersion (Fig. 2.19). Spiral vortex oscillation is widely used because of its ease of operation and low cost. All that is required is for the sample collection tube to be placed on the vortex generator and allowed to oscillate for 10–20 s. Adding 5–6 small sterile glass beads (110–150 μm) into the small tube can help improve sample dispersion. Ultrasonic dispersion yields superior sample suspension, but the disadvantage is that a sonicator, which is an expensive piece of equipment, is required and the microbial detection rate can drop because spirochetes, Porphyromonas, Prevotella, and other Gramnegative bacteria are easily lysed. It is also

2.2.2.2 Sample Dilution As oral clinical samples are mixed bacterial samples containing a great number and variety of bacteria, proper dilution with the correct diluent must be performed prior to inoculation to obtain single colonies after sample suspension. The solution in which the sample was transported can be used as a diluent; otherwise, phosphate buffer (pH 7.2) can also be used. A tenfold dilution series is generally performed. Under aseptic conditions, 0.1 ml of the specimen sample is added to 0.9 ml diluents. After thorough mixing, 0.1 ml of the mixture (101) is added to a tube containing 0.9 ml diluent and mixed again. Using this method, tenfold dilutions are made in series (Fig. 2.20). Due to differences in sample bacteria content, different samples require different dilutions. For example, saliva samples should be diluted to 104–106, gingival groove plaque should be diluted to 101–102, while mixed plaque should be to 103–105.

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Fig. 2.20 Sample dilution (tenfold serial dilution)

Fig. 2.21 Spread method protocol

2.2.3

Inoculation and Incubation of Samples

bacilli. To culture most aerobic and facultative anaerobic bacteria, the blood agar (BA) is used.

In addition to selecting the appropriate medium for inoculation, other considerations include degree of dilution before inoculation, method of inoculation, incubation environment, time needed for colony establishment, purpose of incubation, and microbial species.

2.2.3.2 Sample Inoculation Spread method, drop method, and spiral inoculation method are used for oral clinical bacteriology samples. Appropriate dilutions of the specimen solution are quantitatively inoculated onto the agar plate.

2.2.3.1 Choice of Medium The medium commonly used for oral bacteria include brain-heart infusion (BHI) medium, trypticase soy medium (TSA), and TPY medium. These can be used to cultivate most bacteria from an oral sample. Five percent defibrinated blood (or 5% serum), chlorinated hemoglobin, and vitamin K1 must be supplied to the culture medium for some obligate anaerobic Gram-negative

1. Spread method: Ten microliters from appropriately diluted samples are taken with a micropipettor and placed on the surface of the agar medium. The sample is then evenly coated on the agar surface using a sterile glass spreader (triangle rod and L-shaped rod). At the appropriate dilution, each bacterial cell from the specimen should form a single colony after incubation (Figs. 2.21 and 2.22).

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Fig. 2.22 Colonies on a plate using spread method (periodontal bag samples)

Fig. 2.23 Drop method protocol

2. Drop method: Twenty-five or 50 μl samples at the appropriate dilution are dropped onto the surface of the agar using a micropipettor. Then, the plates are placed directly into the dry incubator (Figs. 2.23 and 2.24). 3. Spiral plater method: The spiral plater the most advanced method for inoculating bacteria for the purpose of counting colony forming units. The sample liquid is automatically diluted and inoculated using a needle tip on the surface of the agar plate by the instrument, and the bacteria colonies grow uniformly along the spiral trajectory after incubation (Fig. 2.25). As a result, sample counting and bacterial colony observation is more accurate and reproducible. Details are included in Sect. 2.4 of this chapter.

2.2.3.3 Sample Incubation Media containing clinical samples are incubated according to the requirements of each specific sample, including oxygen requirement, temperature, and time. Oral clinical specimens such as infected root canal, pericoronitis infection, samples taken after tooth extraction, and subgingival periodontitis plaque samples, which are mostly mixed bacterial samples, may involve different oxygen requirements, as there are a variety of microorganisms each with their respective requirements. Some bacteria require incubation under the anaerobic conditions, while others require incubation under aerobic conditions. The variability in culture conditions requires the researcher to become familiar with and to master

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Fig. 2.24 Colonies on a plate using drop method (saliva samples)

Fig. 2.25 Colonies on a plate using spiral plater (plaque samples)

the growth characteristics of bacteria in the mouth to avoid mistakes in the incubation. Usually, anaerobic cultures grow best in atmospheric conditions with 80% N2, 10% CO2, and 10%

H2, with a temperature of 36  C–37  C and approximately 48–72 h culture time. Some oral bacteria, such as forsyth steiner bacterium, Treponema, and others, require 1 week of anaerobic

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Fig. 2.26 Anaerobic glove box

culture. Commonly used anaerobic incubation devices include the anaerobic glove box, anaerobic incubator, anaerobic bag, etc. (Fig. 2.26). The initial steps of bacterial identification involve observing colonies morphologies following incubation and performing Gram stain. After this, a second round of purification is carried out to further complete phenotypic and genotypic identification using routine protocols for microbial separation and identification. For certain microorganisms, special separation, cultivation, and other identification techniques may be adopted.

2.2.4

Growth Characteristics and Identification

2.2.4.1 Growth Characteristics Bacterial colony morphology can be described in terms of its size, color, shape, growth pattern, and other characteristics. Hemolysis is one of the basic tests used for bacterial identification. Some bacteria produce hemolysis (Figs. 2.27, 2.28, 2.29, 2.30 and 2.31), some produce gas (Fig. 2.32), and some exhibit specific growth patterns, such as the migration of Proteus (Fig. 2.33). 2.2.4.2 Biochemical Tests Biochemical tests are among the most important methods for microbial identification. Routine

biochemical tests include tests for carbohydrate fermentation (Fig. 2.34), methyl red (Fig. 2.35), citric acid utilization (Fig. 2.36), and hydrogen sulfide production (Fig. 2.37). Microbial biochemistry tests shorten the time required to identify microbes, reduce costs, and ensure or enhance the accuracy of identification of an unknown sample. It is the fastest developing trend in microbial identification. In recent years, the rapid commercial test kits for anaerobic bacteria have become available in China and abroad. The most representative biochemical test kits are the Minitek identification system using paper substrates, API-20A system using dry powder substrates, PIZYMAN-IDENT rapid enzyme activity assay system using primary materials, RaPID-ANA systems, and fully automated microbial identification systems. The aforementioned microbial biochemistry reaction plate includes 30 biochemical matrices and their related biochemical test indicators, phosphate buffered saline (PBS), bacterial turbidity standard tube, and a few of identification series (Table 2.1). Due to the different types of experiments performed, the readouts for results are different. For example, esculin hydrolysis can be directly observed: black is positive, while colorless is negative. For sugar and alcohol fermentation acid test, BM (bromothymol blue-methyl red, BTB-MR) must be added to the result as a pH indicator. Red or yellow indicate a positive

44 Fig. 2.27 β-Hemolytic reaction

Fig. 2.28 α-Hemolytic reaction

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Fig. 2.29 White pigment (Staphylococcus epidermidis)

Fig. 2.30 Green pigment (Pseudomonas aeruginosa)

reaction, while green or blue indicate a negative reaction (Figs. 2.38 and 2.39).

2.2.5

Instruments for Microbiological Identification

2.2.5.1 Spiral Plater A spiral plater (Fig. 2.40) is used to inoculate plates to determine viable bacteria count. The

working principle behind a spiral plater is the use of a tip to dispense the liquid inoculum onto a Petri dish in a spiral pattern. The spiral plater deposits a known volume of sample onto a rotating agar plate so that the sample forms a spiral pattern with highest concentration at the center and lowest concentration at the outside of the spiral. Colony counting can be performed manually counting or by using automatic colony counters.

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Fig. 2.31 Black pigment (Porphyromonas gingivalis)

With a high degree of automation, simplicity of operation, and high reproducibility, spiral platers can significantly improve the efficiency and accuracy of bacteria counting while saving manpower, time, and culture space (Fig. 2.41).

2.2.5.2 Microbiology Analyzer The automated microbiology analyzer provides results by capturing images with a high-definition digital camera and analyzing them using its builtin software. Microbiology analyzers can be used for automatic colony counting, inhibition zone measurement to test antibiotic sensitivity test, and measurement of the hemolytic zone (Fig. 2.42). Microbiology analyzers are simple to operate, fast, and accurate and provide highly reproducible results. The instruments have a special lighting system suitable for various types of agar and plates. Other features include powerful analysis software and vivid full-color images which can be saved digitally or printed out, with results labeled beside the images and automatically recorded.

2.2.5.3 Microbial Identification System Microbial cells produce different enzymes during their metabolism of different carbon sources. The MicroStation automated microbial identification system is based on the differences in color and in turbidity that occur when these enzymes react with four azole substances (e.g., TTC, TV, etc.). With the use of a unique technology that detects the characteristic fingerprint of each microorganism and based on a large number of experiments and mathematical models, the corresponding database between the fingerprints and microbial species has been established. Identification results can be derived through comparisons between the unidentified microbial species and the reference database by software (Fig. 2.43). The MicroStation automated microbial identification system is used for microbial identification in clinical settings, food, dairy, pharmaceutical, and cosmetics industries and environmental microbial identification (in rivers, oceans, plants, animals, and insects). It can also be used to analyze microbial communities and aid in ecological research in the analysis of carbon utilization and

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lenses. Two beams are separated by intermediate objectives at an angle of about 12–15 which is called the stereo angle and then are imaged by their corresponding optical lens. The beams are not parallel but at an angle, providing a threedimensional image to both eyes by two separate optical paths. The magnification changes when the distance between intermediate objectives is changed. The digital imaging system connected to the computer to analyze and process images is composed of the stereomicroscope, a variety of digital ports, digital cameras, electronic optical lenses, and the image analysis software. Before using the stereomicroscope, the instrument should be adjusted for focus, diopter, and interpupil distance for each user in order to acquire the best image. The stereomicroscope is used to observe microbial colony morphology (Figs. 2.45, 2.46, 2.47, 2.48 and 2.49) and microbial distribution (Figs. 2.50, 2.51, 2.52 and 2.53).

2.3.2

Fig. 2.32 Production of gas (E. coli)

metabolism. There are four types of microplates specifically designed for the analysis of microbial communities and ecosystems research.

2.3 2.3.1

Microscopy Techniques Stereomicroscopy

The stereomicroscope (Fig. 2.44) is an optical microscope that produces a three-dimensional visualization of the sample being examined. The instrument is also known as a stereoscopic microscope or dissecting microscope. The optical structure of the stereomicroscope includes one shared primary objective and two sets of intermediate objective lenses or zoom

Scanning Electron Microscopy

The scanning electron microscope (SEM, Fig. 2.54) is mainly used to observe the topography of the cells in the samples over a large range of magnification. Sample preparation for SEM is simple. It is adaptable to various samples and does not require producing ultrathin slices. SEM is already a routine method in medical research and is especially crucial for studies on the morphologies and interactions of oral bacteria. SEM can be used to analyze and interpret observations on a micron or nanometer scale. The resolution of a field emission scanning electron microscope can reach as little as 1 nm. Another important feature of the scanning electron microscope is that it can be used to observe and analyze samples three-dimensionally due to its deep depth of field. The greater the depth of field, the more sample information is provided. In microbial identification, SEM is utilized to observe and detect surface morphology and structural characteristics of microbial cells.

48 Fig. 2.33 Migrating growth (Proteus)

Fig. 2.34 Carbohydrate fermentation test

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Fig. 2.35 Methyl red test

Fig. 2.36 Citric acid utilization test

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Fig. 2.37 Hydrogen sulfide production test

Table 2.1 Microbial biochemistry test identification series Main classification series Gram-positive anaerobic cocci Gram-negative anaerobic cocci Gram-positive anaerobic non-spore bacillus Gram-negative anaerobic non-spore bacillus

Sub-classification series I. Staphylococci and micrococci II. Streptococcus

I. Does not produce black pigment II. Produces black pigment

Gram-negative anaerobic Clostridium or Enterobacter Gram-negative facultative anaerobic bacillus Gram-negative Campylobacter

2.3.2.1 Mechanism The scanning electron microscope is used to scan sample areas or micro-volumes with a fine focused beam of electrons, producing various signals including secondary electrons, backscattered electrons, Auger electrons, characteristic X-rays, and photons carrying different levels of energy. When the electron beam scans the

sample surface, the signals will change according to the surface topography. The limited emission of secondary electrons within the volume close to the electron focusing area results in high image resolution. The three-dimensional appearance of images come from the deep depth of field and shadow effect of secondary electron contrast.

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Fig. 2.38 Gram-positive anaerobic cocci series II (Streptococcus series) (Streptococcus mutans)

Fig. 2.39 Gram-negative anaerobic non-spore bacillus series II (black pigment produced) (Porphyromonas gingivalis)

2.3.2.2 Operating Procedure 1. Bacterial sample preparation: Culture bacteria at 37  C for 48 h; identify cells as pure culture by morphological and biochemical tests; make bacterial suspension with 0.2 mol/L phosphate buffer (pH 7.2).

2. Sample washing and fixation: Wash sample twice with phosphate buffer or saline, fix sample for 2 h (at 4  C) with 2.5% or 3% glutaraldehyde, and then wash twice with phosphate buffer or saline.

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Fig. 2.40 WASP spiral plater

Fig. 2.41 The colonies on a plate prepared by spiral plater After inoculation on a 9 cm plate, the sample will be 1000-fold diluted on the outside track and the colonies will be evenly distributed

3. Gradual dehydration: Dehydrate samples with ethanol using a concentration gradient: 30% ethanol for 20 min, 50% ethanol for 20 min, 70% ethanol for 20 min, 80% ethanol for

20 min, 90% ethanol for 20 min, and finally 100% ethanol for 20 min twice. 4. Liquid exchange: Place dehydrated samples into 100% amyl acetate solution for 20 min for exchange.

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Fig. 2.42 Synbiosis automated microbiology analyzer

Fig. 2.43 MicroStation automated microbial identification system

5. Critical point drying: Place samples into a CO2 critical point dryer for CO2 critical point drying. 6. Metal coating: Coat the dried samples with ion sputter coater.

characteristics of oral microbial cells, as shown in the following examples (Figs. 2.55, 2.56, 2.57, 2.58, 2.59, 2.60, 2.61, 2.62 and 2.63).

2.3.3 2.3.2.3 Sample Observation Observe processed samples using the scanning electron microscope. The authors used the Inspect F field emission scanning electron microscope to observe surface morphologies and structural

Transmission Electron Microscopy

Transmission electron microscope (TEM, Fig. 2.64) is mainly used to observe the cell’s internal structures using ultrathin sections.

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Fig. 2.44 Stereomicroscope

Fig. 2.45 Isolated bacteria from dental plaque sample A: Actinomyces israelii colonies; B: Fusobacterium nucleatum colonies (BHI blood agar, anaerobic culture for 48 h, stereomicroscopy)

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Fig. 2.46 Characteristic colonies of saliva bacteria A: Prevotella oris (pink colonies); B: Actinomyces odontolyticus (red colonies) (BHI blood agar, anaerobic culture for 48, stereomicroscopy)

Fig. 2.47 Diffusively growing colonies of Capnocytophaga sputigena (BHI blood agar, stereomicroscope)

2.3.3.1

Preparation of Bacterial Samples Oral bacteria culture is centrifuged at 3000 r/min for 20 min, and the supernatant is removed. The pellet is washed three times with saline and a small amount of serum is added. The bacteria

are harvested by centrifugation at 3000 r/min for 20 min and the supernatant is removed. The pellet is rinsed with 2.5% glutaraldehyde (prepared with sodium cacodylate buffer) and sodium cacodylate buffer and fixed with 1% osmic acid. The sample was sealed with a series of 50%, 70%, 90%, and

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Fig. 2.48 α-Hemolytic colonies of Streptococcus gordonii (BHI blood agar, stereomicroscope)

Fig. 2.49 Candida albicans colonies (BHI blood agar, stereomicroscope)

100% ethanol dehydration and embedded with epoxy resin to make ultrathin slices.

2.3.3.2 Section Ultrathin sections are defined as sections with a thickness between 10 nm and 100 nm. The technology with which these slices are made is called

ultrathin slice technology and includes the collecting samples, fixing, rinsing, dehydrating, penetrating, embedding, sectioning, and dyeing. Compared with optical microscopy, the process of sample preparation for TEM is more sophisticated and stringent.

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Fig. 2.50 Bacterial colonies from saliva sample (BHI blood agar, stereomicroscope)

Fig. 2.51 Bacterial colonies from gingival margin sample (BHI blood agar, stereomicroscope)

2.3.3.3

Uranium Staining and Observation Professional workers use a TEM to observe the inner structure of cells via ultrathin sections (Fig. 2.65).

2.3.4

Confocal Laser Scanning Microscopy

Confocal laser scanning microscopy (CLSM) was developed in the late 1980s [1]. With the

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Fig. 2.52 Bacterial colonies from non-adhesive subgingival plaque (BHI blood agar, stereomicroscope)

Fig. 2.53 Bacterial colonies from periodontal pocket sample (BHI blood agar, stereomicroscope)

unparalleled advantages of high resolution, ease of sample preparation, dynamic recording without damaging the living cell, acquisition of threedimensional sample images through tomography

and 3D reconstruction, and spatial positioning of the target, CLSM has become widely used in almost all areas of cell research in medicine and biology (Fig. 2.66).

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Fig. 2.54 Scanning electron microscope

Fig. 2.55 Proliferating cells of β-hemolytic streptococcus (SEM)

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60 Fig. 2.56 Division phase of α-hemolytic streptococcus cells (SEM)

Fig. 2.57 Proliferating cells of Streptococcus gordonii (SEM)

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Fig. 2.58 Proliferating state of Lactobacillus fermentum cell (SEM)

Fig. 2.59 Self-aggregating cells of Rothia dentocariosa (SEM)

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62 Fig. 2.60 Cell aggregation of Capnocytophaga sputigena (SEM)

Fig. 2.61 Massive extracellular matrix of Porphyromonas gingivalis (SEM)

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Fig. 2.62 Budding cells of Candida albicans (SEM)

Fig. 2.63 Spirochetes in gingival margin plaque (SEM)

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Fig. 2.64 Transmission electron microscope

2.3.4.1 Principles of CLSM A confocal laser scanning microscope is made up of the optical system, the laser light source, the detection system, and the scanning device. The optics behind this type of imaging is a laser emitted from the light source that becomes a parallel beam of expanded diameter when it passes through the pinhole aperture, encounters the dichromatic mirror, and is reflected onto the objective lens. The light beam is reflected 90 when it hits the dichromatic mirror and is focused onto the desired focal plane on the sample when it passes through the objective lens. The fluorescence-emitting sample fluoresces in all directions under excitation from the laser. Part of the fluorescence becomes focused at the focal point of the objective once it passes through the objective lens, dichromatic mirror, and focusing lens. The fluorescent light passes through a pinhole at the focal point and can then be picked up by the detector. When a laser scans the sample point by point, the photomultiplier tube behind the pinhole receives the corresponding point-bypoint confocal optical image. Accordingly,

different focal planes within the sample and optical cross sectional images (also known as optical sectioning) can be analyzed one by one. Using computer image processing and threedimensional image reconstruction software, a high-resolution three-dimensional image can be obtained from the sample. Cell structure, cell content, and dynamic changes can be analyzed by continuous scanning on the same plane. The optical path of a confocal laser scanning microscope is shown in Fig. 2.67.

2.3.4.2

Application in Dental Plaque Research Through a special fluorescent staining, dental plaque in its natural hydration status can be studied directly. Through this process, dead and viable bacteria in dental plaque can be observed in situ, and the relationship between bacteria during the formation of dental plaque can be observed as well. Images of a single cell, a group of cells, or different levels within tissues can be obtained by scanning biofilm of a given thickness continuously using CLSM. A complete 3D structure of

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Fig. 2.65 Adenovirus AdhTR-si in HEK293 cell (TEM)

plaque can be generated by 3D image reconstruction (Fig. 2.68). Compared with SEM, CLSM requires less sample preparation steps before the composition and structure of dental plaque can be studied. During sample preparation in SEM, structural damage to the sample can accumulate during steps such as dehydration, fixing, embedding, and dyeing. Moreover, CLSM can be used to observe structures, specific molecules, and biological ionic changes in living cells. The technique can also be used to track structural changes and physiological processes within living cells over time to

detect changes under the natural state or after stimulation by certain factors. Quantitative and qualitative measurements can be made regarding the perimeter or area or a sample, average fluorescence intensity of cells, in situ determination of cellular contents, composition and distribution of lysosomes, mitochondria, endoplasmic reticulum, cytoskeleton, structural proteins, DNA, RNA, enzymes, cellular receptors, etc. Physiological signals can be dynamically monitored, including quantitative analysis of various ions (mainly calcium ions) with millisecond time resolutions using fluorescent probes.

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Fig. 2.66 Confocal laser scanning microscopy

2.4 2.4.1

Oral Microecology Techniques Methods for Measuring Microbial Growth Curves

In microbiology, growth generally refers to the increase in cell number, rather than the volume of

Fig. 2.67 The optical path of a confocal light scanning microscope

a single cell. The rate of growth, defined as the change in cell number over time, is closely related to the growth cycle, which can be divided into four phases: lag phase, logarithmic (or exponential) phase, stationary phase, and decline phase, according to the characteristics of the growth curve. The rate of growth differs during these distinct phases. We commonly use the growth curve to portray dynamic changes in bacterial cell number during the growth cycle (Fig. 2.69). Lag phase: Bacterial reproduction is slow. Logarithmic phase: Bacterial reproduction is fast, and the number of viable cells exponentially increases. Stationary phase: The rate of growth decreases gradually, while the number of dead cells increases. Decline phase: The growth gradually slows down until it stops completely, the dead cells outnumber viable ones, and the cells show abnormal phenotypes and autolysis.

2.4.1.1 Protocol Inoculate cells into fresh medium and cultivate under desired growth conditions. The number of

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Fig. 2.68 Biofilm of Streptococcus mutans (CLSM)

Fig. 2.69 Typical example of a growth curve

bacterial cells will constantly change during the growth cycle. Graph the growth curve using the number of bacterial cells as the Y-axis and the time of growth as the X-axis.

2.4.1.2 Methods Turbidimetry and viable count methods are commonly used to determine the growth curve.

1. Turbidimetry: After inoculation, measure the optical density (OD) of the cell culture during cultivation. Graph the growth curve using the OD value as the Y-axis and the cultivation time as the X-axis. 2. Viable count method: In microbial ecology research, the number of viable cells reflects dynamic changes in bacterial growth.

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Fig. 2.70 Protocol of the pouring method

Generally, the cells are plated and colonies are counted (more details in “methods for measuring colony forming units”) to measure the number of viable cells, which are the only cells in the culture that can undergo cell division and reproduce. After inoculation and a certain period of cultivation, inoculate a certain volume of the cell culture onto the agar plate. Graph the growth curve using the logarithm of the number of colonies number as the Y-axis and the growth time as the X-axis.

2.4.2

Methods for Measuring Colony Forming Units

Quantification of spatiotemporal changes in the number of organisms in an ecosystem, especially the percentage of viable cells, is an important part of microecological studies. Plate colony-counting methods have been adopted for widespread use internationally to measure the percentage of viable cells in a sample, rather than more traditional absolute viable count methods. Most commonly used methods for plating samples include spread method, drop method, and spiral plating. In addition, there are several reports of the pouring method being a viable way to measure colonyforming units.

2.4.2.1 Protocol After collecting samples from the desired locations and at the appropriate time points, the sample is transported to the laboratory under suitable conditions, suspended, and diluted to an

appropriate concentration. The diluted cells should then be quantitatively inoculated onto a suitable agar plate, placed in a 37  C incubator after the correct atmospheric conditions and culture time are determined according to the species. After incubation, every viable cell in the sample will form a visible colony. Count the number of colonies, and calculate the number of viable cell in the sample based on the degree of dilution.

2.4.2.2 Methods Plating methods for colony-counting can be divided into the spread method, drop method, pour method, and the spiral plater method, based on the approach taken to inoculate a liquid culture of bacteria onto the plate. They share the same basic protocols for sample collection, transportation, dilution, inoculation, incubation, and colony counting, but are different when it comes to the specifics of sample dilution and inoculation. For more details on the spread method, drop method, and spiral plater method, see Sect. 2.2.3.2. These are the most commonly used plate-based colonycounting methods. In addition, several published reports have shown that the pour method may be a viable option as well (Fig. 2.70). Figures 2.71, 2.72, 2.73, and 2.74 show the appearance of colonies formed by the spread method and drop method. The spiral plater uses a spiral plater instrument to automatically perform sample dilution and inoculation. After incubation the colonies grow along the spiral trajectory, and colony counting can be performed manually or using an automatic

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Fig. 2.71 Colonies on a plate prepared with the spread method (gingival marginal plaque sample)

Fig. 2.72 Colonies on a plate prepared with the drop method (plaque sample)

colony counter. For more details, see Sect. 2.2.3.2. 1. Pouring method: Mix the bacterial diluent and the 50  C soft agar in a sterile bottle. Pour the

mixture into a sterile plate (90 mm in diameter), making sure that the mixture is evenly distributed. After incubation, count the

70 Fig. 2.73 Colonies on a plate prepared with the drop method (saliva sample)

Fig. 2.74 Colonies on a plate prepared with the drop method (subgingival plaque sample)

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Fig. 2.75 Protocol for measuring adhesion strength Place medium into a known quantity of culture, and add a known quantity of bacterial suspension. After incubation under appropriate conditions, wash the adhesion medium 3–4 times with KCl buffer to remove non-adhered surface bacteria. Measure adhesion capacity using an isotope

liquid scintillation detector, and express the adhesion capacity in scintillation counts per minute (CPM). The adhesion rate (%) ¼ (experimental group CPM  negative control CPM/positive control CPM  negative control CPM) 3 100%

colonies on the surface and within the agar medium. 2. Drop method: Drop a certain volume of the appropriately diluted sample (105 CFU/ml, 104 CFU/ml, 103 CFU/ml) onto the surface of agar plates (three replicates for each concentration). Count the colonies after incubation.

strength can be improved, and the quality of the results can be improved.

2.4.3

Measurement of Adhesion Strength and Rate of Adhesion Inhibition

Microbial adhesion is the basis of colonization and pathogenesis. Measurement of adhesion strength and the rate of adhesion inhibition contribute to the understanding of mechanisms of bacterial pathogenesis and control.

2.4.3.1

Measurement of Adhesion Strength Medium Adhesion Choose slide, glass rod, hydroxylapatite, or teeth as medium for adhesion. Method Collect adhesive substance on the surface of the medium surface to evaluate adhesion by measuring the quantity of bacteria (Fig. 2.75). By adding artificial saliva or collagen solution into the culture, or by coating the medium for adhesion with saliva or collagen, adhesion

2.4.3.2

Measuring the Rate of Adhesion Inhibition Measuring the rate of adhesion inhibition allows the researcher to quantify the inhibition of bacterial adhesion by drugs or other reagents. The method is identical to that used to measure adhesion described in the previous section. However, inhibitors of adhesion must be added to the experimental group (Fig. 2.76).

2.4.4

Techniques for the Detection of Plaque Biofilm

In nature, many bacteria are attached to the surface of living and inanimate objects, where they survive and grow in the form of biofilm. Biofilms are groups of bacteria attached to a surface and enclosed in a secreted adhesive matrix and are functional, interacting, and growing bacterial communities. Dental plaque is a typical kind of biofilm formed on the tooth surface by oral microbes. Microbes inside the biofilm survive as a group with interdependence and mutual competition. They also form a complex ecological relationship. Technologies used to detect biofilm are used to analyze the natural state of bacteria, the relationship between different bacterial species

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Fig. 2.76 Protocol for measuring the rate of adhesion inhibition Adhesion inhibition rate ¼ [1  Experimental CPM  negative control CPM/positive control CPM  negative

control CPM] 3 100%. When the CPM of the experimental group is less than that of the negative control group, the rate of adhesion inhibition is defined to be 100%

and the host, pathogenic mechanisms, effects of antibacterial reagents, etc.

polysaccharides. Biofilm structure can be observed clearly with this technology (Fig. 2.81).

2.4.4.1 Biofilm Formation Assay The biofilm formation assay is the basis of a series of biofilm detection technologies and can be used to detect single species biofilm or mixed species biofilm formation as well as their structural characteristics, conditions for formation, and factors that influence their formation. The assay lays the foundation for further studies on relationships between bacterial species, mechanisms of pathogenesis, and preventative measures. The early steps of the assay involve steps that are identical to those in adhesion strength measurements, particularly when it comes to bacterial culture. However, the structure and other features of biofilms are evaluated using scanning electron microscopy (SEM) and confocal laser scanning microscope. Currently there is a commercially available micro-well plate for biofilm detection named the 96 MBECTM-Device (MBEC Biofilm Technology Ltd., Calgary, Alberta, Canada, U.S Patent), shown in Fig. 2.77. 2.4.4.2 Biofilm Detection and Analysis By scanning electron microscopy (SEM) and laser confocal microscopy, characteristic biofilm morphology, structure, and extracellular polymers can be detected. Using SEM, biofilm growth can be observed at different times and under different conditions (Figs. 2.78, 2.79 and 2.80). The combined use of fluorescence staining and laser confocal microscopy is a common method to study biofilm structure and extracellular

2.5

Oral Microbiome Techniques

The oral microbiome refers specifically to microorganisms (e.g., bacteria, archaea, fungi, mycoplasma, protozoa, and viruses) that inhabit the human oral cavity. Among them, oral bacteria make up the largest proportion of the oral microbiome and are also the most complex in organization. So far, more than 250 oral bacteria species have been isolated, cultivated, and named. Over 450 species have been identified by culture-independent approaches. These bacteria can be classified into different categories based on their Gram stain results (Gram-positive or Gram-negative bacteria), their shape (coccus, bacillus, or spirochetes), and their tolerance to oxygen (aerobic, facultative anaerobes, microaerobic, or obligate anaerobes).

2.5.1

Denaturing Gradient Gel Electrophoresis and Temperature Gradient Gel Electrophoresis

Denaturing gradient gel electrophoresis (DGGE) [2, 8] and temperature gradient gel electrophoresis (TGGE) [3] are forms of gel electrophoresis that use either a chemical gradient or a temperature gradient to separate samples as they move across an acrylamide gel. DGGE was introduced to microbial ecology by Muyzer et al. [4]. Within a short period of time, this method has become

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Fig. 2.77 Biofilm detection micro-well plate (MBECTM-Device) The MBECTM-Device has 96 removable pegs (adhesion medium) in on its plate cover that sit in the 96 micro-wells in the corresponding base. Biofilm formation on the 96 pegs can be detected at the same time, but every peg can also be removed when the biofilm on that peg must be analyzed independently

widely used in the analysis of microbial diversity in various complex samples including samples from the oral cavity. In both DGGE and TGGE, DNA fragments of the same length but with different sequences can be separated. Separation is based on the reduced electrophoretic mobility of partially melted double-stranded DNA molecules in polyacrylamide gels that contain either a linear gradient of DNA denaturants (a mixture of formamide and urea) in the case of DGGE or a linear temperature gradient in the case of TGGE. Melting domains, i.e., stretches of base-pairs with identical melting temperatures (Tm), lead to the melting of DNA fragments within discrete domains. Once a domain with the lowest Tm reaches its Tm at a particular position in the denaturing or temperature gradient gel, and that segment of the DNA double helix transitions to melted single-stranded DNA, migration of the DNA molecule will virtually stop. Sequence

variations within these melting domains cause the melting temperatures to vary, and molecules with different sequences will stop migrating at different positions in the denaturing or temperature gradient gel, therefore becoming separated. DNA bands in DGGE and TGGE profiles can be visualized using ethidium bromide, SYBR Green I, or silver stain (Fig. 2.82).

2.5.2

Sanger Sequencing

Sanger sequencing, also known as the chain termination method, is a technique for DNA sequencing based upon the selective incorporation of chain-terminating dideoxynucleotides (ddNTPs) by DNA polymer ase during in vitro DNA replication. It was developed by Frederick Sanger and Coulson [5]. It was the most widely used sequencing method for

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Fig. 2.78 Thirty-six-hour biofilm in dentin of Actinomyces viscosus (SEM)

approximately 25 years before it was replaced by next-generation sequencing methods. Classical Sanger sequencing requires a singlestranded DNA template, a DNA polymerase, a DNA primer, normal deoxynucleosidetriphosphates (dNTPs), and modified nucleotides (ddNTPs) that terminate DNA strand elongation. These ddNTPs lack a 30 -OH group that is required for the formation of a phosphodiester bond between two nucleotides, causing the extension of the DNA strand to stop when a ddNTP is added. The DNA sample is divided into four separate sequencing reactions, containing all four of the standard dNTPs (dATP, dGTP, dCTP, and dTTP), the DNA polymerase, and only one of the four ddNTPs (ddATP, ddGTP, ddCTP, or ddTTP) for each reaction. After rounds of template DNA extension, the DNA fragments that are formed are denatured and separated by size using gel electrophoresis with each of the four reactions in one of four separated lanes. The DNA bands can then be visualized by UV

light or autoradiography, and the DNA sequence can be directly read off the gel image or the X-ray film (Fig. 2.83). The ddNTPs may also be radioactively or fluorescently labeled for detection in automated sequencing machines. The four reactions can be incorporated into one reaction run, and the DNA sequence can be read from radioactive or fluorescent labels.

2.5.3

Next-Generation Sequencing

Sanger sequencing enabled scientists to elucidate genetic information from a variety of biological systems. However, wide use of this technology has been hampered due to inherent limitations of throughput, scalability, speed, and resolution. Next-generation sequencing (NGS) [9], also known as massively parallel sequencing or deep sequencing, have been developed to overcome these barriers.

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Fig. 2.79 Forty-eight hour biofilm in dentin of Actinobacillus actinomycetemcomitans (SEM)

Fig. 2.80 Biofilm from infected root canal (SEM)

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Fig. 2.81 Biofilm structure of Streptococcus mutans Under a laser confocal microscope (green laser, 504–511 nm), biofilms appear as a multilayer superimposed image due to yellow fluorescent staining of bacterial colonies and the plaque biofilm structure. The three-dimensional structure of biofilm is mushroom-like, full of channels and pores. Against the dark background, microbial colonies appear as green fluorescence, while dead bacteria show up as red fluorescence.

Fig. 2.82 Negative image of an ethidium bromide stained DGGE gel loaded with 16S rRNA gene fragments

In principle, NGS technology is based on sequentially identifying bases in a small fragment of DNA using emitted signals, while each fragment is re-synthesized from a DNA template strand. NGS proceeds in a massively parallel fashion, which enables rapid sequencing of large stretches of DNA spanning entire genomes.

2.5.3.1 Pyrosequencing Pyrosequencing is a method of DNA sequencing based on the “sequencing by synthesis” principle [6]. It relies on the detection of pyrophosphate release along with nucleotide incorporation. Sequences in each sample are tagged with a unique barcode either by ligation or by using a

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Fig. 2.83 Sanger sequencing

barcoded primer when amplifying each sample by PCR. The method amplifies DNA inside water droplets in an oil solution (emulsion PCR), and each droplet contains a single DNA template attached to a single primer-coated bead. The sequencing machine contains many picoliter-volume wells each containing sequencing enzymes

and a single bead. The sequence of the singlestranded DNA sequence can be determined by the light emitted upon incorporation of the complementary nucleotide because only one of four of the possible A/T/C/G nucleotides can complement the DNA template. The technique uses lucif erase to generate light for detection of the

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Fig. 2.84 Pyrosequencing

individual nucleotides, and the combined data are used to generate the template sequence. Pyrosequencing was commonly used for genome sequencing or resequencing during the last decade. It was widely used in the analysis of the oral microbiome. However, a limitation of the method is that the length of individual DNA reads is approximately 300–500 nucleotides, shorter than the 800–1000 that can be obtained using

Sanger sequencing. This can make the process of genome assembly more difficult, particularly for sequences containing a large amount of repeti tive DNA (Fig. 2.84).

2.5.3.2 Illumina Sequencing Illumina sequencing is based on the incorporation of reversible dye-terminators that enable the identification of single bases as they are incorporated

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Fig. 2.85 Illumina sequencing

into DNA strands [7, 10]. The basic procedure is as follows. DNA molecules are first attached to primers on a slide and amplified so that local clusters are formed. The four types (A/T/C/G) of reversible terminating nucleotides are added, and each nucleotide is fluorescently labeled with a different color and attached to a blocking group. The four nucleotides then compete for binding sites on the template DNA to be sequenced, and non-incorporated molecules are washed away. After each synthesis, a laser is applied to remove the blocking group and the probe. A detectable fluorescent color specific to one of the four bases then becomes visible, allowing for sequence identification and initiating the beginning of the next cycle. The process is repeated until the entire DNA molecule is sequenced. This technique offers some advantages over traditional sequencing methods such as Sanger sequencing. The automated nature of Illumina sequencing makes it possible to sequence

multiple strands at once and obtain actual sequencing data quickly. In addition, this method only utilizes DNA polymerase in contrast with multiple, expensive enzymes required by pyrosequencing (Fig. 2.85).

References 1. Pawley JB. Handbook of biological confocal microscopy. 3rd ed. Berlin: Springer; 2006. 2. Fischer SG, Lerman LS. DNA fragments differing by single base-pair substitutions are separated in denaturing gradient gels: correspondence with melting theory. Proc Natl Acad Sci USA. 1983;80(6):1579–83. 3. Thatcher DR, Hodson B. Denaturation of proteins and nucleic acids by thermal-gradient electrophoresis. Biochem J. 1981;197(1):105–9. 4. Muyzer G, de Waal EC, Uitterlinden AG. Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction- amplified genes coding for 16S rRNA. Appl Environ Microbiol. 1993;59(3):695–700.

80 5. Sanger E, Coulson AR. A rapid method for determining sequences in DNA by Primed synthesis with DNA polymerase. Mol Biol. 1975;94(3):441–8. 6. Ronaghi M, Uhlen M, Nyren P. A sequencing method based on real-time pyrophosphate. Science. 1998;281 (5375):363–5. 7. http://www.illumina.com/technology/next-generationsequencing/solexa-technology.html

X. Peng et al. 8. Strathdee F, Free A. Denaturing gradient gel electrophoresis (DGGE). Methods Mol Biol. 2013;1054:145–57. 9. Mardis ER. Next-generation sequencing platforms. Annu Rev Anal Chem (Palo Alto, Calif). 2013;6:287–303. 10. Meyer M, Kircher M. Illumina sequencing library preparation for highly multiplexed target capture and sequencing. Cold Spring Harb Protoc. 2010;2010(6): pdb.prot5448.

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Supragingival Microbes Xuedong Zhou, Yuqing Li, Xian Peng, Biao Ren, Jiyao Li, Xin Xu, Jinzhi He, and Lei Cheng

Abstract

This chapter mainly introduces the common microbes in supragingival plaque. Supragingival plaque refers to the plaque above the gingival margin of the tooth neck, including groove plaque, smooth surface plaque, adjacent surface plaque, and cervical margin plaque. The main bacteria of supragingival plaque are Gram-positive cocci and bacilli. With the maturity of dental plaque biofilm, the number of Gram-negative cocci, bacilli, and filamentous bacteria increased. The biological and pathogenic characteristics of Gram-positive bacteria, including Actinomyces,

Bifidobacterium, Lactobacillus, Rothia, Staphylococcus, and Streptococcus, and Gramnegative bacteria, including Leptotrichia and Veillonella, were introduced in this chapter. Gram staining, plate culture, colony, and scanning electron microscope (SEM) images of each microorganism were also provided. Keywords

Actinomyces · Bifidobacterium · Lactobacillus · Rothia · Streptococcus · Veillonella

3.1 X. Zhou (*) · L. Cheng State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China e-mail: [email protected] Y. Li · X. Peng · B. Ren · J. Li · J. He State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China X. Xu State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Department of Operative Dentistry and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China

3.1.1

Gram-Positive Bacteria Actinomyces

Actinomyces are irregular Gram-positive bacilli and are also commonly found anaerobic bacteria in oral samples. When they were first discovered, actinomycetes were believed to be fungi or were grouped as “other microorganism.” Recently, numerous studies have shown that actinomycetes have general characteristics common to bacteria and were classified as prokaryotic organisms. In the 1984 edition of the book Bergey’s Manual of Determinative Bacteriology, Actinomyces were included in the group of Gram-positive irregular bacilli. Common members of the Actinomyces genus in oral microbiology are A. israelii, A. naeslundii, A. odontolyticus, and A. viscosus.

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They are characterized by a relatively high GC content (GC content is the percentage of nitrogenous bases on a DNA molecule that are either guanine or cytosine) in their DNA, ranging from 57 to 69% (Tm method). The type species of this genus is A. bovis. The shape and size of bacterial cells can vary greatly, but are often found to be irregular branched bacilli with a diameter of 0.2–1.0 μm and a length of about 5.0–10.0 μm. The cells are usually rod-shaped, but can occasionally be clubshaped with irregular arrangements including single, paired, chain, clusters, and fence-shaped. The cells produce no spores, show no motility, and also do not produce conidia. The main distinguishing characteristic of Actinomyces cells is that the cell wall does not contain DAP and glycine. There are differences at the species level regarding oxygen sensitivity, but a primary culture of Actinomyces requires an anaerobic environment. Spider-like micro-colonies or branched mycelia can form on agar plates after 18–24 h of incubation. These typical spider web-shaped colonies can help identify the genus of bacteria. All strains of actinomycetes can ferment glucose and fructose to produce acid, without producing gas. Other tests, including fermentation of raffinose, xylose, cellobiose, laetrile, ribose, and salicin; catalase production; reduction test using nitrate and nitrite; urea hydrolysis; or gelatin hydrolysis, can help identify different species. Actinomycetes are normal members of the oral flora and are the dominant bacteria in dental plaque [1]. A. israelii, A. naeslundii, A. odontolyticus, and A. viscosus can be detected in human dental plaque, dental calculus, and saliva. The main colonization site of A. mai is in the gingival sulcus. Clinical and epidemiological investigations indicate that A. israelii can cause actinomycosis, conjunctivitis, and lachrymal and other diseases of the face, neck, lung, and abdomen. A. naeslundii and A. mai can be detected in clinical samples of gingivitis, periodontitis, pulp periapical infection, and pericoronitis. A. viscosus is suspected to be a cariogenic bacterium.

3.1.1.1 Actinomyces israelii A. israelii are Gram-positive irregular bacilli (Figs. 3.1, 3.2, and 3.3). The culture atmosphere requires CO2, as the culture grows poorly or not at

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all under ordinary atmospheric environment. Bacterial growth can be inhibited by 4%–6% NaCl, 20% bile, or 0.005% crystal violet. Characteristic colonies are shown in Figs. 3.4, 3.5, and 3.6. The DNA G + C content ranges from 57% to 65% when analyzed using the Tm method. The type strain is ATCC12102 (WVU46, CDCX523, W855). The main habitat of this species is the oral cavity. A. israelii often colonizes the tonsils and plaque, but can also be detected in the human gut and the female reproductive tract. This bacterium is mainly a pathogen of the face and neck and causes lung and abdominal actinomycosis. It can also infect the lachrymal sac and conjunctiva. It is detected in oral mixed infections such as gingivitis, periodontitis, and pericoronitis, but the link between A. israelii and the infections is unclear. A. israelii always display branching rods and short filamentous with no spores; Gram stain is negative. Young colonies of A. israelii are typical filamentous micro-colonies. Mature colonies are characteristically less than 2 mm in diameter, raised, white, opaque, and molar-shaped. The species shows no β-hemolytic reaction on blood agar.

3.1.1.2 Actinomyces naeslundii A. naeslundii is a Gram-positive irregular bacillus (Figs. 3.7 and 3.8). Under aerobic conditions without CO2, the culture may not grow. However, cultures can be grown in an anaerobic environment without CO2. Some strains can be grown at 45  C. Growth can be inhibited by 6% NaCl. Characteristic colonies are shown in Figs. 3.9 and 3.10, and broth culture is shown in Fig. 3.11. The DNA G + C content is approximately 63%–69% using the Tm method. The type strain is ATCC12104 (NCTC10301, WVU45, CDCX454). A. naeslundii is a member of the normal human oral flora and can be found on the tonsils and in dental plaque. It can take part in mixed bacteria infections and is one of the pathogenic bacteria in root caries. It is often detected in clinical specimens of periodontitis or infected root canals, but the pathogenesis is unclear. This bacterium can lead to actinomycosis at many locations, such as the face, neck, chest, abdomen, and eye, and can cause infection of the female genital tract and knee joint empyema.

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Fig. 3.1 A. israelii cells (Gram stain)

Fig. 3.2 A. israelii cells (SEM)

A. naeslundii cells are mainly irregular branched rods or short filaments without spores. The cells are Gram-negative.

Young colonies of A. naeslundii can appear as filiformed micro-colonies. Mature colonies are convex or flat and rough or smooth without center

84 Fig. 3.3 A. israelii branch cells (SEM)

Fig. 3.4 A. israelii colonies (BHI blood agar)

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Fig. 3.5 A. israelii molarshaped colonies (stereomicroscope)

Fig. 3.6 A. israelii colonies detached from plaque samples (stereomicroscope)

sag. The colonies show no hemolytic reaction on blood agar. The culture is muddy in broth culture and sticks to the flask wall.

3.1.1.3 Actinomyces odontolyticus A. odontolyticus is a Gram-positive irregular bacillus (Figs. 3.12 and 3.13). It cannot grow in

an anaerobic environment unless CO2 is added. Growth can be inhibited by 5%–20% bile or 0.005% crystal violet. Characteristic colonies are shown in Figs. 3.14, 3.15, and 3.16. The DNA G + C content is 62% when analyzed by Tm method. The type strain is ATCC17929 (NCTC9935, WVU867, CDCX363).

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Fig. 3.7 A. naeslundii cells (Gram stain)

Fig. 3.8 A. naeslundii cells (SEM)

Dental plaque and dental calculus are the main habitats. This species is often involved in eye infections and is occasionally seen in advanced

actinomycosis. The relationship between A. odontolyticus and periodontosis and dental caries remains to be confirmed.

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Fig. 3.9 A. naeslundii colonies (BHI blood agar)

Fig. 3.10 A. naeslundii colonies (stereomicroscope)

3.1.1.4 Actinomyces viscosus A. viscosus is a Gram-positive irregular bacillus (Figs. 3.17, 3.18, and 3.19). Primary cultures grow under anaerobic conditions and CO2 can stimulate its growth. Subcultures can grow in

common atmospheric conditions. Some cells can grow at temperatures up to 45  C. Characteristic colonies are shown in Figs. 3.20 and 3.21. The DNA G + C content ranges from 59% to 70%

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Fig. 3.11 A. naeslundii liquid culture (BHI broth)

Fig. 3.12 A. odontolyticus cells (Gram stain)

using the Tm method. The type strain is ATCC15987 (WVU745, CDCX603, A828). Humans’ and other warm-blooded animals’ oral cavities, including subgingival plaque, transparent plaque, and dental calculus, are the main sites of growth. A. viscosus is one of the cariogenic bacteria found in root caries and is related to periapical infections, dacryosolenitis, and abdominal and faciocervical actinomycosis.

3.1.2

Bifidobacterium

Bifidobacterium is a genus of bacteria with various forms and non-motile; they are Grampositive, non-sporulating, anaerobic bacilli. These bacteria were first isolated from infant feces and attracted attention because of their important physiological significance to the host organism. Species that are important human gut

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Fig. 3.13 A. odontolyticus cells (SEM) A. odontolyticus cells are mainly irregularly rod-shaped. Ball-like rods, branched rods, or filiform cells can also be observed. The cells are Gram-positive

Fig. 3.14 A. odontolyticus colonies (BHI blood agar)

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Fig. 3.15 Red colonies of A. odontolyticus (stereomicroscope)

Fig. 3.16 A. odontolyticus colonies (stereomicroscope) A. odontolyticus can generate filiformed micro-colonies. The mature colonies on the surface of BHI agar measures are less than or equal to 2 mm in diameter. They are white or gray-white and opaque and do not show a central sag or

filamentous edge. The important identifying feature of this species is that the colony turns dark red after 2 days of culture on the surface of blood agar under an anaerobic environment. The color clears after the cells are placed at room temperature

bacteria include B. bifidum, B. infantis, B. adolescentis, and B. longum. Bacteria isolated from the oral cavity belonging to the Bifidobacterium spp. include mainly B. dentium, B. breve, B. inopinatum, and B. denticolenu. The

percent G + C in Bifidobacterium DNA ranges from 55% to 67% when analyzed by the Tm or Bd method. The type species is B. bifidum. The bacterial cells are short and thin, with pointed ends, and are irregular. They also appear

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Fig. 3.17 A. viscosus cells (Gram stain)

Fig. 3.18 A. viscosus cells (SEM)

as long cells with many branches and slightly branching spoon-shaped cells. Cells are arranged as single cells, chains, polymer-shaped, V-shaped, or palisade-shaped. Their distinct cell morphology can be helpful in differentiating

bacteria belonging to this genus. For example, B. bifidum appear as flask-shaped cells, while B. asteroides are star-shaped. All members of this genus are Gram-positive.

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Fig. 3.19 A. viscosus cells (SEM) A. viscosus cells are mainly irregular short- or moderate-length rod-shaped without spores. Branch rods or short filiform also can be seen; Gram stain is positive

Fig. 3.20 A. viscosus colonies (BHI blood agar)

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Fig. 3.21 A. viscosus colonies (stereomicroscope) A. viscosus can generate filiformed micro-colony. The mature colony do not have center sag. The typical colony is big and sticky

Bifidobacterium are anaerobes and most strains cannot grow under 90% air and 10% CO2. Colonies formed on agar plates are convex, creamy or white, glossy, smooth, neat-edged, sticky, and soft. The main terminal acid products in liquid culture medium containing glucose are acetic acid and lactic acid, but a few species also make formic acid and succinic acid. However, no butyric acid or propionic acid is formed, and no CO2 is generated (with the exception of gluconate degradation). Bifidobacterium can ferment carbohydrates to produce acid and generally do not reduce nitrate or produce urease. They test positive using the catalase test.

3.1.2.1 Bifidobacterium dentium Originally, B. dentium was isolated from pus specimens and named B. appendicitis. Later, researchers isolated similar bacteria from the adult dental caries, feces, and vagina, and they were then named A. eriksonii or grouped into B. adolescentis. According to later research, these bacteria make up an independent branch on the phylogenetic tree, and the species was named B. dentium [2] in the 1970s. The percent G + C in their DNA is 61% (Tm method), and the

type species is ATCC27534 (reference strains B764). The cells are anaerobic Gram-positive irregular bacilli (Fig. 3.22). Some strains are resistant to oxygen in the presence of CO2. The optimum temperature for the growth of this bacterium is 37–41  C, and the optimum pH value ranges from 6.5 to 7.0. TPY culture medium supplemented with neomycin, kanamycin, and various salt solutions is commonly used as the selective culture medium. Characteristic colonies are shown in Figs. 3.23, 3.24, and 3.25. B. dentium is biochemically active. It can ferment D-ribose, L-arabinose, lactose, sucrose, cellobiose, trehalose, raffinose, melibiose, mannitol, salicin, starch, galactose, maltose, fructose, xylose, mannose, and glucose to produce acid, but cannot ferment sorbitol and inulin. It cannot reduce nitrate and tests negative for both the urease test and the catalase test. The distribution of the bacteria in the oral cavity and its pathogenic mechanism are not clearly characterized.

3.1.2.2 Bifidobacterium breve B. breve is an anaerobic Gram-positive irregular bacillus [3] (Figs. 3.26, 3.27, and 3.28). Its culture

94 Fig. 3.22 B. dentium cells (Gram stain)

Fig. 3.23 B. dentium colonies (BHI blood agar)

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Fig. 3.24 B. dentium colonies (B. dentium selective agar)

Fig. 3.25 B. dentium colonies (stereomicroscope) B. dentium forms colonies that can be described as spherical, lustrous, smooth, convex, gray-white, sticky, and soft when plated on BHI blood agar and Bifidobacterium selective culture medium

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96 Fig. 3.26 B. breve cells (Gram stain)

Fig. 3.27 B. breve cells (SEM)

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Fig. 3.28 B. breve cells (SEM) B. breve cells are thin and short bacilli and non-sporulated, non-motile, and Gram-positive

Fig. 3.29 B. breve colonies (BHI blood agar)

characteristics are similar to those of B. dentium, and characteristic colonies are shown in Figs. 3.29 and 3.30. The percent G + C in

B. breve DNA is 58% by the Tm method, and the type species is ATCC15700.

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Fig. 3.30 B. breve colonies (stereomicroscope) B. breve on agar plates form colonies that are spherical, smooth, translucent, graywhite, sticky, and soft

B. breve can ferment D-ribose, lactose, and raffinose, but does not ferment L-arabinose and starch.

3.1.3

Lactobacillus

Lactobacillus is a group of anaerobic or microaerobic Gram-positive bacilli that do not produce spores. Bacteria of this genus form part of the normal flora of the human oral cavity and intestinal tract. This genus is cariogenic, as they are detected in decayed oral cavity materials. This genus includes 44 species according to Bergey’s Manual of Systemic Bacteriology and also contains 7 subspecies. The common species found in the in oral cavity include L. acidophilus, L. salivarius, L. plantarum, L. fermentum, L. brevis, and L. casei. The percentage G + C in the DNA is 40% (by either Tm method or Bd method). The type species is German type Lactobacillus. The shapes and sizes of the bacterial cells can vary greatly. They can be vimineous, stubbed, bent, bacilliform, clavate, club-shaped, etc. However, most Lactobacillus cells are fairly regular with no branching. The cells are square or obtuse

at the ends when compared with other Grampositive non-sporulating bacilli. They produce no spores and no capsules and stain Grampositive. Surface culture on a solid medium is best when performed in anaerobic or microaerophilic conditions. However, some species must be cultivated under anaerobic conditions. Some members of this genus can grow within the 15  C to 45  C range, in the presence of 5%– 10% CO2, which promotes bacterial growth. As acid-producing bacteria, low pH Rogosa agar is the culture medium of choice for many strains of Lactobacillus. The optimum pH for growth is 5.5–6.2. The colonies are round, white or gray, and transparent or non-transparent with a diameter from pinprick-sized to 2 mm on the agar surface. Smooth colonies are soft, raised, and lustrous, and the edge of the colony is neat. The surface of rough colonies is dry, flat, and lackluster, and the edge is not neat. The bacteria normally do not produce pigment. Lactobacilli can ferment glucose to produce acid and are negative for catalase, urease, and cytochrome enzyme. They do not produce benzpyrole, cannot reduce nitrate, cannot

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Fig. 3.31 L. acidophilus cell (Gram stain)

hydrolyze gelatin, and cannot produce H2S. Sugar fermentation test and arginine hydrolysis test can help identify the genus of bacteria. The bacteria can promote the development of tooth decay, as its detection is significantly increased in deep caries material.

3.1.3.1 Lactobacillus acidophilus These are Gram-positive regularly shaped bacteria (Figs. 3.31 and 3.32) that grow well under anaerobic conditions. Cells grow well at 45  C, but do not grow at 15  C. BHI blood agar and Rogosa agar are the commonly used media, and the latter is a selective medium. Characteristic colonies are shown in Figs. 3.33, 3.34, 3.35, 3.36, and 3.37. L. acidophilus are obligate homofermentative bacteria [4]. They can produce D- or L-lactic acid and ferment glucose, and most strains can also ferment starch. L. acidophilus cannot produce ammonia from arginine. The G + C content in its genomic DNA is 32%–37% (by Tm method or Bd method). The type strain is ATCC4356. L. acidophilus is mainly isolated from the gastrointestinal tract of humans and animals, human mouths, and the human vagina. L. acidophilus can be isolated from a minority of neonates’ mouths. As the children grow older, the number

of bacteria found in their mouth decreases gradually, until at age 2, only a very small amount of L. acidophilus can be detected. The main site of colonization of L. acidophilus in the mouth is dental plaque; it is relatively rare in saliva, on the tongue, or in the gingival sulcus. As it is often found in material from deep caries, L. acidophilus is believed to be associated with the development of dental caries.

3.1.3.2 Lactobacillus casei L. casei was originally divided into four subspecies: L. casei subsp. casei, L. casei subsp. pseudoplantarum (now known as L. paracasei subsp. paracasei), L. casei subsp. rhamnosas (now known as L. rhamnosus), and L. casei subsp. toleons (now known as L. paracasei subsp. tolerans). A newly added subspecies is L. casei subsp. alactosus (now known as L. paracasei subsp. paracasei). The cells stain Gram-positive (Figs. 3.38 and 3.39). Cultures grow well under anaerobic conditions [5]. BHI blood agar and Rogosa agar are the commonly used media, while the latter is a selective medium. Characteristic colonies are shown in Figs. 3.40, 3.41, and 3.42. The percent G+ C in its genomic DNA is 45%–47%

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Fig. 3.32 L. acidophilus cell (SEM) Cells of L. acidophilus measure 0.3–0.4  1.5–4.0 μm in size. They are arranged as single cells, in pairs, or as short chains. They have rounded ends and no muramic acid in the cell wall. Cells stain Grampositive

Fig. 3.33 Colonies of L. acidophilus (BHI blood agar)

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Fig. 3.34 Colonies of L. acidophilus (Rogosa agar)

Fig. 3.35 Rough colonies of L. acidophilus (stereomicroscope)

(Bd method). The type strain is ATCC393 (L. casei subsp. casei). The main colonization sites of L. casei are the human intestine, mouth, and vagina. The bacteria can also be detected in milk and other dairy

products. The main site of colonization in the mouth is in dental plaque. As it is often found in material from deep caries, it is believed to be a pathogen of dental caries.

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Fig. 3.36 Smooth colonies of L. acidophilus (stereomicroscope)

Fig. 3.37 Surface features of the L. acidophilus rough colonies (stereomicroscope) L. acidophilus colonies can be separated into rough and smooth type. Hair-like structures can be observed under the stereomicroscope. Colonies do not produce pigment

L. casei is a facultative heterofermentative bacterial species. Other than the subspecies L. casei subsp. rhamnosas, other subspecies grow well at 15  C, but do not grow at 45  C.

3.1.3.3 Lactobacillus fermentum L. fermentum is a Gram-positive bacterium (Figs. 3.43, 3.44, and 3.45). Commonly used media to culture this species are BHI blood agar and Rogosa agar, where the latter is a selective

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Fig. 3.38 L. casei cell (Gram stain)

Fig. 3.39 L. casei cell (SEM) L. casei cells measure 0.7–1.1 μm  2.0–4.0 μm  1.5–5.0 μm and are arranged in a chain. Individual cells usually have rounded ends. Cells stain Gram-positive

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104 Fig. 3.40 colonies of L. casei (BHI blood agar)

Fig. 3.41 colonies of L. casei (Rogosa agar)

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Fig. 3.42 cheese colonies of L. casei (stereomicroscope) L. casei can form milky white colonies about 1 mm in diameter. Colony morphology is rounded, smooth, and non-transparent on BHI blood agar and Rogosa agar

Fig. 3.43 L. fermentum cell (Gram stain)

medium. Characteristic colonies are shown in Figs. 3.46, 3.47, and 3.48. Since this species can be detected in the human mouth and in yeast, dairy products, sourdough, and fermented plants, L. fermentum is considered to be related to the occurrence of oral infectious

diseases such as dental caries and root canal infections. L. fermentum usually grows well at 45  C and does not grow at 15  C. Obligate heterofermentative bacteria. Calcium pantothenate, nicotinic acid, and thiamine are required for growth,

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Fig. 3.45 L. fermentum cell (SEM) The L. fermentum cell is 0.5–0.9 μm in diameter, but its length can vary quite significantly. The ends of the cells are square or obtuse. Most bacterial cells are arranged as a single cell or in pairs. They do not produce spores and are non-motile. L. fermentum stains Gram-positive

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Fig. 3.46 colonies of L. fermentum (BHI blood agar)

Fig. 3.47 Concentric circle structure of L. fermentum colonies (stereomicroscope)

but vitamin B2, pyridoxal, and folic acid are not required. The percent G + C in its genomic DNA is 52%–54% (analyzed by the Bd method or Tm method). The type strain is ATCC14931.

3.1.4

Rothia

Rothia are Gram-positive facultative anaerobic bacilli that do not produce spores.

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Fig. 3.48 Surface of smooth colonies and rough, lined, myxoid colonies of L. fermentum (stereomicroscope) L. fermentum can form gray-white colonies about 1 mm in diameter. The shape of the colony is convex or slightly convex, the surface is smooth, and the colonies are non-transparent on BHI blood agar. A striking feature of

L. fermentum colonies is the concentric circle structure at the center of the colony when observed under stereomicroscope. The smooth spherical colonies are convex and have neat edges, while rough myxoid colonies are slightly convex, have irregular edges, and have a granular surface

R. dentocariosa is the type species of the Rothia genus. Rothia dentocariosa is a Gram-positive bacillus that does not produce spores. The characteristic appearance of the cells is shown in Figs. 3.49, 3.50, and 3.51. The G+ C content of its DNA is 47–57% (Tm method). The type strain is ATCC17931. The bacteria cells can be spherical, pleomorphic (similar to Corynebacterium diphtheria), or filamentous. Cells stain as Gram-negative. The cell diameter is generally 1.0 μm, but cells are irregular in shape, and the ends can reach a diameter of 5.0 μm. Cells appear almost filamentous following culture on solid media, while they appear spherical in broth media. Cells become almost completely spherical after growing for 2–3 days in stale broth media, but the coccoid morphology can be easily altered. R. dentocariosa does not produce spores or a capsule. They are non-motile and are not acid

tolerant. Bacterial cells as viewed by SEM are shown in Figs. 3.50 and 3.51. These bacteria are facultative anaerobes. They grow well in an aerobic environment, although primary cultures require incubation under anaerobic conditions (80% N2, 10% H2, 10% CO2). The optimum growth temperature is 35–37  C. When cultured for 18–24 h under anaerobic conditions, young colonies are always filamentous and appear as spider-like colonies. When inoculated under aerobic conditions, young colonies can reach a diameter of 1 mm. The colony surface is smooth or grainy and often shows an umbrella edge. After 2 d of culture, mature colonies can reach a diameter of 2–6 mm, with a milky, glossy, and smooth appearance (Fig. 3.50). Smooth colonies and rough colonies can co-exist on the same agar plate, which may also show loose, crumbly, or sticky colonies. A handful of roughtype colonies may also take on a dry coil shape.

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Fig. 3.49 R. dentocariosa cells (Gram stain)

Fig. 3.50 R. dentocariosa cells (SEM)

Characteristic colonies are shown in Figs. 3.52 and 3.53. The main acid product is lactic acid, and R. dentocariosa does not produce propionic acid when inoculated into PYG broth. R. dentocariosa

can ferment glucose, maltose, sucrose, trehalose, fructose, and salicylate to produce acid. Cells test positive for catalase, but do not produce indole. They are able to reduce nitrate and nitrite and can hydrolyze esculin, starch, and casein. It is unclear

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Fig. 3.51 R. dentocariosa cells (SEM) The bacteria cells are spherical, similar to Corynebacterium diphtheriae in morphology, but can also be filamentous; however, they mostly exist as mixed morphologies. The cell diameter is generally 1.0 μm, but is irregular, as the apical ends of the rod can reach a diameter of 5.0 μm. The culture is almost filamentous on solid media and spherical in broth medium. It is a Gramnegative species

Fig. 3.52 R. dentocariosa colonies (BHI blood agar)

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Fig. 3.53 R. dentocariosa colonies (Stereomicroscope) When inoculated under aerobic conditions, young colonies can reach a diameter of 1 mm. The colony surface is smooth or grainy and often presents an umbrella-like edge. After 2 days of culture, mature colonies can reach a diameter ranging from 2 to 6 mm, with a milky, glossy, and smooth appearance. Smooth colonies and rough

colonies can exist simultaneously on the same agar plate, while colonies can also appear loose, crumbly, or sticky. A small number of rough-type colonies may also exhibit a dry coil shape. When cultured for 18–24 h under anaerobic conditions, young colonies are always filamentous and appear as spider-like colonies

whether R. dentocariosa can hydrolyze gelatin. It is positive for urease activity and can produce H2S in triple sugar iron agar. R. dentocariosa are detected in the human oral cavity. Its main sites of colonization are the saliva and subgingival plaque. They are non-pathogenic members of the human oral microflora and have no confirmed relationship to oral infections. As an opportunistic pathogen, it has been detected from in endocarditis samples [6] and other clinical infected specimens.

early 1980s, analysis of biochemical reactions (e.g., mannitol fermentation) and cellular components (e.g., the availability of coagulase) resulted in the division of the Staphylococcus genus into subgroups of pathogenic and non-pathogenic species. In Bergey’s Manual of Systematic Bacteriology [7], members of the Staphylococcus genus are divided into 4 groups and 19 species based on cell wall composition and nucleic acid analysis. The cells of Staphylococcus are characterized as being spherical (0.5–1.5 μm in diameter), Gram-positive, aflagellar, and non-motile cocci organized as single cells, pairs, tetrads, and clusters. However, they tend to form botryoid clusters. As is the case with other Gram-positive bacteria, peptidoglycan and teichoic acid are the two main components of the Staphylococcus cell wall. The genomic G + C content of this genus ranges from 30% to 39%.

3.1.5

Staphylococcus

Members of the Staphylococcus genus are Grampositive cocci and belong to the Micrococcus family. The organisms are widely spread in the environment. Early on, three species were isolated from clinical samples: S. aureus, S. epidermidis, and S. saprophyticus. In the

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Fig. 3.54 S. epidermidis cells (Gram stain)

Staphylococci are facultative anaerobes, with the exception of S. saccharolyticus, which is an anaerobic bacterium. The optimum temperature for the growth of Staphylococcus is between 18  C and 40  C. Most members of the Staphylococcus genus can grow in media containing 10% NaCl. The type species of Staphylococcus is S. aureus. Staphylococcus epidermidis is a Grampositive bacterium. Their cell wall teichoic acid formed by polymerized glycerol, glucose, and N-acetyl glucosamine. Cellular characteristics are shown in Figs. 3.54 and 3.55. The G + C content of its genomic DNA ranges from 30% to 37%, and the type strain is ATCC14990. S. epidermidis is a facultative anaerobe, but also grows well under aerobic conditions (Figs. 3.56 and 3.57). Culture conditions for S. epidermidis are similar to those of S. aureus (see 5.1.1), but S. epidermidis grows slowly in medium with 10% NaCl. S. epidermidis mainly colonizes human skin and is a health concern due to its involvement in hospital-acquired infections [8]. The organisms are frequently detected in saliva and dental plaque and are thought to be associated with periodontitis, acute and chronic pulpitis, pericoronitis, dry socket, and angular stomatitis. S. epidermidis is

sensitive to novobiocin, with the minimum inhibitory concentration at no more than 0.2 mg/L.

3.1.6

Streptococcus

The Streptococcus genus makes up the most common Gram-positive facultative anaerobic cocci, and its members are the predominant bacteria in the oral cavity. The name Streptococcus was given because the bacteria belonging to this genus always arrange themselves into chains. In clinical bacteriology, members of Streptococcus are divided into three categories based on their ability to induce hemolysis: α-hemolytic Streptococcus (also known S. viridans), β-hemolytic Streptococcus, and γ-hemolytic Streptococcus (also known non-hemolytic Streptococcus). In the 2004 edition of Bergey’s Manual of Systematic Bacteriology, 89 species were attributed to the Streptococcus genus. The most prevalent species in the oral cavity are S. salivarius, S. sanguinis, S. mutans, S. sobrinus, S. oralis, S. mitis, and S. gordonii.

3.1.6.1 Streptococcus salivarius S. salivarius is a Gram-positive coccus (Figs. 3.58 and 3.59). Most strains of S. salivarius belong to

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Fig. 3.55 S. epidermidis cells (SEM) S. epidermidis cells are spherical (0.5–1. 5 μm in diameter) and Grampositive. The cocci organize into tetrads and clusters. Single cells are occasionally observed

Fig. 3.56 Colonies of S. epidermidis incubated on agar plate

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Fig. 3.57 Colonies of S. epidermidis (stereomicroscope) Colonies of S. epidermidis are round, raised, shiny, and gray and have complete edges. The diameter is approximately 2.5 mm. They usually do not produce a hemolytic zone. Strains that can produce mucus form translucent sticky colonies

Fig. 3.58 S. salivarius cells (Gram stain)

the Lancefield group K. Their genomic G + C content is 39%–42%, and the type strain is ATCC7073. S. salivarius are facultative anaerobes, but the optimal atmosphere condition for bacterial cultures should contain a low percentage of oxygen with 5–10% carbon dioxide. S. salivarius grows quickly at 37  C, although it can also

grow at 45  C. Cultures require nutrient-rich complex media, such as TS or TPY. The final pH of glucose broth incubated with S. salivarius usually falls between pH 4.0 and 4.4. Colony morphology is shown in Figs. 3.60, 3.61, 3.62, and 3.63. Biochemical Reactions S. salivarius can ferment glucose, sucrose, maltose, raffinose, inulin,

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Fig. 3.59 S. salivarius cells (SEM) The cells of S. salivarius are spherical or oval (0.8–1.0 μm in diameter) Gram-positive cocci organized in short or long chains

Fig. 3.60 α-hemolytic zone of S. salivarius incubated on blood BHI agar plate

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Fig. 3.61 Rice ball-like colonies of S. salivarius incubated on MS agar

Fig. 3.62 Colonies of S. salivarius observed under stereomicroscope (incubated on blood BHI agar plate)

salicin, trehalose, and lactic acid. It cannot ferment glycerol, mannitol, sorbitol, xylose, and arabinose. Most strains can hydrolyze esculin

and urea, but not arginine. Meanwhile, most strains can produce acetoin from glucose.

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Fig. 3.63 Colonies of S. salivarius isolated from saliva (stereomicroscope) The ability to synthesize extracellular polysaccharides determines whether colonies of S. salivarius are smooth or rough. On agar plates with sucrose, most strains synthesize soluble fructan and form sticky rice balllike colonies, a characteristic that can be used to identify S. salivarius. Very few strains produce α- or β-hemolytic zones when incubated on agar containing starch or horse blood

Fig. 3.64 S. sanguinis cells (Gram stain)

Colonization Characteristics S. salivarius is mainly isolated from the oral cavity of human and animals, and it is part of the normal flora of tongue and saliva microbial communities. Moreover, S. salivarius is also detected in fecal and blood samples of endocarditis patients. Research

on gnotobiotic animals showed that S. salivarius is cariogenic.

3.1.6.2 Streptococcus sanguinis S. sanguinis is a Gram-positive coccus (Figs. 3.64 and 3.65). Most strains of S. sanguinis belong to

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Fig. 3.65 S. sanguinis cells (SEM) S. sanguinis cells are spherical or oval (0.8–1.2 μm in diameter), Gram-positive cocci that are organized in medium or long chains. Occasionally, the bacteria are rod-shaped or pleomorphic

Fig. 3.66 Colonies of S. sanguinis incubated on blood BHI agar plate

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Fig. 3.67 Smooth colonies of S. sanguinis incubated on MS agar plate

Lancefield group H. The genomic G + C content is between 40% and 46% [9], and the type strain is ATCC10556 (NCTC 7863). S. sanguinis is a facultative anaerobe and the optimum atmospheric condition for cultures should contain 5–10% carbon dioxide. S. sanguinis grows quickly at 37  C, and it cannot grow at 45  C. Bacterial cultures require nutrientrich complex media. Blood BHI and TPY media are used for S. sanguinis isolation, and MS medium is used for selective culture (Figs. 3.66, 3.67, and 3.68). Biochemical Reactions S. sanguinis can ferment glucose, maltose, sucrose, trehalose, and salicin to produce acid. Occasionally, it also ferments inulin, raffinose, and sorbitol. S. sanguinis does not ferment xylose, arabinose, glycerol, and mannitol. More than 50% of S. sanguinis strains hydrolyze esculin. The final pH of glucose broth incubated with S. sanguinis is approximately pH 4.6–5.2. Ammonia is produced from arginine hydrolysis and H2O2 synthesis can be used to distinguish this bacterium from S. mutans.

Colonization Characteristics S. sanguinis is the main component of dental plaque and is only isolated from oral cavities with erupted teeth. This organism is considered to be helpful to the colonization and reproduction of S. mutans due to its ability to synthesize PABA. Meanwhile, S. sanguinis is regarded as a key probiotic in the oral ecosystem and is associated with healthy periodontal tissues, owing to its ability to synthesize H2O2. S. sanguinis colonies are either smooth or rough, and the diameter of colonies is between 0.7 and 1.0 mm. When incubated under aerobic conditions, an α-hemolytic zone can be observed around the colonies of most strains, while a β-hemolytic zone can be observed around the colonies of a few strains. A large number of strains that make up S. sanguinis harbors the capacity to produce extracellular polysaccharides above or surrounding their colonies. A liquid-like structure composed of polysaccharides can be observed on these colonies. S. sanguinis colonies grown on agar containing a high concentration of sucrose are sticky, hard, and rough. These colonies, with a ground-glass appearance, seem

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Fig. 3.68 α-Hemolytic zone of S. sanguinis incubated on blood BHI agar plate (stereomicroscope)

to stretch into the surrounding agar. However, colonies on agar with low sucrose concentration or without sucrose are round, soft, and smooth.

3.1.6.3 Streptococcus gordonii S. gordonii was classified as S. sanguinis serotype II in the past. However, S. gordonii lacks the IgA1 protease. It is a Gram-positive coccus (Figs. 3.69, 3.70, and 3.71). The cell wall components are mainly glycerol, teichoic acid, and rhamnose, while its peptidoglycan type is Lys-Ala. The genomic G + C content is 40%– 43%, and the type strain is ATCC10558 (NCTC 7865). S. gordonii is a facultative anaerobe. Zones of α- or γ-hemolysis can be observed on blood agar, and a green hemolytic zone can be seen on chocolate agar. This species includes three biotypes and all three do not synthesize catalase. Colonies are shown in Figs. 3.72, 3.73, and 3.74. Colonization Characteristics S. gordonii mainly inhabits the oral cavity and pharynx.

The α-hemolytic zone can be observed surrounding S. gordonii colonies incubated on blood agar. Some strains harbor the ability to synthesize extracellular polysaccharides on top of or surrounding their colonies. This layer of polysaccharides appears as a liquid-like structure. In addition, colonies grown on agar containing high sucrose concentration are sticky, hard, and rough. These colonies have a ground-glass appearance and seem to stretch into the surrounding agar. However, colonies grown on agar with low sucrose concentration or without sucrose are round, soft, and smooth.

3.1.6.4 Streptococcus mutans S. mutans, S. sobrinus, S. rattus, S. ferus, S. cricetus, and S. macacae are collectively known as mutans streptococci. These bacteria formerly belonged to serotypes a, b, c, d, e, f, g, or h of S. mutans. Their genomic G + C content is between 36% and 38% [10] and the type strain is ATCC25175. S. mutans is Gram-positive and its colony morphology is shown in Figs. 3.75, 3.76, 3.77, 3.78, and 3.79.

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Fig. 3.69 Spherical cells of S. gordonii (Gram stain)

Fig. 3.70 Spear-shaped cells of S. gordonii (Gram stain)

S. mutans is a facultative anaerobe, but the optimal atmospheric condition for cultures should be anaerobic or contain only a low percentage of oxygen with 5–10% carbon dioxide. S. mutans grows quickly at 37  C and some strains can grow at 45  C. S. mutans cultures require nutrient-rich complex media, such as TS and TPY. MSB is

used for selective culture (Figs. 3.80, 3.81, and 3.82). Cells tend to clump or attach to the bottom of the tube when incubated in glucose broth (Fig. 3.83), and the final pH of bacterial culture in glucose broth is usually between pH 4.0 and 4.3.

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Fig. 3.71 Proliferating cells of S. gordonii (SEM) S. gordonii cells are spherical or spear-shaped and organize into chains. The cells are non-motile and non-sporulating

Fig. 3.72 α-Hemolytic zone of S. gordonii incubated on blood BHI agar

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Fig. 3.73 γ-Hemolytic zone of S. gordonii incubated on blood BHI agar

Fig. 3.74 Sticky colonies of S. gordonii incubated on blood BHI agar (stereomicroscope)

Biochemical Reactions Most strains ferment mannitol, sorbitol, raffinose, lactose, inulin, salicin, mannose, and trehalose to produce acid, but

do not ferment arabinose, xylose, glycerol, and melezitose. S. mutans hydrolyzes esculin, but not

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Fig. 3.75 Spherical cells of S. mutans (Gram stain)

Fig. 3.76 Long chainshaped cells of S. mutans (Gram stain)

arginine, hippurate, and gelatin. S. mutans does not produce H2O2. Colonization Characteristics S. mutans is mainly isolated from the surface of teeth. It synthesizes a variety of extracellular polysaccharides, including water-soluble and non-water-soluble glucan and fructan from sucrose. These polysaccharides promote bacterial colonization and are key virulence factors in the formation dental caries. Due to their adhesive capacity, acid production, acid tolerance, and

water-soluble glucan production, S. mutans has long been regarded as one of the main oral pathogens. It also involved in other secondary infections such as bacteremia and endocarditis. Colonies of S. mutans grown on blood agar after 48 h anaerobic incubation are either regular and smooth or irregular, hard, and sticky. The diameter of colonies is 0.5–1.0 mm. Zones of αor γ-hemolysis can be observed around colonies of most strains, while β-hemolytic zones can also be observed with the colonies of a few strains. On

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Fig. 3.77 S. mutans in short chains (SEM)

Fig. 3.78 S. mutans in long chains (SEM)

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Fig. 3.79 Self-curing cells of S. mutans (SEM) S. mutans are spherical (0.5–0.75 μm in diameter), Gram-positive cocci in pairs or chains. Long chains form in broth and short rod-shaped cells (0.5–1.0 μm in length) can be detected in acidic broth and on some solid media. The phenomenon of selfcuring bacterial cells can be detected under SEM

Fig. 3.80 Colonies of S. mutans incubated on blood BHI agar

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Fig. 3.81 Colonies of S. mutans incubated on MS agar

Fig. 3.82 Smooth and rough colonies of S. mutans incubated on MS agar (stereomicroscope)

agar containing sucrose, most strains form stacked colonies (about 1 mm in diameter) with drop-like or myxoid glucan products above or

surrounding the colonies. MS is the commonly used selective medium, and both smooth and rough colonies can be observed on the same plate.

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Fig. 3.83 Bacterial culture of S. mutans incubated in TPY broth

3.1.6.5 Streptococcus sobrinus S. sobrinus was originally classified as serotypes d and g of S. mutans, and it is named based on its close relationship with S. mutans. Research has shown that S. sobrinus has the second highest rate of cariogenicity after S. mutans [11]. The G + C content of the S. sobrinus genome is 44%–46% (by Tm method), and its type strain is ATCC33478 (SL). The cells are Gram-positive cocci (Figs. 3.84, 3.85, and 3.86). Cultures of S. sobrinus grow under similar conditions as those used to culture S. mutans and their colony characteristics are shown in Figs. 3.87, 3.88, and 3.89. Biochemical Reactions S. sobrinus is able to ferment mannitol, inulin, and lactose to produce acid, but its ability to ferment sorbitol, D-melibiose, and raffinose varies by strain. Moreover, most strains of S. sobrinus can produce H2O2, but cannot metabolize arginine to produce ammonia. Most strains also cannot hydrolyze

aesculin and do not synthesize obvious amounts of extracellular polysaccharide. Colonization Characteristics The main site of colonization of S. sobrinus is on the surface of human teeth. S. sobrinus can form stacked and rough colonies (about 1 mm in diameter) on sucrose agar plates. Liquid-like glucan products can be observed above or surrounding these colonies. On TPY agar plates or BHI blood agar plates, S. sobrinus can form smooth sticky colonies, and α-hemolysis can also be detected for some strains growing on blood agar plates.

3.1.6.6 Streptococcus oralis The cells of S. oralis are Gram-positive (Fig. 3.90), spherical, and arranged in short chains. In addition, S. oralis cells are non-motile, have no capsule, and do not form spores. The genomic G + C content of S. oralis

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Fig. 3.84 S. sobrinus cells (Gram stain)

Fig. 3.85 S. sobrinus cells (SEM) The cells of S. sobrinus are Gram-positive, spherical (about 0.5 μm in diameter), are arranged in pairs or in chains, and often form long chains

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130 Fig. 3.86 S. sobrinus cells (SEM)

Fig. 3.87 Colonies of S. sobrinus (TPY agar plate)

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Fig. 3.88 Colonies of S. sobrinus (BHI blood agar plate)

Fig. 3.89 Alphahemolytic colonies of S. sobrinus (BHI blood agar plate, stereomicroscope)

is 40% (Tm method) and its type strain is NCTC11427 (LUG1, PB182). S. oralis is a facultative anaerobe and most commonly grown on TPY agar (Fig. 3.91). Cells

of this species can grow in medium containing 0.0004% crystal violet, and α-hemolytic reaction can be detected when colonies are grown on

132 Fig. 3.90 Cells of S. oralis (Gram-positive coccus)

Fig. 3.91 Colonies of S. oralis (TPY agar plate)

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Fig. 3.92 Colonies of S. oralis (BHI agar plate)

Fig. 3.93 Colonies of S. oralis (stereomicroscope)

blood agar plates (Fig. 3.92). Characteristic S. oralis colonies are shown in Fig. 3.93. S. oralis can reduce tetrathionate, but does not produce catalases. In fact, S. oralis is relatively biochemically inactive. This species is mainly

isolated from the human oral cavity and is a common member of the oral microflora. Streptococci are clinically divided into three major categories: α-hemolytic, β-hemolytic, and γ-hemolytic.

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Fig. 3.94 Cells of α-hemolytic streptococcus (Gram stain)

3.1.6.7 a-Hemolytic Streptococcus Streptococci that fall in to the α-hemolytic group include all Streptococcus species that form a grass-green hemolytic zone around their colonies when grown on blood agar plates. This category included species such as S. sanguis, S. mitis, and S. vestibularis. Like other streptococci, α-hemolytic cells are Gram-positive (Fig. 3.94), spherical, and non-motile. The great majority of cells are organized in pairs or short chains. Cells observed by SEM are shown in Fig. 3.95. Alpha-hemolytic streptococci are facultative anaerobes and form a characteristic α-hemolytic zone on blood agar plates. The α-hemolytic zone appears as a narrow grass-green zone that is observed around colonies (Fig. 3.96). Members of this group are also known as grass-green streptococci. Colonies observed by stereomicroscope are shown in Fig. 3.97.

streptococci and have the highest pathogenicity. These are the causative agents of various oral infectious diseases, including phlegmon in the maxillofacial region, acute tonsillitis, and periodontal abscess. Like other Streptococcus species, cells of β-hemolytic streptococci are Gram-positive (Fig. 3.98), rounded, and non-motile, and the great majority organize themselves into short chains. However, most cells in liquid culture form long chains. Cells observed by SEM are shown in Fig. 3.99. Beta-hemolytic streptococci are facultative anaerobes that form a broad and completely transparent hemolytic zone around colonies grown on blood agar plates (Fig. 3.100). Colonies observed by stereomicroscope are shown in Fig. 3.101.

3.1.6.8 b-Hemolytic Streptococcus Streptococcal species that fall into the β-hemolytic category include all species that can form a β-hemolytic zone, including S. pyogenes and S. agalactiae. Beta-hemolytic streptococci are also known as pyogenic hemolytic

3.2.1

3.2

Gram-Negative Bacteria Leptotrichia

Leptotrichia is a Gram-negative anaerobic bacillus and is a very commonly observed genus in the human oral cavity. The genus Leptotrichia was first found in 1896 and was named Leptothrix as it was isolated from

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Fig. 3.95 Cells of α-hemolytic streptococcus (SEM)

Fig. 3.96 Colonies of α-hemolytic streptococcus (BHI blood agar plate)

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Fig. 3.97 Colonies of α-hemolytic streptococcus (stereomicroscope)

Fig. 3.98 Cells of β-hemolytic streptococcus (Gram stain)

the rabbit uterus. For a long time, Leptotrichia were considered opportunistic pathogens until recent reports that indicated that they may be pathogenic [12]. Leptotrichia can be isolated from the oral cavity and are mainly found in bacterial biofilms. It can also separate from the vagina and the uterus of pregnant women.

The Leptotrichia cell measures 0.8–1.5  5–20 μm. They can be straight or curved rod shapes. The ends of the cell (either one or both ends) can be sharp or rounded. Cells normally organize as pairs or in a chain (Figs. 3.102, 3.103, and 3.104). The cells do not produce spores and are non-motile. Fresh cultures can be stained Gram-positive. Under the light

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Fig. 3.99 Cells of β-hemolytic streptococcus (SEM)

Fig. 3.100 Colonies of β-hemolytic streptococcus (BHI blood agar plate)

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Fig. 3.101 Colonies of β-hemolytic streptococcus (stereomicroscope)

Fig. 3.102 Leptotrichia cells (Gram stain)

microscope, both Gram-negative and Gram-positive cells can be observed on a single slide. After culturing in anaerobic blood agar for 1–2 days, Leptotrichia can form 1–2 mm, raised, and transparent colonies with smooth and filamentous edges (Fig. 3.105). Sometimes polymorphous colonies are also formed.

Leptotrichia grow best under anaerobic conditions. Cultures require 5%–10% CO2. The ideal temperature for culture growth is between 35  C and 37  C, while Leptotrichia cells stop growing temperatures drop below 25  C. The ideal pH for culturing these cells is between

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Fig. 3.103 Leptotrichia cells (SEM)

pH 7.0 and 7.4. Growth is not inhibited by 20% bile. Leptotrichia is biochemically active. They can ferment amygdalin, cellobiose, fructose, glucose, maltose, mannose, melezitose, salicin, sucrose, and trehalose to produce acid [13]. The terminal products of lactose and starch fermentation are variable. Leptotrichia does not ferment arabinose, dulcitol, glycerol, inositol, inulin, mannitol, melibiose, raffinose, rhamnose, ribose, sorbitol, and xylose. The cells do not produce indole, catalase, urease, H2S, phospholipase, and ammonia gas. The percent G + C in Leptotrichia DNA is 25% (by Tm or Bd). The type strain is ATCC14201.

3.2.2

Veillonella

Veillonella are Gram-negative anaerobic cocci and belong to the family Veillonellaceae. Strains

detected in oral cavities include V. parvula, V. atypica, and V. dispar.

3.2.2.1

Veillonella parvula subsp. parvula V. parvula subsp. parvula are Gram-negative anaerobic cocci and are among the most common bacteria in the oral cavity [14]. Cell characteristics are shown in Figs. 3.106 and 3.107. The DNA G + C content is 38% when analyzed by Tm or 41% when analyzed by Bd. The type strain is ATCC10790. V. parvula subsp. parvula is a strict anaerobe. Several strains require putrescine and cadaverine in their growth medium. Characteristic colonies are shown in Figs. 3.108 and 3.109. The cells are relatively biochemically inactive when tested using classical biochemical tests. They are unable to ferment carbohydrate to acid and do not produce indole. They appear negative with the catalase test, but are able to reduce nitrate to nitrite.

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Fig. 3.104 Leptotrichia cells (SEM)

Fig. 3.105 Leptotrichia colonies

V. parvula subsp. parvula is detected in saliva, on the tongue, and in plaques. They are able to utilize lactate produced by Streptococcus mutans

and are thus considered as beneficial bacteria in dental plaques. They form part of the normal human gut flora.

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Fig. 3.106 V. parvula subsp. parvula cells (Gram stain)

Fig. 3.107 V. parvula subsp. parvula cells (SEM) V. parvula subsp. parvula cells are spherical and often arranged in piles or clumps. The cells are Gramnegative, but can show up as Gram-positive in immature cultures

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142 Fig. 3.108 V. parvula subsp. parvula colonies (BHI blood agar)

Fig. 3.109 V. parvula subsp. parvula colonies (stereomicroscope) V. parvula subsp. parvula require strictly anaerobic condition, forming small gray-white colonies on the surface of BHI blood agar

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References 1. Madigan M, Martinko J. Brock biology of microorganisms. 11th ed. Upper Saddle River: Prentice Hall; 2005. 2. Ventura M, Turroni F, Zomer A, Foroni E, Giubellini V, Bottacini F, Canchaya C, Claesson MJ, He E, Mantzourani M, Mulas L, Ferrarini A, Gao B, Delledonne M, Henrissat B, Coutinho P, Oggioni M, Gupta RS, Zhang Z, Beighton D, Fitzgerald GF, O’Toole PW, van Sinderen D. The Bifidobacterium dentium Bd1 genome sequence reflects its genetic adaptation to the human oral cavity. PLoS Genet. 2009;5(12):e1000785. 3. Bottacini E, O’Connell Motherway M, Kuczynski J, O’Connell KJ, Serafini E, Duranti S, Milani C, Turroni E, Lugli GA, Zomer A, Zhurina D, Riedel C, Ventura M, van Sinderen D. Comparative genomics of the Bifidobacterium breve taxon. BMC Genomics. 2014;15:170. 4. Altermann E, Russell WM, Azcarate-Peril MA, Barrangou R, Buck BL, McAuliffe O, Souther N, Dobson A, Duong T, Callanan M, Lick S, Hamrick A, Cano R, Klaenhammer TR. Complete genome sequence of the probiotic lactic acid bacterium Lactobacillus acidophilus NCFM. Proc Natl Acad Sci USA. 2005;102(11):3906–12. 5. Maze A, Boel G, Zuniga M, Bourand A, Loux V, Yebra MJ, Monedero V, Correia K, Jacques N, Beaufils S, Poncet S, Joyet P, Milohanic E, Casaregola S, Auffray Y, Perez-Martinez G, Gibrat JF, Zagorec M, Francke C, Hartke A, Deutscher J. Complete genome sequence of the probiotic Lactobacillus casei strain BL23. J Bacteriol. 2010;192 (10):2647–8. 6. Ricaurte JC, Klein O, Labombardi V, Martinez V, Serpe A, Joy M. Rothia dentocariosa endocarditis complicated by multiple intracranial hemorrhages. South Med J. 2001;94(4):438–40.

143 7. Vos P, Garrity G, Jones D, Krieg NR, Ludwig W, Rainey FA, Schleifer KH, Whitman W. Bergey’s manual of systematic bacteriology, vol. 3. New York: Springer; 2009. 8. Levinson W. Review of medical microbiology and immunology. 11th ed. New York: MCGraw-Hill Medical; 2010. p. 94–9. 9. Xu P, Alves JM, Kitten T, Brown A, Chen Z, Ozaki LS, Manque P, Ge X, Serrano MG, Puiu D, Hendricks S, Wang Y, Chaplin MD, Akan D, Paik S, Peterson DL, Macrina FL, Buck GA. Genome of the opportunistic pathogen Streptococcus sanguinis. J Bacteriol. 2007;189(8):3166–75. 10. Ajdic D, McShan WM, MCLaughlin RE, Savic G, Chang J, Carson MB, Primeaux C, Tian R, Kenton S, Jia H, Lin S, Qian Y, Li S, Zhu H, Najar F, Lai H, White J, Roe BA, Ferretti JJ. Genome sequence of Streptococcus mutans UA159, a cariogenic dental pathogen. Proc Natl Acad Sci U S A. 2002;99 (22):14434–9. 11. Conrads G, de Soet JJ, Song L, Henne K, Sztajer H, Wagner-Dobler I, Zeng AP. Comparing the cariogenic species Streptococcus sobrinus and S. mutans on whole genome level. J Oral Microbiol. 2014;6:26189. 12. Eribe ER, Olsen I. Leptotrichia species in human intections. Anaerobe. 2008;14(3):131–7. 13. Thompson J. Pikis, a metabolism of sugars by genetically diverse species of oral Leptotrichia. Mol Oral Microbiol. 2012;27(1):34–44. 14. Gronow S, Welnitz S, Lapidus A, Nolan M, Ivanova N, Glavina Del Rio T, Copeland A, Chen E, Tice H, Pit- luck S, Cheng JF, Saunders E, Brettin T, Han C, Detter JC, Bruce D, Goodwin L, Land M, Hauser L, Chang YJ, Jeffries CD, Pati A, Mavromatis K, Mikhailova N, Chen A, Palaniappan K, Chain P, Rohde M, Goker M, Bristow J, Eisen JA, Markowitz V, Hugenholtz P, Kyrpides NC, Klenk HP, Lucas S. Complete genome sequence of Veillonella parvula type strain (Te3). Stand Genomic Sci. 2010;2(1):57–65.

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Subgingival Microbes Quan Yuan, Chenchen Zhou, Jing Xie, Demao Zhang, Liwei Zheng, Yuqing Li, Biao Ren, Xian Peng, and Xuedong Zhou

Abstract

This chapter mainly introduces the common microbes in subgingival plaque. Subgingival plaque refers to the plaque located below the gingival margin and distributed in the gingival groove or periodontal pocket. It can be divided into adherent subgingival plaque and nonadherent subgingival plaque. The adherent subgingival plaque extends from

Q. Yuan Department of Dental Implantology, West China Hospital of Stomatology, Sichuan University, Chengdu, China State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China C. Zhou · J. Xie · D. Zhang · Y. Li · B. Ren · X. Peng State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China L. Zheng Department of Pediatric Dentistry, West China Hospital of Stomatology, Sichuan University, Chengdu, China State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China X. Zhou (*) State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

the supragingival plaque to the periodontal pocket and attaches to the root surface of the tooth. Its structure is similar to the supragingival plaque, mainly Gram-positive cocci, bacilli, and filamentous bacteria, and a small number of Gram-negative short bacillus and spirochetes can be seen. Nonadherent subgingival plaque is located on the surface of adherent subgingival plaque, with loose structure, and is mainly Gram-negative anaerobe. The biological and pathogenic characteristics of Gram-positive bacteria, including Enterococcus, Eubacterium, Peptostreptococcus, and Propionibacterium, and Gram-negative bacteria including Bacteroides, Capnocytophaga, Eikenella, Fusobacterium, Helicobacter, Aggregatibacter, Prevotella, Porphyromonas, and Treponema, were introduced in this chapter. Gram staining, plate culture, colony, and scanning electron microscope (SEM) images of each microorganism were also provided. Keywords

Bacteroides · Fusobacterium · Aggregatibacter · Prevotella · Porphyromonas · Treponema

Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China e-mail: [email protected] # Zhejiang University Press 2020 X. Zhou, Y. Li (eds.), Atlas of Oral Microbiology: From Healthy Microflora to Disease, https://doi.org/10.1007/978-981-15-7899-1_4

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Gram-Positive Bacteria Enterococcus

Enterococci are facultative Gram-positive cocci and belong to Lancefield group D. E. faecalis is also called Streptococcus faecalis and is the most common species in the genus Enterococcus. In recent years, it has been closely studied due to its high detection rate in infected root canals. E. faecalis cells are Gram-positive, oval (0.5–1 μm in diameter), and non-motile. Most cells are arranged in pairs or as short chains (Fig. 4.1). Cellular morphology by SEM is shown in Fig. 4.2. The G + C content of its DNA is 33.5% [1]. The type strain is NCTC775 (ATCC19433, NCDO5681). E. faecalis is a facultative anaerobe. Cells of this species can form smooth, non-transparent, white or creamy, spherical colonies on common nutrient agar plates (Fig. 4.3). However, colonies formed on PSE agar plates containing cholate, esculin, and sodium azide (the selective agar medium for Pfizer Enterococci) are brownish black with brown aureole (Fig. 4.4). This can be used as a characteristic to distinguish E. faecalis from other bacteria. Colonies observed by stereomicroscope are shown in Fig. 4.5. Fig. 4.1 Cells of E. faecalis (Gram-positive cocci)

E. faecalis can ferment most carbohydrates. The main acid produced by glucose fermentation is lactic acid. E. faecalis can also hydrolyze arginine to produce ammonia.

4.1.2

Eubacterium

Eubacterium is a genus of Gram-positive non-sporulating strictly anaerobic bacilli. The name of the genus is still disputed. Bergey’s Manual of Systematic Bacteriology volume 2 (1986) points out that the Greek prefix “eu” means good and useful, rather than “true.” Therefore, the author believes that that “Eubacterium” is the more appropriate name. Currently, bacteria species detected in oral cavity that belong to this genus include E. alactolyticum, E. saburreum, E. lentum, E. limosum, E. nodatum, E. brachy, E. timidum, E. saphenus, and E. minutum. Cells can be homogeneous or polymorphously rod-shaped. No spores are produced. Cells are Gram-positive, but Gram staining old cultures and cultures that have produced acid in the culture medium will yield negative results. Eubacteria are strictly anaerobic. Culturing cells can be difficult due to its strict anaerobic demands, and some strains can only grow in

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Fig. 4.2 Cells of E. faecalis (SEM)

Fig. 4.3 Colonies of E. faecalis (common nutrient agar plate)

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Fig. 4.4 Colonies of E. faecalis (PSE agar plate)

Fig. 4.5 Colonies of E. faecalis (stereomicroscope)

pre-reduced medium. The optimum growth temperature is 37
C, while the optimum pH is 7.0. Eubacteria are chemoheterotrophs and produce energy from mixed organic acids produced by carbohydrates or protein metabolism. These

mainly consist of butyric acid, acetic acid, and formic acid. Most eubacteria from the oral cavity are relatively biochemically inactive. In most cases, cells test negative for catalase and do not hydrolyze

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hippurate. Carbohydrate fermentation, indole production, nitrate reduction, esculin hydrolysis, and other biochemical tests can help differentiate the different species in the genus. Eubacteria mainly colonize the saliva and plaque as a member of the normal oral microflora. E. lentum and E. limosum can be detected in the oral cavity. E. nodatum, E. brachy, E. timidum, E. saphenus, and E. minutum are new species isolated from subgingival plaque of patients with periodontitis and are considered as potential periodontal pathogens. The G + C content in Eubacterium DNA is 30–55% (analyzed by Tm), and the percentage in the type species is 47% (by Tm). In 1999, Kageyama et al. called for a change in the classification of E. lentum and proposed for it to be grouped with Eggerthella lenta, the type species of genus Eggerthella [2]. E. lentum is Gram-positive, irregular, non-sporulating, strictly anaerobic bacillus (Figs. 4.6 and 4.7). As a strict anaerobe, most strains can grow between 30 and 45
C, while some strains can grow at 25
C. Arginine can promote bacterial growth. Characteristic colonies are shown in Figs. 4.8 and 4.9. E. lentum does not ferment carbohydrates. The cells do not hydrolyze aesculin, hippurate, and Fig. 4.6 E. lentum cells (Gram stain)

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gelatin. Ammonia can be produced from arginine, and H2O2 can be produced from agar medium containing 1% arginine. Under anaerobic conditions, the bacteria can produce H2S in the beveled bottom of triple sugar iron agar, but cannot produce H2S in SIM culture medium. The G + C is 62% in DNA, and the type strain is JCM 9979 (¼DSM2243 ¼ ATCC25559 ¼ NCTC11813).

4.1.3

Peptostreptococcus

Peptostreptococcus is the most common Grampositive anaerobic coccus in the human oral cavity and in the clinic. P. anaerobius and P. micros are the most commonly encountered species in this genus. The cells of P. anaerobius are Gram-positive (Fig. 4.10), spherical, approximately 0.5–0.6 μm in diameter, and arranged in pairs or chains. Cells in early cultures have been observed to form long chains. Cells observed by SEM are shown in Fig. 4.11. The G + C content of the P. anaerobius genome is 33%–34%, and its type strain is ATCC27337. The optimal temperature for P. anaerobius growth is 37
C, and cells of this species do not

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Fig. 4.8 E. lentum colonies (BHI blood agar)

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Fig. 4.9 E. lentum colonies (stereomicroscope) E. lentum on horse blood agar forms surface colonies that are 0.5–2 mm in diameter, rounded or convex, low, dim and dark or lustrous, translucent or opaque, and smooth, wedge-shaped, or neatedged. A striped appearance can be observed under incident light

Fig. 4.10 Cells of P. anaerobius (Gram stain)

grow well at 25 or 30
C and do not grow at all at 45
C. Growth is stimulated by 0.02% polysorbate-80. P. anaerobius cells form pinpoint-like or rounded (about 1 mm in diameter), raised, white, glossy, non-transparent

colonies with a smooth surface, without hemolytic zone (Fig. 4.12). Colonies formed on the surface of BHI supplemental medium without addition of blood are gray. Broth cultures of P. anaerobius are usually not muddy, and

152 Fig. 4.11 Cells of P. anaerobius (SEM)

Fig. 4.12 Colonies of P. anaerobius (BHI blood agar plate)

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Fig. 4.13 Colonies of P. anaerobius (stereomicroscope)

granulated or viscous precipitates can be observed. Colonies observed by stereomicroscope are shown in Fig. 4.13. P. anaerobius is relatively biochemically inactive, and cells usually do not ferment carbohydrates. The main acidic metabolic end-products in PVG liquid culture of the type strain are acetic acid, isobutyric acid, butyric acid, isovaleric acid, and isocaproic acid. P. anaerobius can produce CO2 and H2 from pyruvates under anaerobic conditions. In deep glucose agar, P. anaerobius can produce a large amount of gas and generate ammonia from peptone. Dental plaque and gingival sulcus are the main habitats for P. anaerobius in the oral cavity. Moreover, P. anaerobius is often detected in clinical samples of periodontitis, pulpitis, and pericoronitis.

4.1.4

Propionibacterium

Propionibacterium is a genus of Gram-positive polymorphic bacilli that do not sporulate. The genus can be divided into two groups: one that lives on the skin, including inside the oral and intestinal tracts, named the sores and blisters group or P. acnes and another that lives in dairy products, cheese, or green fodder named dairy group or typical propionibacteria.

The cells are polymorphic in form, many have two rounded ends, and others are shaped like Corynebacterium diphtheria, with one rounded end and one tapered end. Cells are 0.5–0.8 μm in diameter and 1–5 μm in length and form two branches or branched rods. Cocci are observed in old cultures, arranged as single cells, in pairs, in chains, or in “Y”- or “V”-shaped branched chains. Cells are Gram-positive, but some cells can also stain Gram-negative. Members of this genus are anaerobic or microaerophilic bacteria. The highest rate of growth takes place 48 h after inoculation. The propionibacteria are chemoheterotrophic bacteria that need a complex nutritional medium such as BHI agar in order to be cultured. Most species can grow in dextrose broth containing 20% bile or 6.5% NaCl. P. acnes colonies on the surface of agar can produce colorful pigmentation including white, gray, pink, red, or yellow. The main acid metabolites are propionic acid and acetic acid when cultured in PYG broth. The production of a significant quantity of propionic acid is the identifying feature of this genus when attempting to differentiate them from other Grampositive non-sporulating anaerobic bacteria. They can also produce some isovaleric acid, formic acid, succinic acid, and lactic acid. All the species in this genus can use glucose to produce acid. They test positive for catalase. Species of Propionibacterium are distinguished using

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indole production, nitrate test, aesculin hydrolysis, gelatin liquification, and other biochemical tests such as the fermentation of sucrose, maltose, and mannitol. The G+ C content of Propionibacterium DNA is 53%–57% by Tm method. The type species is P. freudenreichii. P. acnes is a Gram-positive irregular bacillus [3] (Figs. 4.14, 4.15, and 4.16). It is either anaerobic or microaerophilic. A medium with low redox potential is required for primary cultures. Characteristic colonies are shown in Figs. 4.17 and 4.18. Culture in dextrose broth is cloudy or clear with fine granular precipitates. In old broth, the culture exhibits light red precipitation. The final pH value of a culture is 4.5–5.0 in PYG broth. The terminal acid metabolites are propionic acid and acetic acid. P. acnes is part of the normal microflora, but the number of cells varies greatly between different individuals. P. acnes can be separated from the secretions from acne, wounds, blood, pus, and intestinal contents. The G+ C content of the bacterial DNA is about 57%–60% (by Tm method). The type strain is ATCC6919 (NCTC737).

Fig. 4.14 P. acnes cells (Gram stain)

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4.2 4.2.1

Gram-Negative Bacteria Bacteroides

Bacteroides is a genus of Gram-negative, obligate anaerobic, non-sporulating bacilli that belong to the family Bacteroidaceae. Members of this genus are chemoheterotrophs and can use carbohydrates, peptone, and other intermediate bacterial metabolites. Bacteroides are naturally found in the mouth, tongue, intestinal tract, and vagina. All strains that can tolerate bile (20% oxgall salts) fall collectively into the B. fragilis group, including B. fragilis, B. thetaiotaomicron, etc., that are detected in cultures from appendicitis, peritonitis, and cervicitis clinical specimens. Other species that cannot tolerate bile and that produce melanin were reclassified to the genera Porphyromonas and Prevotella. Cells measure 0.8–1.8  0.8–1.6 μm when grown in glucose broth, often showing visible vacuoles or darker stain at the ends. Cells are arranged singly or in pairs with no spores. Many strains are encapsulated.

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Fig. 4.15 P. acnes cells (SEM)

Fig. 4.16 P. acnes cells (SEM) P. acnes cells are polymorphic, but are mainly thin and long rod-shaped cells. Sero variant type II cells are mainly spherical with no spores. Cells stain Grampositive but some can stain Gram-negative

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Fig. 4.17 P. acnes colonies (BHI blood agar)

Fig. 4.18 P. acnes colonies (stereomicroscope) P. acnes can generate pinprick-sized colonies to colonies 0.5 mm in diameter, white or gray, glossy, translucent or opaque, and pad-shaped or neat-edged 2–3 days postinoculation on the surface of horse blood or rabbit blood agar. On rabbit blood agar, 68% of Sero variant type I strains can generate hemolytic reaction, while Sero variant type II strains cannot

4.2.1.1 Bacteroides fragilis B. fragilis are Gram-negative bacilli (Figs. 4.19, 4.20, and 4.21). They are obligate anaerobes and grow well in nutrient agar supplemented with

chlorinated hemoglobin and vitamin K1. Growth is inhibited by 20% bile. Optimum growth takes place at pH 7.0. Characteristic colonies are shown in Figs. 4.22 and 4.23.

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Fig. 4.19 B. fragilis (Gram stain)

Fig. 4.20 B. fragilis (SEM)

The cells mainly produce succinic acid and acetic acid in PYG liquid medium, but lactic acid, propionic acid, isovaleric acid, isobutyric

acid, formic acid, benzene, and acetic acid are also produced. This species is biochemically active and is capable of fermentating glucose,

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Fig. 4.21 B. fragilis (SEM) B. fragilis appears rounded on both ends, with visible vacuoles or darker stain at the ends. The cells are approximately 0.8–1.3  1.6–8.0 μm when grown in glucose broth. The cells are can be single or in pairs; they form no spores and are Gram-negative

Fig. 4.22 B. fragilis colonies (BHI blood agar)

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Fig. 4.23 B. fragilis colonies (stereomicroscope) B. fragilis can form gray colonies about 1–3 mm in diameter. The colonies are rounded, smooth, translucent, or semi-transparent on blood agar

lactose, maltose, fructose, raffinose, and other carbohydrates, hydrolyzing esculin. B. fragilis tests negative with the indole test. The percent G + C in B. fragilis DNA is 41%– 44% (using the Tm method). The type strain is ATCC25285 (NCTC9343).

4.2.1.2 Bacteroides thetaiotaomicron This species is Gram-negative (Figs. 4.24 and 4.25) and its morphology under SEM is shown in Figs. 4.26 and 4.27. B. thetaiotaomicron are obligate anaerobes and have similar growth requirements and characteristics as B. fragilis. Characteristic colonies are shown in Figs. 4.28 and 4.29. Cells exhibit active biochemistry and are able to ferment glucose, lactose, maltose, fructose, raffinose, arabinose, cellobiose, rhamnose, and other carbohydrates. Cells can hydrolyze esculin and test positive for indole. Cells grow well in medium containing 20% bile. The genomic G + C content is 40%–43% (by Tm). The type strain is ATCC29148 (NCTC10582).

4.2.2

Capnocytophaga

Capnocytophaga are Gram-negative, facultative anaerobic bacteria. They were the earliest bacteria to be isolated and named from the human subgingival plaque. Common Capnocytophaga species are C. ochracea, C. sputigena, C. gingivalis, C. granulosa, and C. haemolytica. Capnocytophaga cells are 0.42–0.6  2.5–2.7 μm in size and are shaped like bent rods or filaments, usually with rounded or slightly pointed ends. The length of the cells varies. In liquid culture, cells are polymorphic or take on a long, filamentous morphology, and tight clumps can be observed. The bacteria produce no capsule and no sheath. They do not form spores, have no flagella, but have sliding motility. Capnocytophaga are facultative anaerobes, but do not grow under aerobic conditions. Cultures grow well in a CO2 added anaerobic environment. Primary cultures should be performed in an aerobic environment with CO2 added.

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Fig. 4.24 B. thetaiotaomicron cells (Gram stain)

Fig. 4.25 B. thetaiotaomicron cells (Gram stain)

Species in this genus often form colonies of wet, thin, flat, diffuse growth with ragged edges on TS blood agar and BHI blood agar. After 24 h of incubation at 35–37
C, the size of colonies is like pinpricks. After incubation for 48–96 h,

colonies become 2–4 mm in diameter and take on the appearance of bumps. Some colonies may become recessed into the agar. Aside from hemolytic Capnocytophaga (which produces β-hemolysis), other species are not hemolytic on

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Fig. 4.26 B. thetaiotaomicron cells (SEM)

blood agar. The concentration of agar in the medium affects the force of sliding motility. Capnocytophaga cultures can produce a special smell, similar to caramel or a bitter almond flavor. Colonies on the agar surface can produce white to pink or orange yellow pigmentation. Centrifuged cells appear to be an orange yellow clump. Capnocytophaga does not produce indole; can ferment glucose, lactose, maltose, mannose, and sucrose acid; and does not ferment mannitol and xylose. It can hydrolyze esculin and tests negative for catalase and oxidase, while testing positive for ONPG and benzidine. Nitrate reduction, dextran hydrolysis, starch or gelatin hydrolysis, and other biochemical tests can help identify this genus of bacteria. Member of the normal microflora of humans and primates, this genus is mainly found to colonize the oral cavity. They are common oral bacteria and can be obtained from various parts of the oral cavity, including plaque, gingival sulcus,

saliva and sputum, and throat specimens. These bacteria are often detected in mixed bacterial infections, such as juvenile periodontitis, infected root canal, dry socket after tooth extraction, oral ulcers, and other clinical specimens, and can also be isolated from bacteremia; soft tissue infections; injuries and abscesses at various locations; cerebrospinal fluid; vaginal, cervical, and amniotic fluid; trachea; and eyes. The G+ C content of Capnocytophaga genomic DNA is 33%–41% (by Tm method). The type species is yellowish Capnocytophaga.

4.2.2.1 Capnocytophaga gingivalis This Gram-negative bacterium is shown in Figs. 4.30, 4.31, and 4.32. Characteristic colonies are shown in Figs. 4.33 and 4.34. C. gingivalis does not ferment lactose, galactose, amygdalin, salicin, cellobiose, esculin, and glycogen. It also does not hydrolyze starch, dextran, and gelatin. Only 8% of the strains can reduce nitrate.

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Fig. 4.27 B. thetaiotaomicron cells (SEM) B. thetaiotaomicron appears spherical or rounded on both ends, with visible dark stain at the ends. Cells are

Fig. 4.28 B. thetaiotaomicron colonies (BHI blood agar)

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0.7–1.1  1.3–8.0 μm when grown in glucose broth and are arranged as single cells or in pairs. Cells stain Gramnegative

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Fig. 4.29 B. thetaiotaomicron colonies (stereomicroscope) B. thetaiotaomicron colonies are about 1–3 mm in diameter. They form bumpy, glossy, soft, white, round colonies

Fig. 4.30 Cells of C. gingivalis (Gram stain)

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on blood agar. An orange peel-like appearance can be observed on the colony surface under stereomicroscope

164 Fig. 4.31 Cells of C. gingivalis (SEM)

Fig. 4.32 Cells of C. gingivalis (SEM) Cells of C. gingivalis are fusobacterium-shaped; the ends are usually rounded. Cells are often arranged in an orderly manner and stain negative by Gram stain

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Fig. 4.33 Colonies of C. gingivalis (BHI blood agar)

Fig. 4.34 Colonies of C. gingivalis (stereomicroscope) Colonies of C. gingivalis on BHI blood agar are irregular, gray colonies, with ragged edges. Typical hair-like diffuse colonies can be seen under the stereomicroscope

The G + C content in C. gingivalis DNA is 40% (by method). The type strain is ATCC33624.

4.2.2.2 Capnocytophaga sputigena This species is a Gram-negative bacillus (Figs. 4.35, 4.36, and 4.37). Characteristic colonies are shown in Figs. 4.38 and 4.39.

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Fig. 4.35 Cells of C. sputigena (Gram stain)

Fig. 4.36 Cells of C. sputigena (SEM)

The cells can ferment lactose, glucose, maltose, and sucrose, but do not ferment mannitol, cellobiose, glycogen, and xylose. C. sputigena

does not hydrolyze starch and dextran. The hydrolysis of gelatin and nitrate reduction are the most important features by which this species

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Fig. 4.37 Cells of C. sputigena (SEM) Cells of C. sputigena are bent bacilli, usually with rounded ends. They produce no spores and stain Gram-negative

Fig. 4.38 Colonies of C. sputigena (BHI blood agar)

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Fig. 4.39 Mesh-like structure on the surface of colonies of C. sputigena (stereomicroscope) Colonies of C. sputigena on BHI blood agar surface are flat, spread orange colonies. Typical hair-like diffuse structure can be seen under the stereomicroscope

can be distinguished from other members of the genus. C. sputigena may be involved in juvenile periodontitis. The G + C content of C. sputigena DNA is 33%–38% (Tm method). The type strain is ATCC33612.

4.2.3

Eikenella

Eikenella is a genus of Gram-negative facultative anaerobic bacteria that do not produce spores. E. corrodens was thus named because it produces typical colonies which can erode agar. It is also known as Bacteroides corrodens and is the only species in the genus Eikenella [4]. Cells stain Gram-negative (Figs. 4.40, 4.41, and 4.42). E. corrodens is a facultative anaerobe, and primary cultures require anaerobic conditions or supplementation with 5%–10% CO2. It is essential to add hemin (5–25 mg/L) to the culture when grown under aerobic conditions. The optimum growth temperature is from 35 to 37
C, the optimum pH is 7.3, and cultures require sufficient humidity. Characteristic colonies are shown in Figs. 4.43, 4.44, and 4.45.

E. corrodens does not grow well in liquid media. Broth supplemented with 0.2% agar, cholesterol (10 mg/L), and 3% serum can promote its growth. Under aerobic conditions, 5% to 10% bile can inhibit growth. However under anaerobic conditions, up to 10% bile can be tolerated. E. corrodens is biochemically inactive. It does not ferment glucose and other carbohydrates or produce acid. It tests negative for catalase, urease, arginine dehydrogenase, and indole, but is positive for nitrate reduction, as well as oxidase and lysine decarboxylase. E. corrodens is a member of the normal flora in the human oral cavity and intestinal tract. It can also be isolated from the upper respiratory tract and urogenital tract. As an opportunistic pathogen, it is often associated with other bacterial pathogens to cause mixed bacterial infections, especially in the mouth and respiratory tract. Its detection rate is higher in lesions of active adult periodontitis and specimens of dry socket after tooth extraction; it is suspected to be related to periodontitis. The G + C content in its genomic DNA is 56%–58% (by Tm method). The type strain is ATCC23834 (NCTC10596).

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Fig. 4.40 Cells of E. corrodens (Gramnegative)

Fig. 4.41 Cells of E. corrodens (SEM)

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Fig. 4.42 Cells of E. corrodens (SEM) Cells of E. corrodens are 0.3–0.4  1.5–4.0 μm in size and are rounded at the ends. They are mostly rod-shaped, short rod-shaped, or club-shaped and can sometimes be found in the shape of a short wire rod. The cells do not produce spores and are non-motile. “Tremorshaped movement” can be seen on the surface of the agar. Cells stain Gramnegative

Fig. 4.43 Colonies of E. corrodens (BHI blood agar)

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Fig. 4.44 Colonies of E. corrodens (stereomicroscope)

Fig. 4.45 A pearlescent ring can be observed at the center of colonies of E. corrodens (stereomicroscope) Agar cultures of E. corrodens have a bleach-like smell, similar to Haemophilus and Pasteurella cultures on agar. Colonies do not produce hemolytic reactions on blood agar, but a light green ring can be seen around colonies. Two different types of colonies can form on blood agar: invasive phenotype and non-invasive phenotype. The invasive strain forms on the surface of blood agar when conditions are 36
C, with 15% CO2 and 100% humidity. Colony diameter ranges 0.2–0.5 mm (after 24 h culture) or

from 0.5 to 1.0 mm (after 48 h culture). Colonies are light yellow and opaque, and the center of the colony has a clear pearlescent ring. The edge of the colony is rough and refractive and has a hair-like diffuse edge; “tremor-shaped movement” can be seen on the surface of the agar. The non-invasive phenotype forms colonies with a diameter of 0.5–1 mm. The colonies are hemispherical and translucent, with no hair-like diffuse edge. They do not invade agar, show no adhesion to the agar, and have no “tremor-shaped movement”

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Fusobacterium

Fusobacterium is a group of Gram-negative obligate anaerobic bacteria that do not form spores. They belong to the family Bacteroidaceae. Species mainly found in the oral cavity are F. nucleatum, F. necrophorum, and F. varium. Most fusobacteria are spindle-shaped cells. They may also appear polymorphous. Polymorphic fusobacteria can form globular or long filiform cells. F. necrophorum can take on many other morphologies, including irregular spherical swollen cells and linear cells. The cells are non-motile, do not form spores, and stain Gramnegative. These obligate anaerobes can be grown in aerobic conditions on agar plates when 5%–10% CO2 is added to the culture conditions. Their sensitivity to oxygen depends on the specific bacterial species, the quantity of cells inoculated, and the type of culture medium. Fusobacterium can be detected in clinical specimens of pus or gangrene infections. F. nucleatum has a high prevalence in saliva and dental plaque and is considered to be one of the bacteria involved in mixed infections of periodontitis, root canal infection, and postextraction infection. Fig. 4.46 F. nucleatum cells (Gram stain)

The genomic G + C content of this genus is 26%–34% (Tm). The type species is F. nucleatum.

4.2.4.1 Fusobacterium nucleatum The species F. nucleatum contains five subspecies: F. nucleatum animal subspecies (F. nucleatum subsp. animalis), fusiform subspecies (F. nucleatum subsp. fusiforme), nuclear subspecies (F. nucleatum subsp. nucleatum), polymorphic subspecies (F. nucleatum subsp. polymorphum), and Vincent subspecies (F. nucleatum subsp. vincentii). The cells are Gram-negative spindle-shaped coli. Cell morphologies are shown in Figs. 4.46, 4.47, and 4.48. F. nucleatum is an obligate anaerobe, but can grow under atmospheric conditions with >6% oxygen by volume. The cells are viable even after 100 minutes of exposure to the air. Characteristic colonies are shown in Figs. 4.49, 4.50, and 4.51. F. nucleatum is not biochemically active. They do not transform lactate into propionate. They can produce indoles and DNase, but do not produce phosphatase. Most of the strains produce H2S and can agglutinate red blood cells from both humans and animals.

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Fig. 4.47 F. nucleatum cells (SEM)

Fig. 4.48 F. nucleatum cells (SEM) F. nucleatum measure 0.4–0.7  3–10 μm in diameter in glucose broth culture, with a fusiform or tapered end. Swelling can often be observed at the center of the cell, and Gram-positive particles can be observed inside the cell. Cell length is generally associated with growth conditions. They have no pili and no flagella. Cells stain Gram-negative

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Fig. 4.49 F. nucleatum colonies (BHI blood agar)

Fig. 4.50 F. nucleatum “breadcrumb” colonies (stereomicroscope) F. nucleatum can form colonies 1–2 mm in diameter. Colonies are rounded or slightly irregular, bumpy, pad-shaped, translucent, and with odor on blood agar plates. Colonies commonly appear to have spots of light that shine through. These are referred to as “breadcrumbs”

colonies. F. nucleatum generally do not produce hemolytic reactions on horse and rabbit blood agar. There is visible flocculent or particle precipitation in glucose broth cultures, but the broth does not necessarily become turbid. The culture has a foul smell, and the final pH value is of a glucose broth culture is pH 5.6–6.2

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Fig. 4.51 F. nucleatum isolated from saliva samples (stereomicroscope)

F. nucleatum are mainly found on transparent gingiva and the gingival groove and make up the oral normal flora. They can also be isolated from upper respiratory tract and chest infections and occasionally from wounds and other sites of infection. F. nucleatum has a high rate of detection in destructive periodontal disease and infectious dental pulp, as it assists other pathogens in establishing oral infectious diseases. The genomic G + C content is 27%–28%. The type strain is ATCC25586 [5].

4.2.4.2 Fusobacterium necrophorum There are two subspecies of F. necrophorum: F. necrophorum fundamental form subspecies (F. necrophorum subsp. funduliforme) [6] and F. necrophorum necrosis subspecies (F. necrophorum subsp. necrophorum). F. necrophorum is a Gram-negative bacillus, with diverse cellular morphologies (Figs. 4.52, 4.53, 4.54, 4.55, and 4.56). Heptose and KDO are among the cell wall lipopolysaccharides. F. necrophorum is an obligate anaerobe. Culture requirements are similar to related species. Characteristic colonies are shown in Figs. 4.57, 4.58, 4.59, 4.60, and 4.61. When cultured in glucose broth medium, F. necrophorum appears muddy, and smooth, flocculent particles or filamentous precipitation

can also be found. The final pH value of a culture in glucose or fructose broth is 5.6–6.3. A few strains have a final pH value of 5.8–5.9 in maltose cultures. F. necrophorum can agglutinate red blood cells from human, rabbit, and guinea pig blood, but not blood from cattle. It cannot hydrolyze glucan. Neither phosphatase, superoxide dismutase, nor lysine decarboxylase is detected, but it can produce DNase. F. necrophorum is mainly isolated from some clinical disease specimens from the human and animal body, including abscess, blood, and necrotic lesions, and especially in liver abscess. The bacteria can be also detected in the mouth. The genomic G + C content is 31%–34%. The type strain is ATCC25286.

4.2.4.3 Fusobacterium varium This Gram-negative species’ cell morphology is shown in Figs.4.62, 4.63, and 4.64. Heptose and KDO make up its cell wall lipopolysaccharides. F. varium is an obligate anaerobe. Its culture conditions are similar to other related species. Characteristic colonies are shown in Figs. 4.65 and 4.66. F. varium cannot hydrolyze glucan and does not produce phosphatase, but it does produce lysine dehydrogenase and DNase.

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Fig. 4.52 Filament of F. necrophorum (Gram stain)

Fig. 4.53 Filament of F. necrophorum with particle content (Gram stain)

F. varium can be isolated from the human mouth and human feces; it can also be detected in suppurative infectious wounds, the upper respiratory tract, and peritonitis; but its pathogenicity is not yet well-defined. The genomic G + C content is 26%–28%. The type strain is ATCC8501 (NCTC10560).

4.2.5

Helicobacter

Helicobacter is a genus of Gram-negative, curved rod-shaped bacteria with polar flagella. Because of the differences in their ultrastructure, fatty acid composition, morphology, growth characteristics, enzyme activity, and 16 s rRNA sequence when

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Fig. 4.54 F. necrophorum bacillus (Gram stain)

Fig. 4.55 F. necrophorum cells (SEM)

compared to the genus Campylobacter, the species in this genus were classified as Helicobacter. The type species is H. pylori.

H. pylori was first isolated and cultivated in vitro from the gastric mucosal tissue of patients with chronic gastritis in 1983 by Marshall and

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Fig. 4.56 F. necrophorum cells (SEM) F. necrophorum is morphologically diverse. The cell length can vary significantly and can be spheroids, rod-shaped to filaments more than 100 μm long. Although cells are approximately 0.5–0.7 μm in diameter, when cultured in glucose broth, they may swell to more than 1.8 μm in length, with round or pointed ends. Filamentous material containing particles are commonly seen in broth cultures, while coliform cells are found in old cultured cells or on agar. Cells stain Gramnegative

Warren [7]. The organism is associated with gastritis, duodenal and gastric ulcers, and gastric cancer, as well as a number of diseases at distant sites. The World Health Organization/International Agency for Research on Cancer (WHO/IARC) listed H. pylori as a Class A carcinogen in 1994. H. pylori are non-sporulating, slender, curved rods measuring approximately 2.5–4.0 μm  0.5–1.0 μm. They are pleomorphic and can typically be found as spiral, S-shaped, or gull-shaped cells (Figs. 4.67, 4.68, and 4.69). Cells stain Gram-negative with one or more flagella. Sometimes rods or coccoid forms can be found in addition to the typical cellular morphologies when grown on solid culture medium. Under electron microscope, the bacteria have 2–6 flagella that are approximately 30 nm in thickness and 1–1.5 times longer than the bacterial cell. Flagella play a role in cellular movement and anchor cells during adhesion.

H. pylori grow under microaerophilic conditions and have high nutritional requirements. They grow well on Columbia blood agar medium containing 5% defibrinated blood or on brain heart infusion blood agar at 37
C, 95% humidity, and under 10% carbon dioxide, 5% oxygen, and 85% nitrogen. Because of the long growth period, primary cultures require 3–7 days, and subcultures need 2–4 days to grow. Colonies are pinprick-sized, circular, neat, raised, colorless, and translucent and measure approximately 0.5–1 mm in diameter (Figs. 4.70 and 4.71). Resistance of H. pylori to various stressors is weak, as they survive for less than 3 h in air and no more than 1 day at 4
C. Cultures are sensitive to heat, and the only way to preserve cells for the long-term is by cryopreservation at –80
C. H. pylori is not biochemically active. It usually does not produce acid from sugar. Although there have been a few reports of acid production, it only

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Fig. 4.57 F. necrophorum colonies producing α-hemolysis (BHI blood agar)

Fig. 4.58 F. necrophorum colonies producing β-hemolysis (BHI blood agar)

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180 Fig. 4.59 F. necrophorum colonies (BHI sheep-blood agar)

Fig. 4.60 F. necrophorum zigzag colonies with mosaic internal structure (BHI sheep-blood agar, stereomicroscope)

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Fig. 4.61 F. necrophorum colonies with thread structure (stereomicroscope) F. necrophorum forms colonies that are 1–2 mm in diameter, rounded, raised above the surface, cream-colored, and translucent to opaque, with a fan-like or serrated edge on blood agar. A mosaic internal structure can be seen in the

Fig. 4.62 F. varium cells (Gram stain)

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light. Most strains can produce α- or β-hemolysis on rabbit blood agar. In general, β-hemolytic strains are positive for lipase (on egg yolk agar), while α-hemolytic and non-hemolytic strains are negative for lipase and do not produce lecithin enzyme

182 Fig. 4.63 F. varium cells (SEM)

Fig. 4.64 F. varium cells (SEM) F. varium cells measure 0.3–0.7  0.7–2.0 μm. They are polymorphic. Both cocci and bacilli can be observed and cells are present as single cells or in pairs. Cells stain Gramnegative, but also show uneven staining

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Fig. 4.65 F. varium colonies (BHI blood agar)

Fig. 4.66 F. varium colonies (stereomicroscope) F. varium forms pinpricksized colonies to colonies 1 mm in diameter. They appear rounded, low, flat, and convex, with a gray center and a colorless, translucent, and neat edge on blood agar

becomes apparent in media containing a low concentration of peptone. Sugar is not usually the sole carbon source. H. pylori cellular vitality significantly weakens at pH 3.5 and below, while physiological

concentrations of bile salts can inhibit culture growth. H. pylori produces a large quantity of highly active extracellular urease, producing more than 400 times the urease activity than Proteus. H. pylori can also produce oxidase, catalase,

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Fig. 4.67 H. pylori cells (Gram stain) Gram-negative cells are shaped like curved, S-shaped, or gullshaped rods

Fig. 4.68 H. pylori cells (SEM)

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Fig. 4.69 H. pylori cells (SEM)

Fig. 4.70 H. pylori colonies (BHI blood agar) are tiny pinprick colonies (circular, neat, raised, colorless, translucent), and about 0.5–1 mm in diameter and almost do not produce hemolytic reaction

alkaline phosphatase, γ-GGTP, etc. Six positive enzymatic reactions are used as the basis for H. pylori biochemical identification. These are oxidase, catalase, urease, alkaline phosphatase, γ-GGTP, and leucine peptidase.

The infection rate with H. pylori is over 50% worldwide. The infection does not resolve itself and is sensitive to antibiotic treatment, but the recurrence rate is high. The oral cavity is considered to be a secondary site of colonization of

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Fig. 4.71 H. pylori colonies (stereomicroscope)

H. pylori and may be associated with its high recurrence rate. H. pylori has been found in supragingival plaque or subgingival plaque and saliva, the decayed teeth, infected root canals, and mucous membranes of the tongue and buccal and palatal mucosa by a large number of researchers using various molecular technologies. Isolating H. pylori using the traditional culturing method is particularly difficult, as there are many different kinds of bacteria in the oral cavity that form the dental plaque biofilm. The complex mutually beneficial relationship of coexistence and competition that is present within the biofilm makes it relatively stable environment resistant to external stimuli. As a result, oral H. pylori can more easily evade drug treatments. Oral H. pylori infections may be related to oral infectious diseases such as periodontal disease, tooth decay, oral ulcer, etc. Studies confirmed that periodontal disease is more likely to arise in H. pylori-positive patients and the rate of infection in H. pylori-positive patients is positively correlated with the degree of inflammation. Triple therapy combining basic periodontal treatment (supragingival and subgingival scaling) can enhance the eradication of H. pylori and reduce the recurrence rate of H. pylori. The genome size of H. pylori is about 1.67  106 bp and the G + C content is 37% [8]. The type strain is ATCC43504.

4.2.6

Aggregatibacter

Formerly classified as Actinobacillus actinomycetemcomitans, this species was subsequently named Haemophilus actinomycetemcomitans, as is now known as Aggregatibacter actinomycetemcomitans. It is a major pathogen in juvenile periodontitis [9]. These are Gram-negative bacteria and the morphology of bacterial cells is shown in Figs. 4.72, 4.73, an130d 4.74. A. actinomycetemcomitans is a facultative anaerobe and grows well in the microaerophilic environment of 5%–10% CO2. Its optimum growth temperature is 37
C and it does not grow at 22
C. Characteristic colonies are shown in Figs. 4.75, 4.76, and 4.77. Cells ferment fructose, glucose, maltose, and mannose and product acid, but do not ferment sucrose, trehalose, lactose, raffinose, melibiose, and arabinose. A. actinomycetemcomitans tests positive for oxidase and catalase, can reduce nitrate, does not hydrolyze esculin and hippurate sodium, and does not produce H2S and indole. The main colonization site of this species is subgingival plaque. It is detected both in normal oral bacteria and in lesions of juvenile periodontitis patients at a higher detection rate. Therefore, A. actinomycetemcomitans is considered an important pathogen.

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Fig. 4.72 Cells of A. actinomycetemcomitans (Gram stain)

Fig. 4.73 Cells of A. actinomycetemcomitans (SEM)

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Fig. 4.74 Cells of A. actinomycetemcomitans (SEM) A. actinomycetemcomitans are 0.5–0.8  0.6–1.4 μm in size. Cells are spherical, club-shaped, or rod-shaped. Rod-shaped cells are common in agar cultures. The cells

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arrange as single cells, in pairs, or in piles. They produce no spores, are non-motile, and do not form capsule. Cells stain Gram-negative

Fig. 4.75 A. actinomycetemcomitans colonies (BHI blood agar)

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Fig. 4.76 A. actinomycetemcomitans colonies (stereomicroscope)

Fig. 4.77 A. actinomycetemcomitans colonies (stereomicroscope) On the surface of the agar, A. actinomycetemcomitans forms small colonies, with a diameter of approximately 0.5–1.0 mm. Primary cultures are often difficult to lift off

the agar surface. Typical colonies are star-shaped or shaped like crossed cigars, with irregular edges. In broth culture, the growth shows small particle-like opacity and often sticks to the flask wall. However, some strains grow into a homogeneously turbid culture after repeated cultures

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The genomic G + C content is 43% (by Tm method). The type strain is NCTC9710.

4.2.7

Prevotella

Prevotella is a genus named after the French microbiologist A. R. Prevol. These bacteria belong to the genus Bacteroides and include bile-sensitive strains and melanin-producing, sugar-metabolizing strains. The main species of Prevotella found in the oral cavity are P. intermedia, P. melaninogenica, P. loescheii, P. nigrescens, P. denticola, and P. corporis. The most frequently found cell morphology is short bacillus, but long bacilli are sometimes observed. The cells are non-sporulating, are non-motile, and stain Gram-negative. Bacteria belonging to this genus are obligate anaerobes and most cultures require supplementing with hemin and vitamin K. On blood agar, Prevotella spp. produce melanin to form black colonies. In PYG liquid medium, these bacteria can produce acetic acid and succinic acid as the major terminal acid products.

Fig. 4.78 P. intermedia cells (Gram stain)

Species of this genus ferment glucose and hydrolyze gelatin. They are sensitive to bile salt and can thus be distinguished from other bacteroides that can tolerate bile salts. Prevotella are dominant bacteria in the human gingival groove and are also the suspected pathogens behind periodontitis.

4.2.7.1 Prevotella intermedia P. intermedia is a Gram-negative, blackpigmented, anaerobic bacterium [10]. Typical cell morphologies are shown in Figs. 4.78, 4.79, and 4.80. As an obligate anaerobe, cultures require hemin and vitamin K. Most species grow well in temperatures between 25 and 45
C. For other cultural demands, refer to the genus Prevotella. Characteristic colonies are shown in Figs. 4.81 and 4.82. Culture in glucose broth is cloudy, with even precipitation and sticky or slightly sticky deposits at times. The final pH value of in glucose broth ranges from pH 4.9 to 5.4. P. intermedia can produce indole and hydrolyze gelatin, but not esculin. Cells ferment glucose and sucrose, but not arabinose, larch sugar, cellobiose, rhamnose, galactose, and salicin. Cells

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Fig. 4.79 P. intermedia cells (SEM) Most P. intermedia cells form short rods and measure 0.4–0.7  1.5–2 μm, while some cells can measure up to 12 μm in diameter. Cells stain Gram-negative

Fig. 4.80 P. intermedia cells (SEM)

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Fig. 4.81 P. intermedia colonies (BHI blood agar)

Fig. 4.82 P. intermedia colonies (stereomicroscope) Most strains of P. intermedia form colonies 0.5–2.0 mm in diameter. Colonies are round, low, convex, and translucent, with a smooth surface. Colonies show hemolytic reaction on the surface of blood agar, while aging or large colonies may appear opaque. After incubation for

48 h under anaerobic conditions, colonies may appear gray, brown, or black. Colonies can produce a hemolytic ring on rabbit blood agar. One-third of P. intermedia strains can produce dark brown to black colonies within 2 days. Colonies fluoresce brick red when exposed to longwavelength UV light

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can produce superoxide dismutase, but do not hydrolyze dextran. The site of colonization is the gingival sulcus, but cells also can be found in saliva, dental calculus, and other plaque specimens. Previous research has shown that this species is the main pathogenic bacteria in pregnancy-related gingivitis. In addition, the bacteria can also be isolated from pericoronitis, the focus of infection after tooth extraction, infected root canals, infections of the head and neck, and pleural infection. P. intermedia is occasionally isolated from blood, abdominal, or pelvic specimens. The genomic G + C content is 41%–44%. The type strain is ATCC25611.

4.2.7.2 Prevotella nigrescens Originally, this species was classified as a strain of P. intermedia. However, since it does not produce lipase, it can be thus distinguished from P. intermedia. P. nigrescens is a Gram-negative, non-sporulating, melanin-producing, obligate anaerobe. Cell morphologies are shown in Figs. 4.83 and 4.84. For culture requirements, refer to the information on P. intermedia. Characteristic colonies are shown in Figs. 4.85 and 4.86. Fig. 4.83 P. nigrescens cells (Gram stain)

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Most strains ferment glucose, maltose, sucrose, and maltodextrin to produce acid. They also produce indole, hydrolyze starch, and hydrolyze gelatin. In liquid medium containing glucose, these bacteria can produce acetic acid, succinic acid, isobutyric acid, and isovaleric acid. The genomic G + C content is 40%–44%. The type strain is ATCC33563 (namely NCTC9336, VPI8944).

4.2.7.3 Prevotella melaninogenica P. melaninogenica is a Gram-negative, non-sporulating, melanin-producing obligate anaerobic bacterium [11]. Bacterial cells are shown in Figs. 4.87 and 4.88. Most strains require hemin (1 mg/L) and vitamin K (0.1 mg/L) to grow and can grow at pH 8.5 and 25
C. Characteristic colonies are shown in Figs. 4.89, 4.90, and 4.91. Usually, glucose broth culture turns turbid and is accompanied by smooth or filamentous precipitation. The final pH value range is from pH 4.6 to 5.0. Clinical specimens are isolated from the gingival sulcus. The genomic DNA G+ C content is 36%– 40%. The type strain is ATCC25845.

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Fig. 4.84 P. nigrescens cells (SEM) Most cells of P. nigrescens is a coccobacillus. In broth culture the cells are 0.3–0.4  1–2 μm in size. Some cells can measure up to 6–10 μm. Cells stain Gram-negative

Fig. 4.85 P. nigrescens colonies (BHI blood agar)

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Fig. 4.86 P. nigrescens colonies (stereomicroscope) P. nigrescens colonies on horse blood agar after 72 h. The colony diameter is 0.5–2 mm and appears circular, with a neat edge and low convex, and smooth and produces

Fig. 4.87 P. melaninogenica cells (Gram stain)

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brown or black pigment. The edge of the colony is usually black; the center is cream-colored to dark brown. Most strains produce weak hemolytic reaction. Few strains produce a ring indicative of α-hemolysis

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Fig. 4.88 P. melaninogenica cells (SEM) P. melaninogenica cells are coccobacilli and measure 0.5–0.8  0.9–2.5 μm. Occasionally, cells longer than 10 μm are observed. Cells stain Gram-negative

Fig. 4.89 P. melaninogenica colonies (BHI blood agar)

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Fig. 4.90 P. melaninogenica colonies (stereomicroscope)

Fig. 4.91 P. melaninogenica colonies (stereomicroscope) P. melaninogenica colonies on blood agar are 0.5–2.0 mm in diameter. Colonies appear round, convex, and glossy,

with a neat edge. The centers of colonies are usually black and the edges are gray to light brown. After 5–14 d of culture, colonies become completely black. Few strains produce β-hemolytic reaction on rabbit blood agar

4.2.7.4 Prevotella corporis P. corporis is a Gram-negative non-sporulating, melanin-producing obligate anaerobe. Examples of cells are shown in Figs. 4.92 and 4.93.

Growth of P. corporis requires chlorinated hemoglobin and vitamin K1. A 10% final concentration of serum can promote growth and improve fermentation in some strains. Characteristic

198 Fig. 4.92 P. corporis cells (Gram stain)

Fig. 4.93 P. corporis cells (SEM) P. corporis cells grown in glucose broth culture are measured 0.9–1.6  1.6–4.0 μm in size. Cells arrange singly, in pairs, or in short chains. Coccoid cells are also commonly seen, and long filamentous cells can sometimes be observed. Cells stain Gram-negative

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Fig. 4.94 P. corporis colonies (BHI blood agar)

Fig. 4.95 P. corporis colonies (stereomicroscope) P. corporis cultured under anaerobic conditions on blood agar form colonies ranging from pinprick-sized to 1.0 mm. Colonies are rounded bumps with neat edges. After 48 h to 72 h of incubation, colonies can turn light yellow with brownish edges, and colonies cultured for 4 d–7 d turn dark brown

colonies are shown in Figs. 4.94 and 4.95. Glucose broth cultures turn cloudy and often have smooth or coarse precipitates adhered to the bottom flask. The final pH value of glucose broth cultures is 4.8–5.1.

P. corporis can be detected in specimens of all types of clinical infection, including the oral cavity. The genomic G + C content is 43%–46%. The type strain is ATCC33457.

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Fig. 4.96 P. loescheii cells (Gram stain)

Fig. 4.97 P. loescheii cells (SEM)

4.2.7.5 Prevotella loescheii P. loescheii is a species of Gram-negative, non-sporulating, melanin-producing, obligate

anaerobic bacteria. Typical cells are shown in Figs. 4.96, 4.97, and 4.98.

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Fig. 4.98 P. loescheii cells (SEM) Cells of P. loescheii grown in glucose broth culture are 0.4–0.6  0.8–15 μm in size. They are mostly coccobacilli, but can also be club-shaped and long rods. Cells organize into single cells, in pairs, or in a chainlike arrangement. Cells stain Gram-negative

Cultures of P. loescheii require chlorinated hemoglobin, while the addition of 10% serum enhances fermentation. Colonies observed by stereomicroscope are shown in Figs. 4.99 and 4.100. Glucose broth cultures turn cloudy and are accompanied by a smooth deposit. The final pH value of cultures grown in glucose broth is between pH 4.9 and 5.4. P. loescheii does not produce H2S in SIM medium. But the hydrolysis of esculin and the ability to ferment cellobiose help distinguish this species from P. melaninogenica and P. denticola. The genomic G + C content is 46% (type strain). The type strain is ATCC15930 (NCTC11321) [12].

4.2.8

Porphyromonas

In 1998, three species of Bacteroides were found to have significantly different biological characteristics than other Bacteroides species.

However, they were similar in their ability to ferment carbohydrates to produce melanin. These species, namely, B. asaccharolyticus, B. gingivalis, and B. endodontalis, have been placed into a new genus: Porphyromonas. Members of this genus most commonly found in the oral cavity are P. gingivalis and P. endodontalis. Broth-cultured cells are typically small rods approximately 0.5–0.8  1.0–3.5 μm in size; occasionally cells can be found that measure 4–6 μm long. These bacteria produce no spores, are non-motile, and are Gram-negative. Porphyromonas are obligate anaerobes with an optimum growth temperature of 37
C. On blood agar plates, they can form colonies 1–3 mm in diameter that are protuberant, lustrous, and with smooth surface (very few with rough surface) . The primary endpoint product in PYG medium is butyric acid and acetic acid. There is a small

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Fig. 4.99 P. loescheii colonies (BHI blood agar)

Fig. 4.100 P. loescheii colonies (stereomicroscope) P. loescheii cultures grown on blood agar under anaerobic condition form colonies 1.0 to 2.0 mm in diameter. Colonies are rounded, low convex, lustrous, and translucent, with a smooth surface and neat edges. On agar plates containing whole blood, colonies turn white or yellow after 48 h. When anaerobic culture is continued for 14 d, colonies turn light brown. Some strains do not produce obvious dark brown or black colonies

amount of propionic acid, isobutyric acid, and isoamyl propionate produced. The genomic G + C content of this genus is 46%–54%. The type species is P. asaccharolytica.

4.2.8.1 Porphyromonas gingivalis P. gingivalis are Gram-negative, obligate anaerobic bacteria that do not form spores and produce melanin, as shown in Figs. 4.101 and 4.102. Its cell wall peptidoglycan contains lysine, while the

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Fig. 4.101 P. gingivalis cells (Gram stain)

Fig. 4.102 P. gingivalis cells (SEM) P. gingivalis cells are 0.5  1–2 μm rods or coccobacilli; cells on solid medium form coccobacilli or very short rods. Cells stain Gram-negative

main respiratory quinone has nine isoprene units of unsaturated methyl naphthalene quinones. As an obligate anaerobe, chlorinated hemoglobin and vitamin K1 are required in the growth

media. Hemoglobin is the main product of porphyrin. Characteristic colonies are shown in Figs. 4.103 and 4.104.

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Fig. 4.103 P. gingivalis colonies (BHI blood agar)

Fig. 4.104 P. gingivalis colonies (stereomicroscope) P. gingivalis form colonies 1–2 mm in diameter on blood agar. Colonies are rounded and lustrous, with a smooth (or occasionally rough) surface. After 4–8 days of culture, melanin spreads from the edge to the center of the colony to form black colonies. A small number of colonies do not produce melanin

When grown in BM or PYG media, the main terminal acids produced by P. gingivalis are butyric acid and acetic acid, with a small amount of propionic acid, isobutyric acid, isoamyl propionate, and phenylacetic acid produced as well.

P. gingivalis produces indoles, does not produce alpha fucosidase, does not reduce nitrate to nitrite, and does not hydrolyze aesculin and starch. Cells test positive for MDH and glutamate dehydrogenase and negative for glucose

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phosphate dehydrogenase and glucose 6-phosphate acid salt dehydrogenase. These enzymatic characteristics as well as its ability to produce acetic acid are the important features that differentiate P. gingivalis from other morphologically similar bacteria. Its DNA G + C content is 46%–48%. The type strain is ATCC33277 [13].

Cells test negative for sheep cell agglutination (SCAT), but are positive for MDH and glutamate dehydrogenase. P. endodontalis is negative for glucose phosphate dehydrogenase and glucose 6-phosphate dehydrogenase. The genomic G + C content is 49%–51%. The type strain is ATCC 35406.

4.2.8.2 Porphyromonas endodontalis This Gram-negative, obligate anaerobe produces melanin [14]. Cells do not form spores. Examples of bacterial cells are shown in Figs. 4.105 and 4.106. Chlorinated hemoglobin and vitamin K1 are required for culture growth. Characteristic colonies are shown in Figs. 4.107 and 4.108. The primary terminal acid products are N-butyric acid and acetic acid, but a small amount of propionic acid, isobutyric acid, and isoamyl propionate are also produced. The cells do not produce phenylacetic acid. P. endodontalis can hydrolyze gelatin and arginine. Its protein hydrolysis activity is extremely low. It does not hydrolyze aesculin and starch and can produce indoles and hydrogen sulfide. It does not produce alpha fucosidase and does not reduce nitrate to nitrite.

4.2.9

Fig. 4.105 P. endodontalis cells (Gram stain)

Treponema

Spirochetes are a type of slender, curved spiral, highly motile Gram-negative bacteria. Common genera of spirochetes are Spirochaeta, Cristispira, Treponema, and Borrelia. Here we introduce the Treponema in details. Treponema is a genus of commonly found oral bacteria that are closely related to periodontitis and the etiology of implant periarthritis. Species commonly detected in the oral cavity are T. denticola, T. scaliodontum, T. macrodentium, T. oralis, T. intermedia, T. maltophilum, T. socranskii, and T. vincentii. In addition, the gastrointestinal tract and the vagina are also major colonization sites for bacteria of this genus in humans.

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Fig. 4.106 P. endodontalis cells (SEM) P. endodontalis organizes as single cells approximately 0.4–0.6  1.0–2.0 μm in size. They do not form spores, are non-motile, and are Gram-negative

Fig. 4.107 P. endodontalis colonies (BHI blood agar)

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Fig. 4.108 P. endodontalis colonies (stereomicroscope) P. endodontalis on blood agar forms rounded, lustrous colonies with a smooth surface. The colonies have a neat edge and gain progressively deeper dark brown or black

pigmentation (hemoglobin is the main product of porphyrin) after culturing for 4–7 days. Most strains form solid adhesion to the surface of blood agar, but are slow growing in liquid medium

Clinical samples of Treponema are ideally observed with a dark field or a phase contrast microscope. Treponema cells are Gram-negative, but most of the strains do not take up stain easily by Gram staining or Giemsa staining. Silver impregnation stain and Ryu’s stain are better for the observation of Treponema cells. Presently, we commonly use the Congo red negative stain method, as it is not only economic and simple, but the helical cells are also very easy to observe (Fig. 4.109). The morphology of the bacterial cells can be seen under SEM (Figs. 4.110 and 4.111). The outer membrane of Treponema cells is similar to the outer membrane of Gram-negative bacteria cells. The content includes lipids, proteins, and carbohydrates. The lipids are mainly made up of phospholipids and glycolipids, while

the cell wall contains muramic acid, glucosamine, and ornithine. Peptidoglycan accounts for 10% of the dry weight of the cell. The genus Treponema consists of obligate anaerobes, but those species that are pathogenic to humans may be microaerophiles. Treponema are heterotrophic bacteria that mainly metabolize through fermentation. They can use a variety of carbohydrates and amino acid as their carbon source and energy. Treponema species are difficult to grow in artificial culture media, the growth of some species requires long-chain fatty acids from serum, and other species require branchedchain fatty acids. T. denticola, T. vincentii, and T. scaliodontum require cocarboxylase in serum. The genomic G + C content of most Treponema species ranges from 25% to 54% (by Tm). The type species is T. pallidum [15].

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Fig. 4.109 Plaque samples of Treponema cells (Congo red negative staining)

Fig. 4.110 Plaque samples of large, medium and small Treponema cells (SEM)

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Fig. 4.111 Plaque samples of Treponema and Borrelia cells (SEM) Treponema cells measure 0.1–0.4  5–20 μm. They form a spiral rod and can be either a tight, regular, or irregular spiral. Cells have one or more axial flagella inserted into the plasma membrane at opposite ends of the rod. In old cultures, huge bubbles and spiral spheres can be observed. Large, medium, and small spirochetes can be observed in clinical samples. The spiral-shaped cells measure 0.2–0.5  3–20 μm and form regular or irregular loose spirals. The cells stain Gram-negative

References 1. Qin X, Galloway-Pena JR, Sillanpaa J, Roh JH, Nallapareddy SR, Chowdhury S, Bourgogne A, Choudhury T, Muzny DM, Buhay CJ, Ding Y, Dugan-Rocha S, Liu W, Kovar C, Sodergren E, Highlander S, Petrosino JE, Worley KC, Gibbs RA, Weinstock GM, Murray BE. Complete genome sequence of Enterococcus faecium strain TX16 and comparative genomic analysis of Enterococcus faecium genomes. BMC Microbiol. 2012;12:135. 2. Kageyama A, Benno Y, Nakase T. Phylogenetic evidence for the transfer of Eubacterium lentum to the genus Eggerthella as Eggerthella lenta gen. nov, comb. nov. Int J Syst Bacteriol. 1999;49 (pt4):1725–32. 3. Bruggemann H, Henne A, Hoster E, Liesegang H, Wiezer A, Strittmatter A, Hujer S, Durre P, Gottschalk G. The complete genome sequence of Propionibacterium acnes, a commensal of human skin. Science. 2004;305(5684):671–3. 4. Eiken M. Studies on an anaerobic, rod-shaped, gramnegative microorganism: Bacteroides corrodens N. sp. Acta Pathol Microbiol Scand. 1958;43(4):404–16. 5. Kapatral V, Anderson I, Ivanova N, Reznik G, Los T, Lykidis A, Bhattacharyya A, Bartman A, Gardner W, Grechkin G, Zhu L, Vasieva O, Chu L, Kogan Y,

Chaga O, Goltsman E, Bernal A, Larsen N, D'Souza M, Walunas T, Pusch G, Haselkorn R, Fonstein M, Kyrpides N, Overbeek R. Genome sequence and analysis of the oral bacterium Fusobacterium nucleatum strain ATCC 25586. J Bacteriol. 2002;184(7):2005–18. 6. Calcutt MJ, Foecking MF, Nagaraja TG, Stewart GC. Draft genome sequence of Fusobacterium necrophorum subsp. funduliforme bovine liver abscess isolate B35. Genome Announc. 2014;2(2):e00412-14. 7. Marshall BJ, Warren JR. Unidentified curved bacilli in the stomach of patients with gastritis and peptic ulceration. Lancet. 1984;1(8390):1311–5. 8. Tomb JE, White O, Kerlavage AR, Clayton RA, Sut-ton GG, Fleischmann RD, Ketchum KA, Klenk HP, Gill S, Dougherty BA, Nelson K, Quackenbush J, Zhou Kirkness EE, Peterson S, Loftus B, Richardson D, Dodson R, Khalak HG, Glodek A, McKenney K, Fitze-gerald LM, Lee N, Adams MD, Hickey EK, Berg DE, Gocayne JD, Utterback TR, Peterson JD, Kelley JM, Cotton MD, Weidman JM, Fujii C, Bowman C, Wat-they L, Wallin E, Hayes WS, Borodovsky M, Karp PD, Smith HO, Fraser CM, Venter JC. The complete genome sequence of the gastric pathogen Helicobacter pylori. Nature. 1997;388(6642):539–47. 9. Slots J. The predominant cultivable organisms in juvenile periodontitis. Scand J Dent Res. 1976;84(1):1–10.

210 10. Ruan Y, Shen L, Zou Y, Qi Z, Yin J, Jiang J, Guo L, He L, Chen Z, Tang Z, Qin S. Comparative genome analysis of Prevotella intermedia strain isolated from infected root canal reveals features related to pathogenicity and adaptation. BMC Genomics. 2015;16 (1):122. 11. Harding GK, Sutter VL, Finegold SM, Bricknell KS. Characterization of Bacteroides melaninogenicus. J Clin Microbiol. 1976;4(4):354–9. 12. Koreeda Y, Hayakawa M, Ikemi T, Abiko Y. Isolation and characterisation of dipeptidyl peptidase IV from Prevotella loescheii ATCC 15930. Arch Oral Biol. 2001;46(8):759–66. 13. Naito M, Hirakawa H, Yamashita A, Ohara N, Shoji M, Yukitake H, Nakayama K, Toh H, Yoshimura F, Kuhara S, Hattori M, Hayashi T, Nakayama K. Determination of the genome sequence of Porphyromonas ginsialis strain ATCC 33277 and

Q. Yuan et al. genomic comparison with strain W83 revealed extensive genome rearrangements in P. gingivalis. DNA Res. 2008;15(4):215–25. 14. van Winkelhoff AJ, van Steenbergen TJ, de Graaff J. Porphyromonas (Bacteroides) endodontalis: its role in endodontal infections. J Endod. 1992;18(9):431–4. 15. Fraser CM, Norris SJ, Weinstock GM, White O, Sutton GG, Dodson R, Gwinn M, Hickey EK, Clayton Y, Ketchum KA, Sodergren E, Hardham JM, McLeod MP, Salzberg S, Peterson J, Khalak H, Richardson D, Howell JK, Chidambaram M, Utterback T, McDonald L, Artiach P, Bowman C, Cotton MD, Fujii C, Garland S, Hatch B, Horst K, Roberts N, Sandusky M, Weidman J, Smith HO, Venter JC. Complete genome sequence of Treponema pallidum, the syphilis spirochete. Science. 1998;281 (5375):375–88.

5

Oral Mucosal Microbes Biao Ren, Lei Cheng, Xian Peng, Yuqing Li, Yan Li, Yujie Zhou, Chengguang Zhu, and Xi Chen

Abstract

This chapter mainly covers the microbes that cause mucosal infection. Many microorganisms can invade the oral mucosa, causing infectious stomatitis. Infectious stomatitis is classified as primary and secondary. The former refers to the inflammatory diseases that occur on the oral mucosa due to the direct invasion of pathogenic microorganisms, such as coccal stomatitis, necrotizing gingival stomatitis, and acute gangrenous stomatitis. The latter refers to the manifestation of systemic infection caused by pathogenic microorganisms in the oral mucosa, such as diphtheria and staphylococcal sepsis. Both primary and secondary infections can be divided into bacterial infection, viral infection, fungal infection, spirochetes infection, actinomycetes infection, mycoplasma

B. Ren (*) · X. Peng · Y. Li · Y. Li · Y. Zhou · C. Zhu · X. Chen State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China e-mail: [email protected] L. Cheng State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China

infection, chlamydia infection, and rickettsial infection according to the different pathogenic microorganisms. The biological and pathogenic characteristics of representative Grampositive bacteria, Gram-negative bacteria, mycoplasma, fungi, and virus were introduced in this chapter. Gram staining, plate culture, colony, and scanning electron microscope (SEM) images of each microorganism were also provided. Keywords

Staphylococcus · Moraxella · Neisseria · Mycoplasma · Fungi · Virus

5.1 5.1.1

Gram-Positive Bacteria Staphylococcus

Staphylococcus aureus is Gram-positive species of bacteria [1]. Its peptidoglycan structure is L-Lys-Gly5-6, and the cell wall teichoic acid is formed by ribitol polymerization. Cellular morphology is shown in Figs. 5.1 and 5.2. The genomic G + C content is 32–36%, and the type strains are ATCC126000, NCTC8352, and CCM885. S. aureus is a facultative anaerobe but also grows well under aerobic conditions (Figs. 5.3, 5.4, and 5.5). After incubation on semisolid thioglycollate medium, colonies grow rapidly

# Zhejiang University Press 2020 X. Zhou, Y. Li (eds.), Atlas of Oral Microbiology: From Healthy Microflora to Disease, https://doi.org/10.1007/978-981-15-7899-1_5

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Fig. 5.1 S. aureus cells (Gram stain)

Fig. 5.2 S. aureus cells (SEM) The cells of S. aureus are characterized as being spherical (0.5–1.5 μm in diameter), Gram-positive cocci arranged as single cells, pairs, tetrads, and clusters

are uniform and dense. The final pH of glucose broth incubated with S. aureus is always between pH 4.3 and 4.6.

S. aureus is associated with human infections, such as facial furuncle and carbuncle, loose connective tissue inflammation, and tumor postoperative wound infections.

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Fig. 5.3 Golden pigment of S. aureus incubated on common agar plate

Fig. 5.4 β-hemolytic zone of S. aureus incubated on blood BHI agar plate

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Fig. 5.5 Colonies of S. aureus (stereomicroscope) S. aureus is able to produce golden pigments on common agar plate. The colonies are round, raised, and shiny opaque and have a smooth surface and a whole edge. The diameter is 2–3 mm. A β-transparent hemolytic zone surrounds the colony when S. aureus is incubated on blood BHI agar

Fig. 5.6 Cells of S. mitis (Gram stain)

5.1.2

Streptococcus

5.1.2.1 Streptococcus mitis S. mitis cells are Gram-positive and spherical or elliptical in shape (about 0.6–0.8 μm in diameter) [2]. They can form long chains in broth culture

(Fig. 5.6). The G + C content of the S. mitis genome is 38–39%, and its type strain is NCTC3165. S. mitis cells grow variably at 45
C and the final pH value of a culture in glucose broth is around pH 4.2–5.8. Cells in broth culture change

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Fig. 5.7 Colonies of S. mitis (TPY agar plate)

Fig. 5.8 Colonies of S. mitis (BHI agar plate)

from rough to smooth following subculture and cells that are grown under aerobic conditions can be smooth or rough. TPY agar is the common medium on which S. mitis is grown (Fig. 5.7).

Significant α-hemolytic reaction can be observed when S. mitis is grown on blood agar plates (Fig. 5.8). S. mitis forms small broken-glass-like colonies on sucrose agar plates, and very few

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Fig. 5.9 Colonies of S. mitis (stereomicroscope)

Fig. 5.10 Cells of S. pyogenes (Gram stain)

strains can form the typical sticky colonies. Colonies observed by stereomicroscope are shown in Fig. 5.9. Biochemical reactions: S. mitis can ferment glucose, maltose, sucrose, lactose, and salicin to produce acid, and some strains can also ferment raffinose and trehalose to produce acid. However, inulin, mannitol, sorbitol, glycerol, arabinose, and xylose cannot be fermented by S. mitis.

Colonization characteristics: S. mitis can be detected in human saliva, oral mucosa, sputum, or excrement. It is a common oral streptococcus and a member of the oral microflora.

5.1.2.2 Streptococcus pyogenes S. pyogenes belongs to Lancefield group A and is Gram-positive (Fig. 5.10). Cells of this species have a diameter of 0.5–1.0 μm and are usually

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Fig. 5.11 Cells of S. pyogenes (SEM)

spherical. However, cells from old cultures may be oval. Moreover, S. pyogenes cells are often arranged in short, medium, or long chains, but most cells in broth culture form long chains. Cells of clinical isolates are usually arranged in pairs. S. pyogenes cells observed by stereomicroscope are shown in Fig. 5.11. The G + C content of the S. pyogenes genome is 34.5–38.5% (Tm method), and its type strain is ATCC12344. S. pyogenes is a facultative anaerobes and the optimal temperature for growth is 37
C. A nutrient-rich medium is required for S. pyogenes growth, and both blood and serum can promote its growth. Colonies growing on MS agar are shown in Fig. 5.12. Cells from overnight bloodagar plate culture have three different colony types (Figs. 5.13, 5.14, and 5.15): 1. Colonies with β-hemolytic zone 2. Mucoid colonies 3. Lackluster colonies In fact, the type of colonies is mainly related to the growth condition and production of hyaluronidases.

S. pyogenes can produce several pathogenic virulence factors, including bacteriocins, hemolysin 0, streptokinase, NADH enzyme, hyaluronidase, and antigen of M protein [3]. The main sites of colonization for S. pyogenes are the dental plaque, hypopharynx, and the upper respiratory tract. Clinical samples can be isolated from skin lesions, inflammatory secretions, or blood. S. pyogenes can also be found in loose connective tissue inflammation in the maxillofacial region, pulpitis, or infection after exelcymosis.

5.1.2.3 Streptococcus pneumoniae S. pneumoniae cells are spherical or elliptic (about 0.5–1.25 μm in diameter) and mostly arranged in pairs. They can occasionally be found in short chains or as single cells. Typical cells of this species are lanceted and arranged in pairs, and the extracellular capsule can be observed by capsule staining. However, cells can easily form chains after continuous culture. Cells of S. pneumoniae are Gram-positive, but can change to be Gram-negative when aged (Figs. 5.16 and 5.17). S. pneumoniae cells

218 Fig. 5.12 Colonies of S. pyogenes (MS agar plate)

Fig. 5.13 Colonies of S. pyogenes with β-hemolytic zone (BHI-blood agar plate)

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Fig. 5.14 Mucoid colonies of S. pyogenes (BHI-blood agar plate, stereomicroscope)

Fig. 5.15 Lackluster colonies of S. pyogenes (BHI-blood agar plate, stereomicroscope)

observed by SEM are shown in Fig. 5.18. The G + C content of the S. pneumoniae genome is 30% (by Tm method) or 42% (by Bd method), and its type strain is NCTC7465.

S. pneumoniae is a kind of facultative anaerobe, but could generate a substantial amount of H2O2 under aerobic conditions. Growth of S. pneumoniae requires a complex medium rich in nutrients, including blood, serum, or ascites, as

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Fig. 5.16 Cells of S. pneumoniae (Gram stain)

Fig. 5.17 Capsule of S. pneumoniae cells (capsule stain)

well as vitamin B. Cells of this species grow well on BHI-blood agar plates and form rooflike, reflective, and smooth colonies or rugose, mycelium-like, rough colonies (Fig. 5.19). Cells of S. pneumoniae can generate a great deal of capsular polysaccharides and therefore often form sticky colonies (Fig. 5.20). In fact, this capsular polysaccharide is the species-specific antigen and virulence factor of S. pneumoniae (SSS).

S. pneumoniae cells observed by stereomicroscope are shown in Fig. 5.21. The final pH value of a culture in glucose broth is pH 5.0. Compared to other streptococci, the addition of bile or cholate is required for the isolation and identification of S. pneumoniae. S. pneumoniae carries out metabolic reactions through fermentation. Cells of this species can ferment glucose, galactose, fructose, sucrose,

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Fig. 5.18 Cells of S. pneumoniae (SEM)

Fig. 5.19 Colonies of S. pneumoniae (BHI-blood agar plate)

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Fig. 5.20 Colonies of S. pneumoniae (BHI-blood agar plate, stereomicroscope)

Fig. 5.21 Colonies of S. pneumoniae (stereomicroscope)

maltose, raffinose, and inulin to produce acid, and some strains can also ferment mannitol, but not dulcite and sorbitol. Bacteriolysis of S. pneumoniae cells by bile is positive, and this characteristic is helpful in distinguishing S. pneumoniae from other streptococci. S. pneumoniae mainly inhabits the upper respiratory tracts of healthy individuals or livestock and can be isolated from clinical samples of

amygdalitis, pneumonia, meningitis, and otitis media [4]. However, the colonization, distribution, and pathogenicity of this species in oral cavity are still unknown.

5.1.2.4 Streptococcus vestibularis S. vestibularis gets its name because this species was first isolated from the vestibule of the human oral cavity. The cells of S. vestibularis are Gram-

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Fig. 5.22 Cells of S. vestibularis (Grampositive coccus)

Fig. 5.23 Cells of S. vestibularis (SEM)

positive, spherical (about 1 μm in diameter), and arranged in chains (Fig. 5.22). S. vestibularis cells observed by SEM are shown in Fig. 5.23. The

G + C content of S. vestibularis genome is 38–40%, and its type strain is TC12166 (¼MM1).

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Fig. 5.24 Colonies of S. vestibularis (MS agar plate)

S. vestibularis is a facultative anaerobe and the optimal temperature for growth is 37
C, but cells of this species can also still grow at 10 and 45
C. Except the type strain MM, the majority of strains belonging to this species can grow in media containing 10% bile. However, the species cannot grow in medium containing 4% NaCl, 0.004% crystal violet, or 40% bile. Alpha-hemolysis can be detected for all strains of this species growing on horse blood agar plates. In addition, black blue, lackluster, and umbilicate colonies (about 2–3 μm in diameter) with wavy edges can be observed when cells are grown on MS agar plates under anaerobic conditions (37
C, 72 h culture), while black blue, glossy, and raised colonies (about 1–2 μm in diameter) with neat edges can be observed when cells are grown on MS agar plates under aerobic conditions (Fig. 5.24). Colonies observed by stereomicroscope are shown in Fig. 5.25. S. vestibularis can ferment N-acetylglucosamine, myricitrin, fructose, galactose, glucose, lactose, maltose, mannose, salicin, and sucrose to produce acid, but they do not ferment adonitol, arabinose, dextrin, dulcite, fucose, glycerol, glycogen, inositol, inulin,

mannitol, melezitose, D-Melibiose, raffinose, ribitol, sorbitol, starch, and xylose. The majority of strains of this species can ferment cellulose and amygdalin to produce acid, and a minority of strains of this species can ferment trehalose and D-glucosamine to produce acid. S. vestibularis can also generate urease and H2O2, but not extracellular or intracellular polysaccharides. It also does not generate ammonia from arginine. In addition, the majority strains of this species can produce butanone with two hydroxyl groups and can hydrolyze aesculin and starch [5]. S. vestibularis is mainly isolated from the mucosa or vestibule of the human oral cavity.

5.2 5.2.1

Gram-Negative Bacteria Escherichia

Here we introduce Escherichia coli in details. Commonly known as E. coli, this is the most common bacteria belonging to the family Enterobacteriaceae found in the human body. E. coli is the first Gram-negative bacillus detected

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Fig. 5.25 Colonies of S. vestibularis (stereomicroscope)

Fig. 5.26 E. coli cells (Gram stain)

in the neonatal oral cavity and is thought to be transmitted from the mother during vaginal birth. E. coli is a Gram-negative bacillus (Figs. 5.26 and 5.27). It is a facultative anaerobe with low nutritional requirement, which can grow well

under aerobic condition. Characteristic colonies are shown in Figs. 5.28 and 5.29. E. coli occasionally can cause maxillofacial infections.

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Fig. 5.27 E. coli cells (SEM) The size of E. coli cell is (0.4–0.7)  (1–3) μm. Gram stain is negative

Fig. 5.28 E. coli colonies (BHI blood agar)

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Fig. 5.29 E. coli colonies (stereomicroscope) E. coli colonies are soft gray white with a diameter of 2–3 mm

5.2.2

Haemophilus

Haemophilus is a genus of Gram-negative facultative anaerobic bacilli, named due to their requirement for blood during growth. Most bacteria of this genus are part of the normal microflora of the oral cavity and of the nasopharynx in human and animals. The main colonization site within the oral cavity is dental plaque, followed by saliva and soft palate. Species detected in the oral cavity include H. actinomycetemcomitans, H. influenzae, H. parainfluenzae, H. aphrophilus, H. paraphrophilus, H. parahaemolyticus, H. paraprohaemolyticus, and H. segnis. In the latest classification, H. actinomycetemcomitans, H. aphrophilus, and H. segnis were classified into a new genus, Aggregatibacter, and renamed A. actinomycetemcomitans, A. aphrophilus, and A. segnis, respectively. Generally, Haemophilus cells are club-shaped or rod-shaped and less than 1 μm in width. The length of cells can be short to medium. Sometimes cells are filamentous and can be pleiomorphic. Bacteria of this genus are Gramnegative, do not produce spores, have no capsule, and are non-motile.

Haemophilus are chemoorganotrophic bacteria. After 48 h culture on blood agar surface at 37
C, most species will form flat or raised, colorless or pale yellow, smooth colonies with a diameter of 0.5–2.0 mm. A few species like H. parainfluenzae form rough colonies. H. parahaemolyticus, and H. paraprohaemolyticus can produce β-hemolytic reaction on blood agar. In broth containing glucose, the terminal acid metabolites are acetic acid, lactic acid, and succinic acid. Most oral strains of Haemophilus are non-pathogenic members of the normal human oral mucosa and upper respiratory tract. Occasionally, they can cause mixed infections such as periodontal abscess or jaw infection as conditional pathogens. The genomic G + C content ranges from 37% to 44% (by Tm method). The type species is H. influenzae [6]. H. influenzae is a Gram-negative bacillus. Cell morphology is shown in Figs. 5.30 and 5.31. H. influenzae is facultative anaerobe. Atmosphere supplemented with 5–10% CO2 is the most suitable growth environment. Cultures require growth factors from blood, especially factor X and factor V. Chocolate agar is the preferred

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Fig. 5.30 H. influenzae cells (Gram stain)

Fig. 5.31 H. influenzae cells (SEM) H. influenzae cells are Gram-negative, clubshaped, or small rods, with dimensions of 0.3–0.5  0.5–3.0 μm. The cells have no spores or capsule and are non-motile

growth medium. Characteristic colonies are shown in Figs. 5.32, 5.33, and 5.34. H. influenzae can ferment glucose and sucrose to produce acid, but fermentations of galactose, fructose, maltose, and xylose do not produce acid.

They can also reduce nitrate, but not nitrite. The species tests positive for catalase, while the o-nitrophenyl-β-D-galactopyranoside (ONPG) test show up negative.

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Fig. 5.32 H. influenzae colonies (BHI blood agar)

Fig. 5.33 H. influenzae viscous colonies (stereomicroscope)

H. influenzae can be detected in the nasopharynx of up to 75% of healthy children. However, the infection rate in adults is very low. The

detection rate of capsule-producing strains in the healthy nasopharynx is only 3–7%.

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Fig. 5.34 H. influenzae colonies (stereomicroscope) After cultivation on chocolate agar for 24 h, convex, gray, translucent smooth colonies form with a diameter of 0.5–1 mm. Strains with capsule usually form 1–3 mm large viscous colonies

The genomic G + C content is 39% (by Tm method). The type strain is NCTC8143 (biological variant type II without capsule).

5.2.3

but can cause catarrhal inflammation, such as acute pharyngitis and otitis media. The genomic G + C content is 40–43%. The type strain is ATCC25238 (NCTC11020).

Moraxella 5.2.4

Moraxella, Neisseria, and Kingella all belong to the family Neisseriaceae. Here we introduce Moraxella catarrhalis in details. M. catarrhalis are common bacteria belonging to the genus Moraxella found in the oral cavity. They were previously known as Neisseria catarrhalis and Branhamella catarrhalis. The cells are Gram-positive cocci and are shown in Figs. 5.35 and 5.36. M. catarrhalis are aerobic and can grow on agar medium. However, cultures grow better on blood agar. Characteristic colonies are shown on Figs. 5.37 and 5.38. The main site of colonization is the upper respiratory tract [7] which is the main habitat. This species generally does not cause disease

Neisseria

Neisseria are aerobic Gram-negative diplococci belonging to the family Neisseriaceae, which mainly colonize the human oral cavity and nasopharynx. Most Neisseria are members of the normal microflora of the human body and are usually non-pathogenic. However, N. meningitidis and N. gonorrhoeae are important pathogens. Common Neisseria in the oral cavity are N. sicca and N. subflava. Neisseria are less nutrition-demanding aerobic bacteria that can grow easily on agar medium. Most Neisseria cells are spherical, but occasionally short rods are observed with a diameter of 0.6–1.0 μm. Many are arranged in pairs with a flat adjacent surface. Neisseria cells may have a capsule and pili, but no endospores and flagella.

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Fig. 5.35 M. catarrhalis cells (Gram stain)

Fig. 5.36 M. catarrhalis cells (SEM) M. catarrhalis cells are spheres or short rods and are typically kidneyshaped. Nearly kidneyshaped diplococci are observed in clinical specimens. The cells do not produce spores and are non-motile. Cells stain Gram-negative

Also known as meningococcus, this is the pathogen behind meningococcal meningitis [8]. N. meningitides are Gram-negative cocci (Figs. 5.39 and 5.40). They are aerobic, but

grow better under 5–8% CO2. Growth requires nutrient agar with blood serum or blood. Typical colonies are shown in Figs. 5.41, 5.42, and 5.43.

232 Fig. 5.37 M. catarrhalis colonies (BHI blood agar)

Fig. 5.38 M. catarrhalis colonies (stereomicroscope) After 48 h culture on blood agar, smooth, opaque, relatively flat, gray colonies form with a diameter of about 2 mm

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Fig. 5.39 N. meningitides cells (Gram stain)

Fig. 5.40 N. meningitides cells (SEM) N. meningitidis cells are ovoid or spherical, kidneyshaped, or bean-shaped diplococci; Gram stain is negative

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234 Fig. 5.41 N. meningitides colonies (BHI blood agar)

Fig. 5.42 N. meningitides colonies (BHI chocolate agar)

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Fig. 5.43 N. meningitides colonies (stereomicroscope) The colony size of N. meningitides depends on medium on which they are grown. After 18–24 h culture, the colony diameter reaches 1.0–1.5 mm on the surface of nutrient-rich blood agar. They appear as round, smooth, transparent, dew-like colonies without hemolysis. The colonies grow best on chocolate blood agar. On MuellerHinton agar, colonies are round, smooth, shiny, and translucent

The genomic G + C content is 50–52% (by Tm method). The type strain is ATCC13077.

5.3

Mycoplasma

Mycoplasma can live independently with no cell wall. They are the smallest prokaryotic microbial cells and can be isolated from normal human and animal respiratory mucosa. In the oral cavity, M. pneumoniae, M. oralis, M. salivalis, and M. nominis can be isolated. Mycoplasma are the smallest of all prokaryotic microbial cells (Figs. 5.44 and 5.45). Mycoplasma have high nutritional demands and can grow on PPLO agar with beef heart infusion and 10–20% horse serum. The serum provides Mycoplasma with the cholesterol and long-chain fatty acids required for growth. The optimum pH for Mycoplasma culture is pH 7.8–8.0. Cells may die when the pH drops below pH 7.0. Mycoplasma are aerobic or facultative anaerobic microorganisms, but they usually grow better in an aerobic environment. The best culture

environment for initial isolation is atmospheric conditions supplemented with 5% CO2 or anaerobic conditions with 5% CO2 and 95% N2. Mycoplasma colonies are small and can only be observed under a light microscope at low magnification or a dissecting microscope. Characteristic colonies are shown in Figs. 5.46 and 5.47. Mycoplasma and L type bacteria are similar: 1. they both lack a cell wall and the cell is pleomorphic; 2. they can both pass through an antimicrobial filter. The main difference between the two are as follows: 1. Mycoplasma are independent microbes and L type bacteria are variants of normal bacterial cells that have a cell wall (most L type cells will revert to their original form when the induction factor is eliminated); 2. Mycoplasma growth requires cholesterol (10–20% serum in the medium), while the growth of L-type bacterial does not; 3. L-type bacteria fade easily after Diane staining while Mycoplasma do not fade easily. Mycoplasma usually colonize the throat, bacterial biofilm, or tartar found in the oral cavity. M. pneumoniae is one of the common causes of

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Fig. 5.44 M. pneumoniae cells (Gram stain) The signet-ring-shaped cell of Mycoplasma is Gramnegative, and the size of the cell is 0.2–0.3 μm and is normally smaller than 1.0 μm. Cells have no cell wall. Protein and lipids form the outer cell membrane and the cells are obviously pleomorphic, with spherical, rod-like, bar-like, and filamentous morphologies visible under the microscope. The typical cell is shaped like a signetring. Cells are Gramnegative, light purple with Giemsa stain

Fig. 5.45 M. pneumoniae signet-ring cell (Giemsa stain) M. pneumoniae cell is light purple with Giemsa stain

acute respiratory infections [9]. Although previous studies reported that Mycoplasma can be separated from root canal infections, gingivitis, and periodontitis clinical specimens, the distribution and pathogenicity in the oral cavity are still unclear.

5.4 5.4.1

Fungi Saccharomyces

The family Saccharomycetaceae belongs to the phylum Ascomycota and includes the common

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Fig. 5.46 M. pneumoniae omelet -like colonies

Fig. 5.47 Mulberry-shaped colonies of M. pneumoniae Typical colonies of the Mycoplasma are omelet-like in shape. Colony diameter is about 10–15 μm, the center is round and opaque and extends into the medium; the edge of the colony is thin and flat, forming a transparent or

semi-transparent area. Other characteristic colonies of the Mycoplasma include as mulberry-shaped colonies of M. pneumoniae. Mycoplasma from the mouth and saliva form visible comet-like colonies in semi-solid medium (mostly in the middle and bottom of the culture medium)

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Fig. 5.48 C. albicans cells (Gram-positive)

genera Candida and Saccharomyces. Common species that can be detected in the mouth include C. albicans, S. tropicalis, S. candidaglabrata, S. parapsilosis, S. krusei, S. guilliermondii, and S. dulbilin, the most commonly detected of these is C. albicans. Saccharomyces is common symbiotic yeast that inhabits the gastrointestinal tract, respiratory tract, and the vaginal mucosa and can only infect the host under specific conditions. Therefore, they are known as conditional pathogens. The thallus of this group of yeast is circular or ovoid and consists of a cell wall, cell membrane, cytoplasm, and nucleus. They reproduce by budding. Spore elongate into the germ tube, but do not detach from the thallus and form long pseudohyphae.

5.4.1.1 Candida albicans C. albicans is common fungus found in the oral cavity [10] and can be detected in the oral cavity of 30–35% of healthy adults. The percentage of detection is even higher in the neonate oral cavity. The main sites of colonization are the mucosa (buccal mucosa, palate, etc.), saliva, oral prosthetics, dentures, etc. C. albicans can cause acute or chronic oral candidiasis such as pseudomembranous candidiasis (thrush), denture stomatitis, and Candida leukoplakia. Oral

candidiasis and other oral Candida infections are often secondary infections in AIDS patients. The Candida cell is large and spherical and stains Gram-positive (Fig. 5.48). The cell in its budding form can be visualized by SEM (Figs. 5.49 and 5.50). Other strains are shown in Figs. 5.51, 5.52, and 5.53. Candida is an aerobic microorganism and grows best in temperatures ranging from 30 to 37
C, with 24–48 h incubation time. Sabouraud agar is the standard medium used for its cultivation (Figs. 5.54, 5.55, and 5.56). CHROMagar Candida agar is a commonly used selective medium, as different strains form colonies with different colors on the surface of the medium, which can be used to identify strains from a clinic sample (Fig. 5.57). C. albicans colonies are shown in Figs. 5.58 and 5.59. C. tropicalis colonies and cells are shown in Fig. 5.60. C. glabrata colonies and cells are shown in Fig. 5.61 and 5.62. C. krusei colonies and cells are shown in Figs. 5.63 and 5.64. Germ tube formation assay (Figs. 5.65 and 5.66) and thick-walled spore formation assay (Figs. 5.67, 5.68, and 5.69) are commonly used to identify the C. albicans strains in a sample. Candida can ferment glucose, maltose, and sucrose and produces acid, but does not ferment lactose.

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Fig. 5.49 C. albicans cells (SEM)

Fig. 5.50 C. albicans cells (SEM) The yeast cell is 2  4 mm in size, ovoid or spherical, Gram-positive. The budding cell is visible under SEM

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Fig. 5.51 C. tropicalis cells (Gram stain)

Fig. 5.52 C. glabrata cells (Gram stain)

5.5

Virus

Here we introduce herpes simplex virus (HSV) in details. Herpesvirus is an enveloped DNA virus of medium size. More than 100 strains have been discovered. Of these, HSV, Varicella-zoster

virus, cytomegalovirus, and Epstein-Barr virus are related to human oral mucosal infections. These viruses share a icosahedral protein nucleocapsid formed by 162 subunits that protects the double-stranded linear DNA There is a lipoprotein envelope surrounding the nucleocapsid. The

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Fig. 5.53 C. krusei cells (Gram stain)

Fig. 5.54 C. albicans colonies (Sabouraud’s agar)

outer diameter of the virus is approximately 150–200 nm. HSV is the type virus of the herpesviruses. It causes vesicular lesions of the skin and mucosa and is a common virus found in humans [11].

Research has shown that 30–90% of humans possess antibodies against HSV, indicating that they have previously been infected. HSV can easily invade ectodermal tissues including neurons, skin, and mucosal layers.

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Fig. 5.55 C. albicans colonies (Sabouraud’s agar slant)

Fig. 5.56 C. albicans colonies (Sabouraud’s agar, stereomicroscope) C. albicans grow well in Sabouraud’s agar and form milk white, smooth surface, and softer yeast-like colony and yeast smell. When the hatching time extended, it can form rough surface and faviform or folding rough colony

The double-stranded DNA contained within the HSC core encodes the genetic information necessary for virus production. The DNA consists of two fragments, one long and one short, linked by a covalent bond. The long fragment makes up 82% of the viral genetic material and the short fragment, 18%. The percentage GC content is approximately 68%, which is fairly high. The

nucleocapsid is made up of protein subunits, each of which measures 9.5  12.5 nm, with a 4 nm hole at the center. The envelope is a lipid structure rich in proteoglycans and lipoproteins, within which six viral antigens are located: gB, gC, gD, gE, gG, and gH. In humans, HSV infection is very common, and patients and healthy carriers can be sources

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Fig. 5.57 C. albicans colonies (CHROMagar Candida agar)

Fig. 5.58 C. albicans green colonies (CHROMagar Candida agar)

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Fig. 5.59 C. albicans colonies (CHROMagar Candida agar, stereomicroscope)

Fig. 5.60 C. tropicalis blue colonies (CHROMagar Candida agar)

of infection. Infection is mainly caused by droplets or coming into contact with saliva or other herpesvirus-containing fluids. The fetus can be infected by passage through the birth canal. In oral leukoplakia, HSV can be isolated

from oral squamous cell carcinoma, and there is evidence to support that HSV may be an important factor in oral precancerous lesions or squamous cell carcinoma. The proliferation of HSV in Vero cells is shown in Figs. 5.70 and 5.71.

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Fig. 5.61 S. C. glabrata purple colonies (CHROMagar Candida agar)

Fig. 5.62 S. C. glabrata purple colonies (CHROMagar Candida agar, stereomicroscope)

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246 Fig. 5.63 C. krusei pink colonies (CHROMagar Candida agar)

Fig. 5.64 C. krusei pink colonies (CHROMagar Candida agar, stereomicroscope)

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Fig. 5.65 C. albicans sprouting spores and germ tube (Crystal violet stain)

Fig. 5.66 C. albicans sprouting spores and germ tube (lactophenol stain) Germ tube formation assay involves inoculating the isolated strain on 0.5–1 ml human or sheep serum. Incubate for 2–4 h at 37
C, stain with crystal violet stain or lactophenol stain, and observe under the microscope. Sprouting spores and germ tubes are visible under oil immersion

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248 Fig. 5.67 C. albicans colonies (1% Twain cornmeal agar)

Fig. 5.68 Pseudohypha and chlamydospores of C. albicans (1% Twain cornmeal agar, crystal violet stain)

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Fig. 5.69 Pseudohypha and chlamydospores of C. albicans (1% Twain cornmeal agar, lactophenol staining)

Fig. 5.70 Proliferation of HSV in Vero cells (inverted microscope)

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Fig. 5.71 Proliferation of HSV in Vero cells (TEM)

References 1. Kuroda M, Ohta T, Uchiyama I, Baba T, Yuzawa H, Kobayashi I, Cui L, Oguchi A, Aoki K, Nagai Y, Lian J, Ito T, Kanamori M, Matsumaru H, Maruyama A, Murakami H, Hosoyama A, MizutaniUi Y, Takahashi NK, Sawano T, Inoue R, Kaito C, Sekimizu K, Hirakawa H, Kuhara S, Goto S, Yabuzaki J, Kanehisa M, Yamashita A, Oshima K, Furuya K, Yoshino C, Shiba T, Hattori M, Ogasawara N, Hayashi H, Hiramatsu K. Whole genome sequencing of meticillin-resistant Staphylococcus aureus. Lancet. 2001;357(9264):1225–40. 2. Denapaite D, Bruckner R, Nuhn M, Reichmann P, Hen- rich B, Maurer P, Schahle Y, Selbmann P, Zimmermann W, Wambutt R, Hakenbeck R. The genome of Streptococcus mitis B6-what is a commensal? PLoS One. 2010;5(2):e9426. 3. Ferretti JJ, McShan WM, Ajdic D, Savic DJ, Savic G, Lyon K, Primeaux C, Sezate S, Suvorov AN, Kenton S, Lai HS, Lin SP, Qian Y, Jia HG, Najar EZ, Ren Q, Zhu H, Song L, White J, Yuan X, Clifton SW, Roe BA, McLaughlin R. Complete genome sequence of an M1 strain of Streptococcus pyogenes. Proc Natl Acad Sci U S A. 2001;98(8):4658–63.

4. Tettelin H, Nelson KE, Paulsen IT, Eisen JA, Read TD, Peterson S, Heidelberg J, DeBoy RT, Haft DH, Dodson RJ, Durkin AS, Gwinn M, Kolonay JF, Nelson WC, Peterson JD, Umayam LA, White O, Salzberg SL, Lewis MR, Radune D, Holtzapple E, Khouri H, Wolf AM, Utterback TR, Hansen CL, McDonald LA, Feldblyum TV, Angiuoli S, Dickinson T, Hickey EK, Holt IE, Loftus BJ, Yang E, Smith HO, Venter JC, Dougherty BA, Morrison DA, Hollingshead SK, Fraser CM. Complete genome sequence of a virulent isolate of Streptococcus pneumoniae. Science. 2001;293 (5529):498–506. 5. Delorme C, Abraham AL, Renault P, Guedon E. Genomics of Streptococcus salivarius, a major human commensal. Infect Genet Evol. 2014; https:// doi.org/10.1016/j.meegid.2014.10.001. pii: s15671348(14)00372-4 6. Hood DW. The genome sequence of Haemophilus influenzae. Methods Mol Med. 2003;71:147–59. 7. de Vries SP, van Hijum SA, Schueler W, Riesbeck K, Hays JP, Hermans PW, Bootsma HJ. Genome analysis of Moraxella catarrhalis strain BBH18, a human respiratory tract pathogen. J Bacteriol. 2010;192 (14):3574–83.

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8. Schoen C, Kischkies L, Elias J, Ampattu BJ. Metabolism and virulence in Neisseria meningitidis. Front Cell Infect Microbiol. 2014;4:114. 9. Loens K, Goossens H, Ieven M. Acute respiratory infection due to Mycoplasma pneumoniae: current status of diagnostic methods. Eur J Clin Microbiol Infect Dis. 2010;29(9):1055–69.

251 10. Hellstein J, Vawter-Hugart H, Fotos P, Schmid J, Soll DR. Genetic similarity and phenotypic diversity of commensal and pathogenic strains of Candida albicans isolated from the oral cavity. J Clin Microbiol. 1993;31(12):3190–9. 11. Arduino PG, Porter SR. Herpes simplex virus type 1 infection: overview on relevant clinico-pathological features. J Oral Pathol Med. 2008;37(2):107–21.

6

New Oral Microbial Isolations Yuqing Li, Xian Peng, Biao Ren, Boyu Tang, Tao Gong, Zhengyi Li, and Xuedong Zhou

Abstract

More than 700 different species of bacteria, viruses, fungi, mycoplasma, and chlamydia live in the human oral cavity, collectively known as the oral microbiome. The ecological imbalance of oral microbiome can induce oral infectious diseases, such as dental caries and periodontal disease. Oral microorganisms are also closely related to tumors, diabetes, rheumatoid arthritis, cardiovascular diseases, premature delivery, and other systemic diseases, which have a significant impact on human health. So far, more than 65% of oral microbiota can be cultured in vitro. In order to better study the relationship between oral microbes and human health, several human oral microbiome collection and oral microbiome database have been established. In this chapter, the biological and pathogenic characteristics of several newly isolated oral microbes from dental plaque were introduced. Y. Li · X. Peng · B. Ren · B. Tang · T. Gong · Z. Li State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China X. Zhou (*) State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China e-mail: [email protected]

Gram staining, plate culture, colony, and scanning electron microscope (SEM) images of each microorganism were also provided. Keywords

Leuconostoc lactis · Stenotrophomonas maltophilia · Chryseobacterium indologenes · Elizabethkingia anophelis · Klebsiella pneumoniae · Campylobacter jejuni

6.1 6.1.1

Gram-Positive Bacteria Leuconostoc lactis

Leuconostoc lactis is one of the species of Leuconostoc belonging to Leuconostocaceae family [1]. It is the only Leuconostoc species that does not hydrolyze esculin. L. lactis is a Gram-positive facultative anaerobic bacteria (Fig. 6.1). L. lactis requires a rich and complex media to grow well at 25
C. They are generally ovoid cocci and often form chains (Fig. 6.2). Compared to staphylococci, L. lactis is intrinsically resistant to vancomycin and are catalasenegative. It is mainly employed in the food industry, but also reported to cause osteomyelitis at times. These bacteria are generally slimeforming. Blood brain heart infusion (BHI) media is used for L. lactis isolation (Figs. 6.3 and 6.4). Its genomic GC content is 37–40% (mol%), and the type strain is L. lactis ATCC15520.

# Zhejiang University Press 2020 X. Zhou, Y. Li (eds.), Atlas of Oral Microbiology: From Healthy Microflora to Disease, https://doi.org/10.1007/978-981-15-7899-1_6

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254 Fig. 6.1 Cells of L. lactis (Gram stain)

Fig. 6.2 Cells of L. lactis (SEM)

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Fig. 6.3 Colonies of L. lactis (BHI blood agar)

Fig. 6.4 Colonies of L. lactis (stereomicroscope)

Biological characteristics: L. lactis can be heterofermentative and produce dextran from sucrose. Its catalase is negative and arginine cannot be hydrolyzed. It cannot produce indole

and is not hemolytic. Moreover, it cannot reduce nitrate to nitrite, and the final pH of its culture in liquid glucose medium is between 4.4 and 5.0.

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Fig. 6.5 Cells of C. argentoratense (Gram stain)

Colonization characteristics: Colonies are generally less than 1.0 mm, smooth, round, and graywhite. In blood BHI media, the colonies have gray-white concentric outer ring and dull yellow pigmented ring at the center.

6.1.2

Corynebacterium argentoratense

Corynebacterium argentoratense, one of the species of Corynebacterium genus, is an irregular, Gram-positive rod-shaped bacterium [2, 3] (Figs. 6.5 and 6.6). It contains a type IV cell wall and short-chain mycolic acids. Its cell wall contains arabinose, galactose, and mesodiaminopimelic acid. The bacterium is non-lipophilic and has typical coryneform morphology. Blood BHI media was used for C. argentoratense isolation (Figs. 6.7 and 6.8). Its genomic GC content is 60–61% (mol %), and the strain type is IBS B10697 (CIP 104296). Biological characteristics: C. argentoratense is a catalase-positive, oxidase-negative, and facultative anaerobe. Nitrate is not reduced, and

tyrosine, gelatin, DNA, esculin, and urea are not degraded or hydrolyzed. Acid is produced from D-glucose and fructose but not from sucrose, maltose, lactose, galactose, D-xylose, trehalose, glycogen, or D-mannitol. Acidification from ribose occurs variably. Colonization characteristics: C. argentoratense forms creamy colonies (2 mm diameter) at 37
C, with an absence of hemolysis on blood BHI media.

6.2 6.2.1

Gram-Negative Bacteria Stenotrophomonas maltophilia

Stenotrophomonas maltophilia is an aerobic, non-fermentative, Gram-negative bacterium (Figs. 6.9 and 6.10) that is ubiquitously found in aqueous environments, soil and plants. However, it also is the third most common nosocomial pathogen with multidrug resistance [4, 5]. It was called as Pseudomonas maltophilia according to its flagellum characteristics in 1961 and Xanthomonas maltophilia in 1983 according to its nucleic acid homology and cellular fatty acid

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Fig. 6.6 Cells of C. argentoratense (SEM)

composition. In 1993, however, it was renamed as Stenotrophomonas maltophilia as Xanthomonas would imply plant pathogenicity. Blood BHI media is used for S. maltophilia isolation at 37
C (Figs. 6.11 and 6.12). S. maltophilia is a common conditional pathogen in clinic infection that is difficult to treat in immunocompromised patients. Its genomic GC content is 66.7% (mol%), and the type strain is S. maltophilia ATCC 13637. Biological characteristics: S. maltophilia can reduce nitrate to nitrite and is oxidase negative. It can hydrolyze gelatin, and it has potent lipolytic properties. It is DNase, aescin, and lysine decarboxylase positive. In the oxidative fermentation experiment, S. maltophilia ferments maltose, but acid is produced very slowly and not evidently. S. maltophilia contains beta-lactamase and can be naturally resistant to imipenem. Colonization characteristics: S. maltophilia can give out a strong odor of ammonia in the blood plate and show no hemolysis. The colonies are smooth edged, pale yellow, or grayish-white

in color, with a diameter of 0.5–1 mm and central protuberance.

6.2.2

Chryseobacterium indologenes

Chryseobacterium indologenes is a yellow pigmented, Gram-negative, filamentous (Fig. 6.13), nonmotile, rod-shaped bacteria that can be found in soil, plants, food material, water sources, and hospitals [6]. It is approximately 0.5 μm in diameter and 1.0–3.0 μm in length, with rounded ends and parallel sides (Fig. 6.14). C. indologenes is a non-fastidious and chemoorganotrophic organism that can rapidly grow at 37
C. Blood BHI media can be used for the isolation of C. indologenes (Figs. 6.15 and 6.16). Its genomic GC content is 37–39% (mol %), and the type strain is C. indologenes ATCC 29897. Biological characteristics: C. indologenes can survive in anaerobic conditions with nitrate or fumarate used as the terminal electron acceptor.

258 Fig. 6.7 Colonies of C. argentoratense (BHI blood agar)

Fig. 6.8 Colonies of C. argentoratense (stereomicroscope)

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Fig. 6.9 Cells of S. maltophilia (Gram stain)

Fig. 6.10 Cells of S. maltophilia (SEM)

C. indologenes ferments D-fructose, D-glucose, glycerol, maltose, trehalose, glycogen, and mannose to produce acid, but not lactose, L-arabinose,

ethanol, sucrose, or D-xylose. Catalase, oxidase, and phosphatase are observed in C. indologenes, as well as strong proteolytic activities and

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Fig. 6.11 Colonies of S. maltophilia (BHI blood agar)

Fig. 6.12 Colonies of S. maltophilia (stereomicroscope)

oxidation of carbohydrates. However, glycerol and trehalose are not oxidized. C. indologenes is also capable of degrading esculin, DNA, starch,

and Tween 80 and produces indole. It does not, however, produce β-galactosidase or L-phenylalanine deaminase.

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Fig. 6.13 Cells of C. indologenes (Gram stain)

Fig. 6.14 Cells of C. indologenes (SEM)

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Fig. 6.15 Colonies of C. indologenes (BHI blood agar)

Fig. 6.16 Colonies of C. indologenes (stereomicroscope)

Colonization characteristics: Colonies are circular, convex, entire, smooth with up to 2 mm diameter and aromatic odor. They are a

deep yellow in color and thinner in the central portion than in the periphery.

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Fig. 6.17 Cells of E. anophelis (Gram stain)

6.2.3

Elizabethkingia anophelis

Elizabethkingia anophelis, belonging of the Flavobacteriaceae family, is a slightly yellow-pigmented, nonmotile, non-sporeforming, Gram-negative, rod-shaped bacterium (Figs. 6.17 and 6.18). It can grow well at 30–31
C and 37
C. It is a dominant resident in the gut of the malaria vector mosquito Anopheles gambiae. Its type strain is R26(T) (¼ CCUG 60038(T) ¼ CCM 7804(T)). Blood BHI media can be used for its isolation (Figs. 6.19 and 6.20). E. anophelis has natural antibiotic resistance to several antibiotics, including β-lactam antibiotics and fluoroquinolones. However, recent clinical cases have reported that it’s developing multidrug resistance [7–9]. Biological characteristics: E. anophelis can ferment fructose, glucose, lactose, maltose, mannitol, and trehalose to produce acid and hydrogen sulfide. However, arabinose, raffinose, salicin, sucrose, and xylose cannot be metabolized by E. anophelis. It cannot make use of malonic acid and can hydrolyze aescin. It cannot reduce nitrate to nitrite. Colonization characteristics: E. anophelis forms smooth, slightly yellow-pigmented, clear

boundary, round colonies which are transparent or translucent. In blood BHI media, no zone of hemolysis is observed.

6.2.4

Klebsiella pneumoniae

Klebsiella pneumoniae is a facultative anaerobic Gram-negative bacterium. It is rod-shaped and measures 2  0.5 μm (Figs. 6.21 and 6.22). It can be single, in pairs, or in short chains. It has no spores and no flagella, but has thick capsule layer surrounding the bacterium. It can be found in surface water, sewage, and soil. As a pathogen, K. pneumoniae can cause pneumonia which can cause destructive changes to human and animal lungs, if aspirated, specifically in alveoli, resulting in bloody sputum. In recent years, Klebsiella species have become an important pathogen in nosocomial infections [12]. Its genomic GC content is about 57% (mol%), and the type strain is ATCC13883. Biological characteristics: K. pneumoniae is negative for oxidase test. It can ferment glucose, lactose, and sucrose. The result of triple sugar iron is A/A. Indole, methyl red, motility, ornithine decarboxylase, and arginine dihydrolase

264 Fig. 6.18 Cells of E. anophelis (SEM)

Fig. 6.19 Colonies of E. anophelis (BHI blood agar)

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Fig. 6.20 Colonies of E. anophelis (stereomicroscope)

Fig. 6.21 Cells of K. pneumoniae (Gram stain)

tests are negative, whereas the Voges-Proskauer, citrate utilization, nitrate reduction, urease, and lysine decarboxylase tests are positive.

Colonization characteristics: Colonies are round, convex, gray-white colored, non-hemolytic, and mucoid in blood BHI media

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Fig. 6.22 Cells of K. pneumoniae (SEM)

after 18–24 h of culturing at 37
C (Figs. 6.23 and 6.24).

6.2.5

Escherichia coli

Escherichia coli is a Gram-negative, facultative anaerobic, rod-shaped, coliform bacterium of the genus Escherichia (Figs. 6.25 and 6.26). It is commonly found in the lower intestine of warmblooded organisms (endotherms). Most E. coli strains are harmless, but some serotypes can cause serious food poisoning in their hosts and are occasionally responsible for product recalls due to food contamination. The harmless strains are part of the normal microbiota of the gut and can benefit their hosts by producing vitamin K2 and preventing colonization of the intestine with pathogenic bacteria, having a symbiotic relationship. E. coli is expelled into the environment via fecal matter. The bacterium grows massively in fresh fecal matter under aerobic conditions for 3 days, but its numbers decline slowly then after.

Biological characteristics: E. coli can thrive on a wide variety of substrates and uses mixed-acid fermentation in anaerobic conditions, producing lactate, succinate, ethanol, acetate, and carbon dioxide. In mixed-acid fermentation, many pathways produce hydrogen; its levels are required to be low, such as in the case of E. coli which colonies with hydrogen-consuming organisms, like methanogens or sulfate-reducing bacteria. Colonization characteristics: Colonies are off-white or beige in color with a shiny texture (Figs. 6.27 and 6.28).

6.2.6

Campylobacter jejuni

Campylobacter jejuni is one of the most common causes of food poisoning in Europe and the United States. The vast majority of these cases occur as isolated events, and not as a part of recognized outbreaks. A surveillance by the Foodborne Diseases Active Surveillance

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Fig. 6.23 Colonies of K. pneumoniae (BHI blood agar)

Fig. 6.24 Colonies of K. pneumoniae (stereomicroscope)

Network (FoodNet) indicated that about 14 cases are diagnosed each year for every 100,000 persons in the population. Biological characteristics: C. jejuni is a helical-shaped, non-spore-forming, Gram-negative, microaerophilic, and non-fermenting bacterium, forming motile rods with a single polar flagellum (Figs. 6.29 and 6.30). It is also

oxidase-positive and grows optimally between 37 and 42
C. When exposed to atmospheric oxygen, C. jejuni is able to change into a coccal form. This species of pathogenic bacteria is one of the most common causes of human gastroenteritis [10, 11]. Colonization characteristics: The colonies are small, mucoid, usually grayish in coloration, flat

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Fig. 6.25 Cells of E. coli (Gram stain)

Fig. 6.26 Cells of E. coli (SEM)

with irregular edges, and non-hemolytic at 24–48 h of culturing (Figs. 6.31 and 6.32). An alternate colonial morphology that appears to be strain related consists of round colonies 1–2 mm

in diameter that are convex, and glistening. Colonies tend to spread or swarm, especially when initially isolated from fresh clinical specimens.

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Fig. 6.27 Colonies of E. coli (BHI agar)

Fig. 6.28 Colonies of E. coli (stereomicroscope)

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270 Fig. 6.29 Cells of C. jejuni (Gram stain)

Fig. 6.30 Cells of C. jejuni (SEM)

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Fig. 6.31 Colonies of C. jejuni (BHI blood agar)

Fig. 6.32 Colonies of C. jejuni (stereomicroscope)

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Fig. 6.33 Cells of V. atypica (Gram stain)

6.2.7

Veillonella atypica

Veillonella atypica is a lactate fermenting bacteria (Figs. 6.33 and 6.34). It is a normal bacterium of the intestines and oral mucosa in mammals. In humans, they have been known to cause osteomyelitis and endocarditis. The relative abundance of V. atypica found in the gut of endurance athletes is associated with increased treadmill run-time performance. This effect was found to be due to the propionate metabolite produced by the organism from lactic acid. Biological characteristics: Lactate is fermented by V. atypica to propionate and acetate by the methylmalonyl-CoA pathway, and a small amount of ATP is produced in this process due to high substrate affinity. Colonization characteristics: Colonies are round, convex, white, non-hemolytic, and mucoid in blood BHI media after 18–24 h of culturing at 37
C (Figs. 6.35 and 6.36).

6.3 6.3.1

Fungi Candida tropicalis

Candida tropicalis is an asexual, diploid, opportunistic pathogen, with a shape ranging from

round to oval and approximately 2–6 μm in size (Figs. 6.37 and 6.38). It is more likely to be found in tropical climate, where the temperature and humidity enhance its adaptability. It is a widespread species of yeast that colonizes in food, plants, gastrointestinal tract, and the mucocutaneous membranes of humans causing a variety of diseases. C. tropicalis has a strong vitality and is highly infectious. It is responsible for approximately half of the beyond-surface Candida infections with the second most virulence observed in Candida species. It is more often associated with deep fungal infections than normal mucosa and can be easily identified using phenotypic and molecular methods. Antifungal agents can be used to treat the infections caused by C. tropicalis. The size of the diploid genome is approximately 14 Mb, with 33.2% GC content. Biological characteristics: Candida tropicalis reproduces asexually by the production of blastoconidia through budding. It can survive in high salt concentration and can develop fungal persistence in saline environments. The optimum pH value for the growth of C. tropicalis is 5.5, and the optimum temperature ranges from 25 to 35
C. There are different media on which C. tropicalis can grow effectively. The most common medium used is the Sabouraud agar and Corn Meal Agar.

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Fig. 6.34 Cells of V. atypica (SEM)

Fig. 6.35 Colonies of V. atypica (BHI agar)

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Fig. 6.36 Colonies of V. atypica (stereomicroscope)

Fig. 6.37 Cells of C. tropicalis (Gram stain)

Colonization characteristics: Colony of C. tropicalis are white, smooth, and butyrous with a fringed border (Figs. 6.39 and 6.40). When cultured on CHROMagar™ Candida at

35
C for 48 h, approximately 1.5-mm-sized iron blue colonies could be found according to the enzyme substrate method.

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Fig. 6.38 Cells of C. tropicalis (SEM)

Fig. 6.39 Colonies of C. tropicalis

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Fig. 6.40 Colonies of C. tropicalis (stereomicroscope)

6.3.2

Candida glabrata

Candida glabrata, formerly known as Torulopsis glabrata, is a species of opportunistic pathogens (Figs. 6.41 and 6.42). It is often considered as the second most common cause of candidiasis and also a commensal of human mucosal tissues. Unlike C. albicans, C. glabrata is a species of haploid yeast with 13 chromosomes, whose mating types are common. The genome of C. glabrata undergoes frequent rearrangements, contributing to high resistance to antifungal agents such as azoles. The identification of C. glabrata usually requires culturing for several days. Thorough treatment is difficult to accomplish as the infection spreads rapidly and drug resistance. The median total length of the genome size is approximately 12 Mb, with 38.6% median GC content. Biological characteristics: C. glabrata do not form hyphae. The culturing methods and

conditions required are same as most of the Candida species, i.e., on Sabouraud medium and Corn Meal Agar, except for the slow growth. It should be noted that C. glabrata ferments and assimilates only glucose and trehalose, which could be used for its identification. The frequent genome rearrangements contribute to its fitness to thrive under stressful conditions, helping the fungus to tightly adhere and develop a resistance to the drugs. C. glabrata has innate tolerance to most azoles; however, it is vulnerable to polyenes. Its resistance to echinocandin is gradually increasing, making its treatment more difficult. Colonization characteristics: The growth on Sabouraud medium is slow. Colony of C. glabrata are small, gray, and smooth after 2–3 days of incubation (Figs. 6.43 and 6.44). Pseudohyphae and chlamydospore cannot be formed on 1% Tween80-Corn Meat Agar Medium. However, when cultured on

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Fig. 6.41 Cells of C. glabrata (Gram stain)

Fig. 6.42 Cells of C. glabrata (SEM)

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Fig. 6.43 Colonies of C. glabrata

Fig. 6.44 Colonies of C. glabrata (stereomicroscope)

CHROMagar™ Candida at 35
C for 48 h, approximately 2-mm-sized lavender colonies could be found.

6.3.3

Candida parapsilosis

Candida parapsilosis is a species of clinically important yeast-like fungus, previously known

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Fig. 6.45 Cells of C. parapsilosis (Gram stain)

Fig. 6.46 Cells of C. parapsilosis (SEM)

as Monilia parapsilosis, and was considered nonpathogenic (Figs. 6.45 and 6.46). It is considered an important emerging nosocomial pathogen, which often infects patients via catheters and internal medical devices in the hospitals. It is now the second most common non-C. albicans

species of Candida from blood cultures in Asia, Europe, Canada, and Latin America. It is most commonly isolated from the human skin. C. parapsilosis is not an obligate human pathogen and is widely distributed in the environment. Immunocompromised individuals and those

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Fig. 6.47 Colonies of C. parapsilosis

undergoing surgery of the gastrointestinal tract can contribute to C. parapsilosis colonization. Treatment of invasive candidiasis includes the removal of foreign bodies and the administration of antifungal agents. The median total length of genome size of C. parapsilosis is approximately 13 Mb, with 38.6% median GC content. Biological characteristics: The shape of C. parapsilosis cell is oval, round, or cylindrical under microscope. It can exist in yeast phase or pseudohyphal form, meaning not forming a true hyphae. It can produce active molecules to exert cytotoxic effects on the other organisms. As it is an unobligated human pathogen, C. parapsilosis is frequently encountered in nature than the other species of Candida and is usually transmitted by external sources. Colonization characteristics: Colonies are generally white, creamy, and shiny on Sabouraud medium (Figs. 6.47 and 6.48). C. parapsilosis is smooth or cratered in yeast form, while wrinkled or concentric in pseudohyphal form. The colonies may appear white or lavender when cultured on CHROMagar™ Candida at 35
C for 48 h.

6.3.4

Candida krusei

Candida krusei is species of opportunistic pathogen, but also yeast used in food processing (Figs. 6.49 and 6.50). Candida krusei is also referred to as Pichia kudriavzevii, Issatchenkia orientalis, and Candida glycerinogenes known as an industrial yeast. Candida krusei is an emerging fungal nosocomial pathogen with higher mortality than C. albicans, result in patients that obtain this fungus, got the lowest 90-day survival period among all Candida species, as mainly found in the immunocompromised and patients with hematological malignancies. The identification of C. krusei becomes easy with its typical colony characteristics on Sabouraud medium and 2.5–5.5  7.5–21.5 μm “long-grain rice” appearance on microscopy. Voriconazole, polyenes, and echinococcins could be administrated in the treatment of C. krusei, except fluconazole, to which the fungus has natural resistance. The median total length of genome size is approximately 11 Mb, with 38.33% median GC content.

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Fig. 6.48 Colonies of C. parapsilosis (stereomicroscope)

Fig. 6.49 Cells of C. krusei (Gram stain)

Biological characteristics: C. krusei could be cultured on Sabouraud agar and Corn Meal Agar with the maximum temperature of 43–45
C. Interestingly, C. krusei is the only Candida that can grow in vitamin-free media. Grown in media containing lactose, C. krusei could metabolize

complex varieties of fatty acids and also produce a number of short-chain carboxylic acids when cultured in saliva containing glucose. Colonization characteristics: In contrast to the other Candida species, C. krusei demonstrate significantly different spreading colonies with a

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Fig. 6.50 Cells of C. krusei (SEM)

matte or a rough whitish yellow surface on Sabouraud dextrose agar (Figs. 6.51 and 6.52). Cultured on CHROMagar™ Candida at 35
C for 48 h, approximately 4–5 mm, larger pink colonies could be recognized as C. krusei.

6.3.5

Aspergillus fumigatus

Aspergillus fumigatus is a common fungus of Aspergillus species that is widespread in the nature and playing an important role in the carbon and nitrogen recycling (Figs. 6.53 and 6.54). It is widely distributed in areas with distinct climates and environment; however, it demonstrates low genetic variation and lacks population genetic differentiation. Aspergillus fumigatus is also a species of opportunistic pathogens, causing a range of diseases termed as aspergillosis, especially in lungs of immunocompromised individuals. With its increasing prevalence, invasive aspergillosis has overtaken candidiasis as the most frequent fungal infection in the world. Therefore, a study

of the A. fumigatus has become a priority. It is estimated that A. fumigatus could be responsible for over 600,000 deaths annually, with a mortality rate of 25–90%. The life cycle of A. fumigatus consists of two phases: a hyphal growth phase and a sporulation phase. The colonies of this fungus are produced from conidiophores, and the spores are ubiquitous in the atmosphere. Once the immune functions are impaired, the conidia emerge from dormancy and undergo a morphological switch to hyphae by germinating in the warm, moist, nutrient-rich environment in the human body. This in turn could result in intravascular thrombosis and localized tissue infarction. Current treatment consists of a class of drugs, known as azoles, which have good antifungal activity. Aspergillus fumigatus has a stable haploid genome of 29.4 million base pairs, and median GC content is 49.5%. Biological characteristics: Aspergillus fumigatus is a highly aerobic organism, capable of growing at 37–50
C, with its conidia surviving even at 70
C. The spores are ubiquitous in the atmosphere. Czapek-Dox medium is a growth

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Fig. 6.51 Colonies of C. krusei

Fig. 6.52 Colonies of C. krusei (stereomicroscope)

medium for propagating fungi and other organisms, which is conducive for the growth of A. fumigatus. When this fungus is incubated on Sabouraud agar for 24–72 h, the spores are formed with a change of color. Conidia have a

short stalk, and one segment bulges to form a parietal capsule, which is like a green-colored flask under microscope. In 80% of capsule, the spores are arranged in a beaded manner in radial pattern, attached to the stalk.

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Fig. 6.54 Cells of A. fumigatus (SEM)

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Fig. 6.55 Colonies of A. fumigatus

Fig. 6.56 Colonies of A. fumigatus (stereomicroscope)

Colonization characteristics: On Sabouraud agar, colonies are white, flocculent, and alike, initially (Figs. 6.55 and 6.56). However, once

the spores are formed, the colonies are floured, with color changing from pale gray, green, dark green, and then smoky green to black.

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References 1. Moon JS, Choi HS, Shin SY, Noh SJ, Jeon CO, Han NS. Genome sequence analysis of potential probiotic strain Leuconostoc lactis EFEL005 isolated from kimchi. J Microbiol. 2015;53(5):337–42. 2. Fernández-Natal I, Sáez-Nieto JA, RodríguezLázaro D, Valdezate-Ramos S, Parras-Padilla T, Medina MJ, Rodríguez-Pollán RH, Blom J, Tauch A, Soriano F. Phenotypic, molecular characterization, antimicrobial susceptibility and draft genome sequence of Corynebacterium argentoratense strains isolated from clinical samples. New Microbes New Infect. 2016;10:116–21. 3. Bomholt C, Glaub A, Gravermann K, Albersmeier A, Brinkrolf K, Rückert C, Tauch A. Whole-genome sequence of the clinical strain Corynebacterium argentoratense DSM 44202, isolated from a human throat specimen. Genome Announc. 2013;1(5): e00793-13. 4. Brooke JS. Stenotrophomonas maltophilia: an emerging global opportunistic pathogen. Clin Microbiol Rev. 2012;25(1):2–41. 5. An SQ, Berg G. Stenotrophomonas maltophilia. Trends Microbiol. 2018;26(7):637–8.

Y. Li et al. 6. Imataki O, Uemura M. Chryseobacterium indologenes, a possible emergent organism resistant to carbapenem antimicrobials after stem cell transplantation. Clin Case Rep. 2016;5(1):22–5. 7. Janda JM, Lopez DL. Mini review: New pathogen profiles: Elizabethkingia anophelis. Diagn Microbiol Infect Dis. 2017;88(2):201–5. 8. Chew KL, Cheng B, Lin RTP, Teo JWP. Elizabethkingia anophelis Is the Dominant Elizabethkingia Species Found in Blood Cultures in Singapore. J Clin Microbiol. 2018;56(3):e01445-17. 9. Raygoza Garay JA, Hughes GL, Koundal V, Rasgon JL, Mwangi MM. Genome Sequence of Elizabethkingia anophelis Strain EaAs1, Isolated from the Asian Malaria Mosquito Anopheles stephensi. Genome Announc. 2016;4(2):e00084-16. 10. Young KT, Davis LM, Dirita VJ. Campylobacter jejuni: molecular biology and pathogenesis. Nat Rev Microbiol. 2007;5(9):665–79. 11. Davis L, DiRita V. Growth and laboratory maintenance of Campylobacter jejuni. Curr Protoc Microbiol. 2008., Chapter 8:Unit;10:8A.1.1–7. 12. Chen L, Mathema B, Chavda KD, DeLeo FR, Bonomo RA, Kreiswirth BN. Carbapenemase-producing Klebsiella pneumoniae: molecular and genetic decoding. Trends Microbiol. 2014;22(12):686–96.

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The Oral Microbiome Bank of China Xian Peng, Xuedong Zhou, Xin Xu, Yuqing Li, Yan Li, Jiyao Li, Xiaoquan Su, Shi Huang, Jian Xu, and Ga Liao

Abstract

X. Peng · Y. Li · Y. Li · G. Liao (*) State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China X. Zhou (*) State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China e-mail: [email protected] X. Xu State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Department of Operative Dentistry and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China J. Li State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China X. Su · S. Huang · J. Xu Single-Cell Center, CAS Key Laboratory of Biofuels and Shandong Key Laboratory of Energy Genetics, Qingdao Institute of BioEnergy and Bioprocess Technology, Chinese Academy of Sciences, Qingdao, Shandong, China

The Human Microbiome Project (HMP) promoted further understanding of human oral microbes. However, research on the human oral microbiota has not made as much progress as research on the gut microbiota. Currently, the causal relationship between the oral microbiota and oral diseases remains unclear, and little is known about the link between the oral microbiota and human systemic diseases. To further understand the contribution of the oral microbiota in oral diseases and systemic diseases, a Human Oral Microbiome Database (HOMD) was established in the USA. The HOMD includes 619 taxa in 13 phyla, and most of the microorganisms are from American populations. Due to individual differences in the microbiome, the HOMD does not reflect the Chinese oral microbial status. Herein, we established a new oral microbiome database— the Oral Microbiome Bank of China (OMBC, http://www.sklod.org/ombc). Currently, the OMBC includes information on 289 bacterial strains and 720 clinical samples from the Chinese population, along with lab and clinical information. The OMBC is the first curated description of a Chinese-associated microbiome; it provides tools for use in investigating the role of the oral microbiome in health and diseases and will give the community abundant data and strain information for future oral microbial studies.

# Zhejiang University Press 2020 X. Zhou, Y. Li (eds.), Atlas of Oral Microbiology: From Healthy Microflora to Disease, https://doi.org/10.1007/978-981-15-7899-1_7

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Keywords

Human Microbiome Project · Oral microbiome · Oral diseases · Systemic diseases · 16s rRNA sequence

7.1

Introduction

With the development of microbiome research in recent years, studies have found that the human microbiome is closely related to systemic diseases [1–4]. At the end of 2007, the NIH launched the Human Microbiome Project (HMP), which aimed to resolve microorganisms by mapping the genomes of five major parts of the human body (buccal, nasal, vaginal, gut and skin microbiomes) [5, 6]. The oral microbial community and oral and systemic health are closely related; they can induce not only dental caries, apical periodontal disease, periodontal disease, pericoronitis, oral mucosal disease and other oral diseases but also many systemic diseases [7–12]. Based on the clinical features of oral disease diagnosis and treatment, the oral microbiome has more advantages than other parts of the body in terms of convenience, ease of operation and number of patients. Currently, the Human Oral Microbiome Database (HOMD, http://www. homd.org) includes 619 taxa in 13 phyla, 65% of which can be cultured in the laboratory [13, 14]. The human oral cavity is a complex ecological niche, containing both exfoliated surfaces (mucosa) and non-shedding solid surfaces (tooth surfaces), and contains fluid saliva. Therefore, the oral microflora colonized on different surfaces have significant spatial specificity [15, 16]. In the process of individual development and growth, the oral microbiome changes dynamically; as age increases and dentition changes, physiological changes occur in the oral microbiome, and the composition of microorganisms in different age groups has large specificity [2, 12, 17]. Keijser et al. found that oral microorganisms covered 318 genera of 22 phyla, 5600 and 10,000 of which were

colonized in saliva and plaque, respectively [15, 18, 19]. As early as 1891, Miller proposed the theory that oral lesion infection may cause a variety of systemic diseases due to oral microorganisms entering other parts of the body through oral infection [20]. In recent years, with in-depth study of the microbiome, the correlation between the oral microbiome and various systemic diseases has gradually been confirmed, including digestive system diseases, cardiovascular diseases, tumours, premature birth, diabetes and rheumatoid arthritis [9, 21–25]. It has been reported that an increased intracellular C-reactive protein levels caused by bacterial infections in the mouth have an important correlation with the development of atherosclerotic vascular disease [26, 27]. C-reactive protein levels in saliva can predict the occurrence of acute myocardial infarction, which will benefit the early diagnosis and control of disease [28, 29]. Lactobacilli and streptococci, which are closely related to dental caries, are involved in the pathogenesis of infective endocarditis [30, 31]. Among them, the early colonization of oral biofilm—Streptococcus sanguinis in patients with the highest detection rate of the endocardium—and endocarditis are closely related to the occurrence and development of infective endocarditis [32, 33]. Currently, the pathogenesis occurs with the formation of thrombus-like processes on the surface of normal vascular endothelial cells, promoting bacterial adhesion, which in turn leads to infection [34, 35]. Researchers found that the expression of Streptococcus gordonii virulence-associated factor was significantly increased in the process of endocardial infection, which may play a vital role in the endocarditis model [32, 36]. In patients with severe periodontal infection, the oral microbiome interacts with the immune system in a complex manner, producing persistent chronic inflammation, and some oral microbes can also enter the bloodstream [24, 37–39]. A large number of studies have confirmed that diabetes and chronic inflammation in the human body related to periodontitis can lead to systemic inflammatory cytokines, such as cytokines TNF-α, IL-1β and IL-6, and can

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increase the body’s oxidative stress, affecting insulin sensitivity and glucose metabolism [38, 40–43]. Human chronic inflammation caused by periodontal microbial infections is likely to be one of the mechanisms that contributes to the development of diabetes [15, 44–46]. Moreover, a recent study showed that periodontal pathogens involved in the occurrence of rheumatoid arthritis (RA) also exhibit a certain correlation [47]. Actinobacillus actinomycetemcomitans can induce the dysregulation of PAD activity in host neutrophils by the secretion of toxin P or toxin A (LtxA), leading to a high degree of citrullination of proteins in neutrophils and the formation of autoantigens [48]. LtxA also alters neutrophil morphology, mimicking neutrophil extracellular trapping nets and causing neutrophil lysis, eventually releasing massive citrullinated antigens [43, 49]. Unlike Porphyromonas gingivalis, which synthesizes bacterial PAD, Actinobacillus actinomycetemcomitans induces autoantigen formation by inducing endogenous PAD activity, leading to the formation of ACPA and rheumatoid factor and the development of RA [30, 50]. In addition, the oral microbiome may also serve as a “fingerprint” for the development and treatment of rheumatoid arthritis [51]. A recent study found that there is significant ecological imbalance in the oral microbiome of patients with rheumatoid arthritis, which can be recovered by the treatment of rheumatoid arthritis. Based on a metagenomic association analysis between the oral and gut microflora, a diagnostic classification model of the human population in the diagnosis of healthy people and rheumatoid arthritis patients was built with a diagnostic accuracy of nearly 100%, suggesting that the oral microorganism group is linked to the occurrence, development and prognosis of RA [51]. The human oral microbiome constantly interacts and evolves with the human body, and the composition of the human oral microbiome varies greatly in different ethnicities and regions; studying the oral microbiome in the Chinese population is indispensable. Given the importance of the oral microbiome and with the aims to further study the Chinese oral microbiology group and serve oral microbiology researchers in China, the

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first goal of this study was to develop a provisional taxonomic scheme for unnamed Chinese oral bacterial isolates and phylotypes and provide this information in an online publicly available database, namely, the Chinese Oral Microbiome Bank of China (OMBC) (http://www.sklod.org/ ombc). The second goal was to analyse the 16S rRNA gene oral clone sequences to determine the number of clones observed for each Chinese oral taxon and to identify additional taxa that were not included in the initial setup of the OMBC.

7.2 7.2.1

Results Overview of the Online Database

The Chinese Oral Microbiome Database has two main parts currently; one component is bacterial strain information in the Chinese population, which is based on the 16S rRNA gene sequences that were used to define individual Chinese oral taxa and create the taxonomic structures in the database. The other component is clinical sample information, which was obtained through secondgeneration sequencing and multi-omics analysis to collect biological information on samples for correlation analysis. Currently, information on a total of 289 bacterial strains was stored in our online database, and we also collected multidimensional characteristics of the bacterial strains by identifying their biochemistry and molecular properties. At the time of writing, the online database contained 60 (20.76%) 16S rRNAsequenced bacterial strains, 102 (35.29%) of which had biochemical evidence and 117 (40.48%) of which had biochemical descriptions. A detailed exhibition of the phylogenies of the 289 bacteria can be seen in the phylogenetic tree (Fig. 7.1). The evolutionary tree was inferred using the Neighbour-Joining method [52]. The bootstrap consensus tree inferred from 1000 replicates was assumed to represent the evolutionary history of the analysed taxa [53]. Branches corresponding to the partitions reproduced in less than 50% bootstrap

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Fig. 7.1 Evolutionary relationships of cultured bacteria. All positions containing gaps and missing data were eliminated. There was a total of 1255 positions in the

final dataset. Evolutionary analyses were conducted in MEGA7

replicates are collapsed. The evolutionary distances were computed using the Kimura two-parameter method [54] and are in the units of number of base substitutions per site. The name of each bacteria is followed by its designated OMBC accession number. An overview of the phylogenetic distribution of these 289 bacteria is shown in Table 7.1. Three phyla, Firmicutes, Proteobacteria, and Actinobacteria,

contain ten families, Micrococcaceae, Actinomycetaceae, Neisseriaceae, Enterobacteriaceae, Pseudomonadaceae, Moraxellaceae, Staphylococcaceae, Lactobacillaceae, Streptococcaceae and Veillonellaceae. It is remarkable that all the 289 bacteria have been cultured and phenotypically analysed, including their colony colonial morphology, Gram staining and image recording through a scanning electron microscope,

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Table 7.1 Phylogenetic distribution of 289 bacteria in OMBC Phylum/number NULL Actinobacteria

195 22

Class/number NULL Actinobacteridae

Proteobacteria

12

Betaproteobacteria Gammaproteobacteria

Firmicutes

60

Bacilli

Clostridia

195 22 5 7

57

3

Order/number NULL Actinomycetales Neisseriales Enterobacteriales Pseudomonadales Bacillales Lactobacillales Clostridiales

195 22 5 4 3 5 52 3

Family/number NULL Micrococcaceae Actinomycetaceae Neisseriaceae Enterobacteriaceae Pseudomonadaceae Moraxellaceae Staphylococcaceae Lactobacillaceae Streptococcaceae Veillonellaceae

195 3 19 5 4 1 2 5 2 50 3

transmission electron microscope and laser confocal microscope (Fig. 7.2). Furthermore, our database supports views, queries and basic local alignment search tool (BLAST), and free downloads of the information on bacterial strains are available. New services, such as a

comprehensive analysis system and bacterial strain application system, are under development.

Fig. 7.2 Phenotypic records of a cultured microorganism, a Streptococcus mutans strain (COCC139), are shown. (a) Separated microbe on a Mitis Salivarius (MS) Agar plate. (b) Different colony forms on a blood agar plate. (c) Gram

staining. (d) Scanning electron microscopy. (e) Transmission electron microscopy. (f) Observation of extracellular polysaccharides (red) and bacterial cells (green) by confocal laser scanning microscopy

7.2.1.1 Web-Accessible Functions The upper navigation menu provides entries for each of the functions of the database. Registration

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Fig. 7.3 Description of the functions of the database. (a) Layout of the OMBC homepage; (b) layout of the query and view database page; (c & d) BLAST function and result page

and login functions are located at the upper right. A quick start of the database view and database query is located in the middle of the page, following the overall statistics of the datasets, which is updated real time (Fig. 7.3a). The layout of the view database page contains a search module and list of bacteria by taxa ID. Bacterial information can be searched using a keyword, and the search results can be listed in ascending or descending order by genus, family or order. The results are listed by four main variables, including unique taxa ID, separate no., bacteria name and disease association status. Clicking on the taxa ID or separate ID will lead to the detailed information page of a particular bacterial strain. Clicking each of the keywords (underlined phrase) of the bacterial strain name and health association status column within the result list can trigger a new set of results in the same category with the same keyword. A very convenient function is to determine bacterial strains that have the same characteristics

(Fig. 7.3b). Clicking on the bacteria library ID or separate ID leads to the detailed information page of a particular bacterial strain, and this page displays the most important characteristics of the bacterial strain, which includes the following fields: Bacteria_Library_Id/Separate_Id: The official library ID and original separate ID of the isolated microorganisms Bacteria_Name: The name of the isolated microorganisms in both Latin and Chinese Description: Description of the physiological characteristics of the isolated microorganisms Kingdom/Phylum/Class/Order/Family/Genus: The official biological classification of the isolated microorganisms Separate_Method/Separate_Source: The separate method and source of the isolated microorganisms Identify_Method: The method used for the identification of the isolated microorganisms

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Relativity_Healthy: The health status of the individuals from whom the isolated microorganisms were isolated from Hemolysis/Gram_Stain/H2O2_Enzyme/Oxidase: The metabolic abilities of the isolated microorganisms 16S_Sequence/Molecule_Confidence: The sequence of 16S rRNA and the identity of the 16S rRNA sequence

7.3

Discussion

Currently, aside from traditional microbiome research techniques, many new high-throughput analytical techniques have been developed and adopted in modern microbiome metagenomic studies. We now have a completely new understanding of oral microbes from analysing the characteristics of the oral microbiome samples of twins [46, 55, 56], newborns [57, 58], infants [59, 60] and children [61–64], adolescents [65, 66], adults [56, 67–69] and the elderly [70– 72]. The new concept of the “oral core microbiome” suggests that this phenomenon should be personalized in accordance with the ecological characteristics of the oral environment in certain populations at different ages or grades of disease [15, 16, 39, 73]. By monitoring the dynamic changes in the structure and function of microorganisms, the relationship between oral bacterial flora and these diseases, such as periodontal disease [22, 74, 75] and lichen planus [76, 77], was revealed. By studying the oral microbiome of patients with systemic diseases, the role of the oral microbiome in the occurrence and development of systemic diseases (leukemia [42, 78], head and neck cancer [79–81], HIV [82, 83], diabetes [42, 84], etc.) was clarified. Chinese researchers also proposed that the microbial indices of caries (MiC), the microbial indices of gingivitis (MiG) and the relative microbial recovery indices (RMRI) were used to evaluate gingival healthcare programmes based on the distribution pattern of bacteria in different sites within the oral cavity. The potential role of oral microbes in the

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diagnosis and prognosis of systemic diseases was demonstrated through applications in studies on rheumatoid arthritis [50, 85]. The oral cavity is an interactive environment. Oral diseases, such as oral lichen planus and oral cavity cancer, are closely related to oral microorganisms. Therefore, it is of great significance for multifactor studies on oral diseases to construct a comprehensive oral microbial database. By combining the clinical data of oral microbial communities and the microbiological metagenomics data of healthy and Chinese patients from multiple regions, ethnicities and ages, the OMBC was established, and clinical samples were collected, isolated and identified. The OMBC is the first publicly available human oral microbiome database for the Chinese population. Moreover, the OMBC has several key features. First, this well-organized database was built according to industrial standards for maximum expansion and migration capacity, and regulations regarding microbial data management were established. Second, the OMBC contains all kinds of metadata, such as 16S rRNA sequencing data and key property information on the microbiome, and can compare bacterial 16S rRNA sequences and predict oral microbial classifications and related clinical information based on clinical sample gene sequences. Third, the taxa ID that we created can be used as a unique identifier of the Chinese oral microbiome to facilitate connections and communication among different studies. Fourth, all data can be filtered and sorted in many precise methods to maximize query efficiency. Lastly, all of the data can be downloaded freely via our website. However, there are still many limitations of our database. The current quantity of the microbiome is to be improved as more samples and data are collected continuously. Additional registries will make this database more useful for future multicentre studies and will more accurately reflect the overall distribution and evolution trends of the microbiome of the Chinese population. In addition, we also plan to expand our data dimension by adding more detailed data and external data in addition to the current datasets. In the meantime, we will improve data quality.

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To improve the visibility and usability of the OMBC, we are working to carry out extensive big data studies with the database to obtain more profound insights into the oral microbiome, such as interaction networks among bacteria. It is also our mission and responsibility to build this database into a national platform as an important component of the world microbiome field and to benefit global microbiome investigators. The characteristics and commonality of microbiomerelated phenomena are receiving more and more attention. Understanding microbiomes closely related to humans helps us better understand human beings and comprehensive studies on the microbiome help with the prevention, diagnosis and treatment of many major human diseases in modern precision medicine.

7.4 7.4.1

Materials and Methods Collection and Transport of Samples

Collection of saliva samples: Unstimulated salivary samples were collected as described previously, which was followed by a gentle rinse with warm water to remove the food residue. In addition, 0.5–1.0 mL of naturally secreted saliva was collected, or saliva samples in the oral cavity were absorbed directly by the sterile pipette tips of a micro-concentrator. Collection of plaque samples: Plaque samples were collected in different ways depending on different clinical requirements and purposes. Plaque indicators were used to display plaque for some collections. Before sample collection, subjects gargled with warm water to remove food residue in their mouths. Then, sterile gauze or a yarn ball was used to isolate saliva and collect plaque samples or decayed materials. A sterile probe was used in the collection of plaque in the occlusal surface fissure. Adjacent surface plaque samples were collected with both a sterile probe and dental floss or with a fine wire used in orthodontics. A sterile curette was used in the collection of root surface plaque samples. Plaque

samples on the gum or gingival margin were collected by a spoon scaler. Collection of other infection samples: Infection samples were generally sampled with sterile cotton swabs. A purulent fluid sample of the periodontal abscess was collected with sterile syringes. The tissue pieces in the alveolar socket were generally collected as samples of dry sockets developed after tooth extraction. Collection of oral mucosal diseases samples: White membrane materials were collected with curettes or cotton swabs. To collect quantitative samples, filter papers with a particular area are used. Sample delivery: Among oral clinical specimens, the detection of specimens of fungus, aerobic bacteria and general facultative anaerobic bacteria was cotton swab-sampled and sent in sterile tubes directly. For the detection of most anaerobic bacteria or microaerophilic bacteria, samples were sent to the laboratory as soon as possible in an anaerobic way. In addition, a pus or saliva sample was inserted directly into the needlepoint sterile rubber stopper of a syringe needle tube for transport; most of the samples were put in the prereduction of anaerobic culture media and transported to the lab after immediate acquisition in order to reduce the death of bacteria that were sensitive to oxygen in the carrying process. Vaccination beside the chair and anaerobic delivery were used to improve the detection rate of obligate anaerobic bacteria. For clinical specimens that could not be inspected in a timely manner or delivered over a long distance, anaerobic bags (commercially available) or prereductions of liquid spiral tubules stamped with liquid paraffin were used for delivery.

7.4.2

Dispersion and Dilution of Samples

Oral clinical infection is generally a mixed infection of many bacteria, with mixed species and varying quantities of bacteria in a concentrated plaque mass. Thus, an oral clinical specimen is usually required for inoculation after

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decentralized processing and dilution to achieve a single colony of pure culture. Sample dispersion: Generally, two methods, named spiral vortex oscillation and ultrasonic dispersing, are adopted. Sample dilution: Oral clinical samples are mixed bacterial infection samples, with mixes in the number and variety of bacteria. Therefore, proper concentrations of diluent dilution are required before vaccination to obtain a single colony after the sample dispersion process. The transporting fluid can be used as a diluent, and a phosphate buffer with a pH value of 7.2 is also available. Tenfold dilution series are usually used. Under an aseptic operating status, a specimen liquid of 0.1 mL is added to 0.9 mL of diluent, and 0.1 mL of mixture after the diluent is thoroughly incorporated (10 1) is added to a tube containing 0.9 mL of diluent for blending. According to the above method, tenfold dilution series are achieved. Different samples have different diluted concentrations, such as a saliva sample dilution degree of 10 4–10 6, a gingiva groove plaque dilution of 10 1–10 2 and a plaque collection dilution of 10 3–10 5.

7.4.3

Inoculation and Incubation of Samples

In addition, to select the appropriate medium before inoculation, vaccination dilution degrees, inoculation method and incubation environment and times must be established according to the specimen type, purpose and microbial species. Selection of the medium: The basic media commonly used for oral bacteria include cardiocerebral immersion (BHI) medium, pancreatic enzyme-hydrolysed soy agar (TSA) and TPY agar medium. These media can be used to cultivate most bacteria in oral samples. Approximately 5% of fibre blood serum (or 5% serum) and chlorinated haemoglobin and vitamin K1 were supplied to the culture medium for some obligate anaerobic Gram-negative bacterium. To

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train general aerobic and facultative anaerobic bacteria, ordinary medium containing blood agar (BA) was used. Inoculation of samples: The spread method, drop method and spiral vaccination method were adopted for oral clinical bacteriology samples, which makes the appropriate dilution degrees of the specimen solution and quantitative inoculation on the agar plate. Incubation of samples: For a medium that has received clinical samples, its incubation conditions are determined according to the requirements of cultivation, including atmospheric conditions, temperature and time. Oral clinical specimens, such as an infected root canal, pericoronitis infection after tooth extraction and samples under the gums of periodontitis plaque, whose main characteristics are mixed bacterial infection, are generally involved different atmospheric conditions with a variety of microorganisms with their respective characteristics. An anaerobic culture containing 80% N2, 10% CO2 and 10% H2 at atmospheric conditions with a temperature of 37  C and a time of 48–72 h was used to grow the bacteria. Some bacteria in the mouth, such as Treponema, were trained in the anaerobic environment for approximately 1 week. Common anaerobic incubation devices include an anaerobic glove box, anaerobic incubation and anaerobic bag.

7.4.4

Smear and Stain

Smear test and slide stain are basic techniques for the identification of microbes, and they are primarily used for morphological observation. A combination of smear and stain tests is widely used in oral microbial research for differentiating spirochete, bacteria, fungi and protozoa and to identify specific cellular structures, including spores, capsules and flagella. Therefore, we used a direct smear test and stained smear test under a microscope for cellular morphological examination, including Gram staining and Congo red staining.

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Growth Characteristics and Identification

Phenotypic organism identification is the most basic and important part of microbiology. The classical method for bacterial identification is to observe the phenotypic characteristics on a foundation of pure bacterial culture, including colony characteristics (size, colour, shape, etc.), cell characteristics (size, shape, arrangement and dying), special structure (with or without spores, capsule and flagellum), culture characteristics (sensitivity to oxygen, optimum growth temperature and pH, requirements for nutrients and growth factors, etc.) and metabolites. Bergey’s Manual of Systematic Bacteriology, which is an authoritative reference book for bacterial isolation, is the reference for the growth characteristics of bacteria. Bacterial colony morphology includes size, colour, shape, growth patterns and characteristics. Haemolytic reaction is one of the basic characteristics of bacterial identification. Some bacteria can produce a haemolytic reaction, whereas some produce pigmentation; this test has been used for differentiation; some bacteria produce gas, and some bacteria exhibit their own growth patterns, such as the migration of Proteus growth.

7.4.6

Biochemical Tests

Biochemical tests are an important microbial identification method. Routinely, biochemical tests include the carbohydrate fermentation test, methyl red test, citric acid utilization test and hydrogen sulphide production test. The microbiochemical test reaction plate we used included 30 biochemical matrixes and relevant biochemical test indicators, phosphate-buffered saline (PBS), a bacterial turbidity standard tube and eight identification series and was used in a VITEK-2 COMPACT.

7.4.7

Molecular Method

The 16S rRNA gene was used as the standard for the classification and identification of microbes because it is present in most microbes and shows proper changes. Type strains of 16S rRNA gene sequences for most bacteria and archaea are available in public databases (GenBank).

7.4.8

DNA Extraction and Sequencing

Total DNA was extracted from collected samples from each respective host. The barcoded 16S rRNA amplicons [86] (V1–V3 hypervariable region) of all samples were sequenced on using a Roche 454 FLX Titanium. Pyrosequencing data were analysed using scripts from MOTHUR [87], QIIME [88] and custom R scripts. All raw sequences were deposited at the OMBC [89].

7.4.9

16S rRNA Alignment

All bacterial 16S rRNA gene sequences that we believe represent oral taxa and named human oral species in GenBank were entered. Evolutionary analyses were conducted by exporting aligned sequences from our database in MEGA7 [90]. Phylogenetic trees were made using the Neighbour-Joining method [52]. Bootstrapping was performed using 1000 replicates [53].

7.4.10

Database and Web Design

The backbone of our platform is an industrial standard LAMP system (Fig. 7.4). Linux (CentOS) provides maximum stability and a multithread computation environment as the operating system; Apache provides the most important and fundamental function as the web service; MySQL works as the relational database,

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297

Fig. 7.4 Description of the essential elements and main methodology used in the establishment of the OMBC. The backbone of the database is the LAMP model

and PHP is adopted for dynamic web page rendering. PHP is also used to code the Common Gateway Interface (CGI) to the relational database. Our database can be accessed via the URL http://www.sklod.org/ombc

7.4.11

Curation

The project investigators Peng Xian, Zhou Xuedong, Xu Xin, Li Yuqing, Li Yan, Li Jiyao, Su Xiaoquan, Huang Shi, Xu Jian and Liao Ga carried out the curation of the database. These investigators reviewed each item on the taxon description page.

7.4.12

Service and Function

The database and query and statistical functions deployed were designed and developed according

to the suggestions of more than 20 professionals in microbiome research with more than 10 years of expertise in this field. The key features of our database include the following: (1) The database has been designed with a user-friendly interface. Users can start using the core functions immediately without professional training. (2) The backoffice management system provides add, delete and modify record functions for administrators. We plan to make the system a public platform that enables users to upload their own bacterial strain information in the near future. (3) Query provides statistical functions that can efficiently analyse the trends and distributions of each variable of the dataset as a whole. (4) Export and backup functions plus a complete restoration mechanism ensure data security and integrity. Acknowledgments This work was supported by the National Key R&D Program of China (2017YFC0840100 and 2017YFC0840107), the Key Project for Frontier Research of Science and Technology

298 Department of Sichuan Province (2016JY0006 to X.Z.) and the National Natural Science Foundation of China (81670978 and 81430011 to X.Z, 81470746 and 81772275 to G.L, 81700963 to X.P). The content of this chapter was modified from a paper reported by our group in Int J Oral Sci (Peng X et al. 2018). The related contents are reused with permission. Conflicts of Interests The authors declare that they have no conflicts of interests.

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8

Invasion of Oral Microbiota into the Gut Bolei Li, Yang Ge, Lei Cheng, Benhua Zeng, Jinzhao Yu, Xian Peng, Jianhua Zhao, Wenxia Li, Biao Ren, Mingyun Li, Hong Wei, and Xuedong Zhou

Electronic Supplementary Material: The online version of this chapter (https://doi.org/10.1007/978-981-15-78991_8) contains supplementary material, which is available to authorized users. B. Li · Y. Ge · J. Yu State Key Laboratory of Oral Diseases, Sichuan University, Chengdu, China National Clinical Research Center for Oral Diseases, Sichuan Unziversity, Chengdu, China Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China L. Cheng · X. Zhou (*) State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China e-mail: [email protected] B. Zeng · H. Wei (*) Department of Laboratory Animal Science, College of Basic Medical Sciences, Third Military Medical University, Chongqing, China X. Peng · B. Ren · M. Li State Key Laboratory of Oral Diseases, Sichuan University, Chengdu, China National Clinical Research Center for Oral Diseases, Sichuan Unziversity, Chengdu, China J. Zhao Shanghai Majorbio Bio-pharm Technology Co., Ltd, Shanghai, China W. Li State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

Abstract

The oral microbiota is associated with oral diseases and digestive systemic diseases. Nevertheless, the causal relationship between them has not been completely elucidated, and colonization of the gut by oral bacteria is not clear due to the limitations of existing research models. The aim of this study was to develop a human oral microbiota-associated (HOMA) mouse model and to investigate the ecological invasion into the gut. By transplanting human saliva into germ-free (GF) mice, a HOMA mouse model was first constructed. 16S rRNA gene sequencing was used to reveal the biogeography of oral bacteria along the cephalocaudal axis of the digestive tract. In the HOMA mice, 84.78% of the detected genus-level taxa were specific to the donor. Principal component analysis (PCA) revealed that the donor oral microbiota clustered with those of the HOMA mice and were distinct from those of specific pathogen-free (SPF) mice. In HOMA mice, OTU counts decreased from the stomach and small intestine to the distal gut. The distal gut was dominated by Streptococcus, Veillonella, Haemophilus, Fusobacterium, Trichococcus, and Actinomyces. HOMA mice and human microbiotaassociated (HMA) mice along with the GF mice were then cohoused. Microbial communities of cohoused mice clustered together and were significantly separated from those of HOMA mice

# Zhejiang University Press 2020 X. Zhou, Y. Li (eds.), Atlas of Oral Microbiology: From Healthy Microflora to Disease, https://doi.org/10.1007/978-981-15-7899-1_8

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and HMA mice. The Source Tracker analysis and network analysis revealed more significant ecological invasion from oral bacteria in the small intestines, compared to the distal gut, of cohoused mice. In conclusion, a HOMA mouse model was successfully established. By overcoming the physical and microbial barrier, oral bacteria colonized the gut and profiled the gut microbiota, especially in the small intestine. Keywords

Oral microbiota · Gut microbiota · Mouse model · Ecological invasion

8.1

Introduction

Clinical trials have indicated that the oral microbiota is associated with dental caries and periodontitis [1–4], both of which give rise to an extensive loss of natural teeth in older people and are identified as public health problems worldwide [5]. Accumulating evidence has even linked the human oral microbiota to oral cancer [6, 7]. In recent years, oral microecology dysbiosis has been proven to cause periodontitis [4, 8] and regarded as an indicator to predict early childhood caries (ECC) [9]. Thus, the oral microbiota plays a key role in the initiation of oral diseases. An increasing number of clinical research studies of the oral microbiota are being designed. However, the clinical investigations are usually restricted by complex conditions, including ethical issues. Regardless, a prospective cohort clinical study [9] found that shifts in the microbiota preceded the manifestation of clinical symptoms of ECC. Unfortunately, most of the other studies were cross-sectional and could barely address whether the oral microbiota was the cause or effect in the development of oral diseases. In vitro models also have limitations due to the abundant uncultivated phylotypes in the mouth [10]. Animal models would have been considered a good choice to study the oral microbiota; however, the oral microbiota of mice, the most

common experiment animal model, differs from that of humans. Therefore, a HOMA mouse model, with an oral microbiota similar to the human donors, must be established to reveal the cause and effect relationships between the oral microbiota and host pathologies, like the HMA mouse model [11, 12]. Not only oral diseases but also oral bacteria are linked to various digestive systemic diseases, including inflammatory bowel disease [13, 14], colorectal cancer (CRC) [15], pancreatic cancer [16, 17], liver carcinoma [18], and liver cirrhosis [19]. Seedorf et al. [20] demonstrated that mouthderived bacteria such as Actinobacteria, Bacilli, Clostridia, Fusobacteria, and Epsilonproteobacteria are able to overcome the host physical barrier and persist in the germ-free distal gut. Comparing the gut microbiome of patients suffering from liver cirrhosis with that of healthy control individuals, Qin et al. [21] found that most (54%) of the patient-enriched faecal microbial species originated from the oral cavity, demonstrating that the oral microbiota had invaded the gut of patients with liver cirrhosis. These studies indicated that the oral microbiota influenced host health by invading and colonizing the gut. The colonization of oral microbiota in the gut is a key point to understand pathologic colonization, facilitating studies of the pathogenic mechanisms of oral bacteria in systemic digestive diseases. However, invasion by oral microbiota by overcoming host physical barriers and gut microbiota barriers at various regions along the cephalocaudal axis of the gut is not well described. To develop the HOMA mouse model, we introduced the human salivary microbiota into GF mice and created a well-defined, representative animal model of the human oral microbial ecosystem. Using the HOMA mouse model, we investigated the colonization of gut-selected oral bacteria along the longitudinal axis. Furthermore, we studied the competition of oral microbiota with the native gut microbiota in various regions of the gut and identified key bacteria during the ecological invasion, by cohousing HOMA mice, HMA mice, and GF mice (Fig. 8.1).

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Human Salivary Sample

GF Mouse

HOMA Mouse

HMA Mouse

Cohoused Mouse

Sacrifice

5 weeks

4 weeks

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5 weeks

Cohouse

Cohouse

Fig. 8.1 Design of the human microbiota transplant and cohousing experiments

8.2 8.2.1

Results The Oral Microbiota of the HOMA Mouse Model

The surveys of oral samples revealed the engraftment of the human oral microbiota: all bacterial phyla, classes, orders, 27 of 28 bacterial families, and 84.78% (39 of 46) of genus-level taxa were detected among the recipient mice. All seven genus-level taxa missed by the humanized mice exhibited a low abundance in the donor sample (0.21% on average). The oral microbiota of the donor was dominated by 11 genus-level taxa, with a high relative abundance (> 1%), of which 5, Veillonella, Fusobacterium, Streptococcus, Porphyromonas, and Haemophilus, maintained a high abundance (>1% on average) among the recipient mice. The others were depleted to a low abundance among the recipient mice (Table S1). To further identify the advantages of the HOMA mouse model, we compared the oral microbiota of HOMA mice with SPF mice. PCA revealed that the donor oral microbiota clustered

closely with the HOMA mouse but were distinct from SPF mouse microbiota, especially in PC1 (57.91%) (Fig. 8.2a). The oral microbiota of HOMA mice differed from that of SPF mice in taxonomic structure. Dominant genus-level taxa present in the donor saliva sample were significantly more abundant among HOMA mice than SPF mice, including Veillonella, Fusobacterium, Streptococcus, and Haemophilus (Fig. 8.2b, c).

8.2.2

Biogeography of the Host Gut-Selected Oral Microbiota

The 16S rRNA gene sequencing survey revealed that the oral bacteria colonized various segments of the gut. In the stomach, 18 genus-level taxa were detected, with a relative abundance of more than 0.1% on average, 11 of which had a relative abundance exceeding 0.5% on average. In the small intestine, the relative abundances of 23 genus-level taxa exceeded 0.1% on average. Those with a relative abundance greater than 0.5% on average were Streptococcus, Veillonella, Haemophilus, Enterococcus, Fusobacterium,

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Fig. 8.2 Advancement of the HOMA mouse model. (a) PCA score plot of the oral microbiota of the human donor (Donor_O, red), HOMA mice (HOMA_O, green), and SPF mice (SPF_O, blue) at the genus level. (b) Taxonomic cladogram for HOMA mouse-enriched taxa (red) and SPF mouse-enriched taxa (green) obtained by LEfSe analysis

of 16S sequences. (c) The HOMA mouse-enriched taxa are indicated by a negative LDA score (red), while the taxa enriched by SPF mice have a positive score (green). Taxa at the genus level with different abundances between groups and with an LDA score greater than 3.0 are shown

Acinetobacter, Enterobacteriaceae_unclassified, and Bacteroides. In the caecum, only six genuslevel taxa were detected, with a relative abundance greater than 0.1% on average, including Veillonella, Streptococcus, Haemophilus, Fusobacterium, Bacteroides, and Trichococcus. Genus-level taxa with a relative abundance greater than 0.1% in the colon were the same as those in the caecum. The main genus-level taxa in the whole gut were Streptococcus, Veillonella, Haemophilus, Fusobacterium, Trichococcus,

and Bacteroides (Fig. 8.3a, Table S2). All six main genus-level taxa in the gut were also the dominant genus-level taxa (>1%) in the mouth of the HOMA mouse (Table S1). Although the microbial communities colonizing various regions shared some main bacteria, the differences among them were clear. Principal coordinates analysis (PCoA) showed that microbial communities present in the caecum, colon, and faeces clustered together and were distinct from those in the stomach and small intestine

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Invasion of Oral Microbiota into the Gut

Fig. 8.3 Biogeography of gut-selected oral microbiota. (a)Heatmap of specimens showing the relative abundance of the main identified bacteria at the genus taxonomic level in each segment of HOMA mouse guts, including the stomach (St), small intestine (Si), caecum (Ce), colon (Co), and faeces (F). (b) PCoA score plot of the microbiota

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from each segment of HOMA mouse guts based on unweighted UniFrac metrics. (c)The Kruskal–Wallis test was used to compare the difference between each segment of HOMA mouse guts in the OTU count and Chao index (*P < 0.05, **P < 0.01)

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(Fig. 8.3b). OTU counts significantly decreased from the stomach and small intestine to the distal gut and from the caecum to faeces, as did the Chao index (Fig. 8.3c). Distal gut communities were depleted to a low diversity consortium. The relative abundances of Acinetobacter, Enterobacteriaceae_unclassified, Lactobacillus, Turicibacter, Proteobacteria_unclassified, and Moraxella decreased from the stomach and small intestine to the distal gut and faeces. The relative abundances of Parabacteroides, Lachnoclostridium, and Blautia decreased from the caecum and colon to the faeces (Fig. 8.3a). These results indicated that the oral bacteria were filtered out by the distal gut.

8.2.3

HOMA mouse (Fig. 8.4c). In the small intestine, seven genus-level taxa were significantly increased from HMA mice to cohoused mice, six of which were dominant genera in the mouth of the HOMA mouse: Enterococcus, Streptococcus, Empedobacter, Porphyromonas, Moraxella, and Trichococcus (Fig. 8.4d). In the distal gut, four genus-level taxa were significantly increased from HMA mice to cohoused mice, but none was the dominant genera in the mouth (Fig. 8.4e, f). Microbial Source Tracker was used to analyse the effects of cohousing on the flow of microbes between cage mates, which allowed us to determine whether the assembly processes were involved in shaping the communities. The results revealed significant ecological invasion by oral bacteria in the small intestine (Fig. 8.4g).

Ecological Invasion by Oral Microbiota in the Gut

PCoA revealed that the microbial communities in every segment could not be distinguished by the original grouping 28 days after cohousing (Fig. 8.4a). Therefore, the gut microbiota of the cohoused mice could be regarded as an aggregate, regardless of the original mouse group. The microbial communities of cohoused mice were closely clustered with those of HMA mice and distinct from those of HOMA mice in every segment (Fig. 8.4a), suggesting that the oral microbiota was unable to challenge the dominant position of the gut microbiota in the gut. Interestingly, further analysis without HOMA mice showed that the microbial communities of cohoused mice could also be separated from HMA mice in every segment (Fig. 8.4b). These results indicated that although the oral microbiota was almost protected by the gut microbiota barrier, it reshaped the native gut microbiota. To further understand the effect of the oral microbiota on the community composition of the gut microbiota, LEfSe analysis was used. In the stomach, seven genus-level taxa were significantly increased from HMA mice to cohoused mice. One of the seven genus-level taxa was Streptococcus, which was the dominant genus (relative abundance >1%) in the mouth of the

8.2.4

Porphyromonas Competed for Colonization with the Small Intestinal Microbiota

To further study the functional positions of oral bacteria in the microbial community colonizing the small intestine, the co-occurrence network of the top 50 abundant genus-level taxa was used. Porphyromonas was found to correlate negatively with Turicibacter (Fig. 8.5). Before invasion by the oral microbiota, Turicibacter was the most dominant genus in the small intestine with the highest relative abundance (40.40% on average). Following invasion by the oral microbiota, the relative abundance of Porphyromonas increased significantly, and the abundance of Turicibacter decreased to 8.79% on average (Fig. 8.4d, Fig. S1). Moreover, Porphyromonas was found to correlate positively with these genera dominating the mouth of the HOMA mouse, including Streptococcus, Enterococcus, Acinetobacter, Moraxella, Trichococcus, Fusobacterium, Flavobacterium, and Lactobacillus (Fig. 8.5, Table S1). These results suggested that Porphyromonas, as common oral bacteria, played a key role in competing for colonization with the native main genus in the small intestine.

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Fig. 8.4 The shift in microbial composition after cohousing. (a) PCoA score plot of the microbiota from each gut segment of HOMA mice, HMA mice, and cohoused mice. (b)PCoA score plot of the microbiota from each gut segment from HMA mice and cohoused mice. c HMA mouseenriched genus level taxa in the stomach are indicated by a positive LDA score (green), while the cohoused (c)

mouse-enriched taxa have a negative LDA score (red). (d) The HMA mouse-enriched genus level taxa in the small intestine are indicated by a positive LDA score (green), while the cohoused mouse-enriched taxa have a negative LDA score (red). (e) The HMA mouse-enriched genus level taxa in the caecum are indicated by a positive LDA score (green), while the cohoused mouse-enriched

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Fig. 8.5 The co-occurrence network was generated from the small intestinal microbiota of the cohoused mice. Different coloured edges represent a positive (red) and a negative (blue) correlation, respectively. Each node

8.3

Discussion

In the past years, cumulative research data have implied a tight association between dysbiosis of the oral microbiota and diseases [3, 6, 7, 16, 22]. However, it has been difficult to verify the contribution of the oral microbiota to diseases via clinical studies due to their limitations. The lack of understanding of the effect and pathogenic mechanism of dysbiotic oral microbiota manifests in a great gap between the large amount of data and clinical applications [23]. Thus, for oral microbiota investigations, the establishment of a

Fig. 8.4 (continued) taxa have a negative LDA score (red). (f) The HMA mouse-enriched genus level taxa in the colon are indicated by a positive LDA score (green), while the cohoused mouse-enriched taxa have a negative LDA score (red). (g) Microbial Source Tracker analysis

represents a genus-level taxon, and the size of each node is proportional to the abundance. The colour of the nodes indicates their classification at the phylum level

HOMA mouse model can play an important role in translational medicine, similar to the HMA mouse model. In the present study, 84.78% (39 of 46) of the genus-level taxa were from donor saliva, similar to the HMA mouse model receiving 11 of 12 bacterial classes, and 88% (58 of 66) of the genus-level taxa were human [12]. Additionally, in subsequent study, we inoculated the contents of another two donor salivary glands into GF mice and obtained similar results [24]. Additionally, the HOMA mouse was a better representative for the donor than traditional SPF mice (Fig. 8.2a). Therefore, it is not difficult to conclude that the HOMA mouse

showed the proportions of the different sources present in the microbiota of the cohoused mice in each gut segment. The Kruskal–Wallis test was used to compare the proportions of the oral sources present in each gut segment of the cohoused mice (*P < 0.05, **P < 0.01)

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model was established successfully. Currently, the HMA mouse is an ideal model to study the role of the disease-associated gut microbiome [11]. In future, we believe that the HOMA mouse model could be used to investigate the effect of a dysbiotic oral microbiota on oral diseases, such as dental caries, periodontics, and oral cancer. In addition to oral disease, the HOMA mouse model will be applied to verify whether the oral microbiota is associated with some digestive systemic diseases. In most previous studies, the faecal microbiota was collected to represent the gut microbiota; however, some researchers have had different opinions and have suggested to divide the digestive tract into different sections to study the gut microbiota [25]. By collecting ileostomy samples from humans, Zoetendal et al. [26] found that the small intestine was enriched with Streptococcus sp. and Escherichia coli. Interestingly, in the present analysis, invasion by oral bacteria into the small intestine increased the relative abundance of Streptococcus and Enterobacteriaceae (Fig. 8.4d). Furthermore, in the small intestines of the cohoused mice, nearly 40% of the taxa were from oral microbial communities, which reshaped the community composition in the small intestine of the HMA mouse (Fig. 8.4g). Thus, especially in the small intestine, the oral microbiota played an important role in building the integrated gut microbiota. In the present study, oral bacteria overcame the host physical barrier and colonized the gut in HOMA mice (Fig. 8.3a, Table S2). However, in cohoused mice, the oral bacteria showed minimal colonization of the gut, especially the distal gut (Fig. 8.4g). This result is consistent with a previous study [20], in which all the distal guts of HMA mice cohoused with mice with the microbiota from soil or zebrafish were dominated by caecum-derived microbiota at 7 days after cohousing. These results indicated that gut microbiota plays an important role as a barrier in resisting the foreign bacteria from mouth. This resistance might due to greater acceptability in the gut of the gut microbiota than the oral microbiota and the creation of a more stable microenvironment by the gut microbiota to resist

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foreign bacteria. However, the microbiota barrier of the gut was not consistently indestructible, especially in the small intestine, where six of seven increasing genus-level taxa in the cohoused mice were dominant genera in the mouth of the HOMA mouse, including Porphyromonas (Fig. 8.4d). As a key oral genus to overcome the gut microbiota barrier, Porphyromonas was tightly associated with these genera that dominated the mouth of the HOMA mouse, but it correlated negatively with Turicibacter, the most dominant genus in the small intestine of HMA mice. Prior to invasion by oral microbiota, the relative abundance of Turicibacter in other regions of the gut was lower than that in the small intestine (Fig. S1), which might explain why more oral bacteria invaded the small intestine instead of the other regions. The small intestine is responsible for the majority of substance transformation [27] and is covered by a thinner mucin layer than the distal gut [28]. Thus, the small intestinal microbiota more effectively impacts digestive systemic health, suggesting that ecological invasion in the small intestine by Porphyromonas had a marked effect on digestive systemic health. For example, oral administration of Porphyromonas gingivalis, belonging to Porphyromonas, has been confirmed to induce gut microbiota dysbiosis and impair mucosal barrier function, leading to the dissemination of Enterobacteriaceae to the liver [29, 30]. Another interesting phenomenon is revealed by the barrier function of the gut microbiota. Fusobacterium overcame the physical barrier and became the dominant genus in the gut of the HOMA mouse. However, after receiving the gut microbiota by cohousing, the abundance of Fusobacterium decreased dramatically, and even the gut microbiota barrier was partly overcome by oral microbiota in the small intestine. Fusobacterium was still stopped by the microbiota barrier, but the resistance to Fusobacterium was supported by the gut microbiota from a healthy donor here. Those individuals suffering CRC fail to resist Fusobacterium [15, 31, 32]. The accumulating Fusobacterium nucleatum overcome the defective gut microbiota barrier from the CRC patient

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and further promote tumour development [33– 35]. In conclusion, resistance from various gut microbial communities is a key point to understand the effect of oral microbiota on gut microbiota and digestive systemic health, and it should be investigated in future studies. Overall, we first established a HOMA mouse model, which copied the oral microbiota of the human donor. Using this animal model, we found that both physical and microbiota barriers filtrated the oral microbiota in the digestive tract. Additionally, the oral microbiota invaded and profiled the gut microbiota, especially in the small intestine. Oral Porphyromonas was the key bacterial species competing with the small intestinal microbiota.

8.4 8.4.1

Materials and Methods Sample Collection from Humans

The study was authorized by the Ethical Committee of Sichuan University (WCHSIRB-D-2016070). The saliva was collected using a sterilized tube from an adult donor with natural dentition without periodontitis or active caries and without the use of antibiotics in the previous 3 months. The donor was required not to brush teeth for 24 h and abstain from food/drink intake for 2 h prior to donating saliva. Faeces were collected from the same person using a sterilized sealable plastic bag. A portion of the saliva and faeces was sent to the lab and inoculated into GF mice within 30 min. The rest was stored immediately at
80  C.

were bred in plastic gnotobiotic isolators, where the temperature and humidity were maintained at 20–26  C and 40–70%, respectively. They were fed a standard diet (GB-T14924.3–2001) sterilized by 60 co gamma radiation. Thirteenweek-old SPF mice were also maintained in the Experimental Animal Research Center. They were fed in the barrier housing facility.

8.4.3

To establish the HOMA mouse model, swabs dipped in 200 μL fresh saliva from the male donor were used to seed oral microbiota in the GF mice (n ¼ 13) by swabbing without anaesthesia. Swabbing was performed only once. The HMA mouse model was developed as previously described [36]. The faeces were resuspended in 10 mL sterile potassium phosphate buffer (0.1 mol•L
1, pH 7.2). Eight GF mice were inoculated by intragastric gavage with 1 mL human faeces suspension each, and 2-mL aliquots were spread on the fur. HOMA mice and HMA mice were bred in separated plastic gnotobiotic isolators. After 35 days, oral microbial samples were collected from the HOMA mice with swabs. The oral microbial samples from SPF mice were collected in the same way. Faeces of HOMA mice were also collected. Six of thirteen HOMA mice and six of eight HMA mice were subsequently sacrificed randomly, and the contents of the stomach, small intestine, caecum, and colon were collected. All the samples were immediately stored at
80  C.

8.4.4 8.4.2

Establishment of the HOMA and HMA Mouse Models

Cohousing Experiment

Animal Husbandry

The animal experimentation protocols were approved by the Ethical Committee of Sichuan University (WCHSIRB-D-2016-118) and the Third Military Medical University. Six-week-old GF male Kunming mice were maintained in the Experimental Animal Research Center at the Third Military Medical University. All GF mice

Two HOMA mice and two HMA mice was transferred to a new germ-free plastic isolator containing two GF mice (Fig. 8.1). These six mice were then distributed into two triads, each of which included a HOMA mouse, a HMA mouse, and a GF mouse housed in one cage, by which the animals could exchange components of their microbiota. After 28 days, the cohoused

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mice were sacrificed, and the contents of the stomach, small intestine, caecum, and colon were collected. All these samples were immediately stored at
80  C.

8.4.5

16S rRNA Gene Sequencing

The samples were processed by Shanghai Majorbio Bio-Pharm Technology Co., Ltd. (Shanghai, China). Total DNA was extracted, amplified, and sequenced according to standard procedures [37, 38]. Briefly, microbial DNA was extracted using the E.Z.N.A.® Soil DNA Kit (Omega Bio-Tek, Norcross, GA, USA) according to the manufacturer’s protocol. The DNA concentration was assessed using a NanoDrop (Thermo Scientific), and the quality was determined by agarose gel electrophoresis. Bacterial 16S rRNA gene sequences spanning the variable regions V4–V5 were amplified using the primer 515F_907R. The amplicons were then extracted from 2% agarose gels and further purified using the AxyPrep DNA Gel Extraction Kit (Axygen Biosciences, Union City, CA, USA) and quantified by QuantiFluor™-ST (Promega, USA). Purified amplicons were pooled in equimolar amounts and subjected to paired-end sequencing (2  300) on an Illumina MiSeq platform.

8.4.6

Bioinformatics and Statistical Analysis

Raw fastq files were demultiplexed and qualityfiltered by QIIME (version 1.9.1) [39]. Operational taxonomic units (OTUs) were clustered with a 97% similarity cut-off using UPARSE (version 7.1). The taxonomy of each 16S rRNA gene sequence was analysed using the RDP Classifier (http://rdp.cme.msu.edu/) against the SILVA rRNA database (http://www.arb-silva.de) with a confidence threshold of 70%. After the elimination of interference sequence, alpha diversity estimator calculations were performed using Mothur v.1.30.2. Phylogenetic beta diversity measures, such as unweighted UniFrac distance

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metrics analysis, were determined using the representative sequences of OTUs for each sample, and PCA and PCoA were conducted according to the distance matrices. LEfSe analysis (linear discriminant analysis [LDA] coupled to effect size measurements) was conducted to calculate bacteria with significant difference in relative abundance between the groups. Using a normalized relative abundance matrix, LEfSe showed taxa with significantly different abundances, and LDA estimated the effect size of the feature [37, 40]. In this study, a P value threshold of 0.05 (Wilcoxon rank-sum test) and an effect size threshold of 3 were used for all bacteria discussed. Microbial Source Tracker analysis was performed using the Source Tracker package based on Bayesian inference [20, 41]. The co-occurrence network of the top 50 abundant genus-level taxa was inferred based on the Spearman correlation matrix with a strict p-value threshold (P < 0.05) and a high correlation value (r > 0.6) to filter strong correlations. The combined result was exported to Cytoscape V.3.2.1 [37]. The data were subjected to nonparametric Kruskal–Wallis analysis. Differences were considered significant when P < 0.05. SPSS21.0 software (SPSS Inc., Chicago, IL, USA) was used for statistical analysis. Data Availability The raw reads were deposited into the NCBI Sequence Read Archive (SRA) database (Accession Number: SRP116564). Competing Financial Interests The authors declare that they have no competing financial interests. Acknowledgments This study was supported by the National Key Research and Development Program of China 2016YFC1102700 (X.Z.); National Natural Science Foundation of China grant 81372889 (L.C.), 81370906 (W.H.), 81600858 (B.R.), and 81430011 (X.Z.); Youth Grant of the Science and Technology Department of Sichuan Province, China 2017JQ0028 (L.C.); and National Basic Research Program of China 973 Program 2013CB532406 (W.H). The content of this chapter was modified from a paper reported by our group in Int J Oral Sci (Li B et al. 2019). The related contents are reused with permission.

312 Conflict of Interests The authors declare that they have no conflicts of interest. Authors’ Contributions Lei Cheng, Xuedong Zhou, and Hong Wei conceived and designed the experiments; Bolei Li, Jinzhao Yu, Benhua Zeng, Xian Peng Wenxia Li, Biao Renand, and Mingyun Li performed the experiments; Bolei Li, Yang Ge, and Jianhua Zhao analysed the data; Bolei Li and Yang Ge wrote the manuscript; and Hong Wei and Lei Cheng revised the manuscript. Supplementary information accompanies the manuscript at the International Journal of Oral Science website: http://www.nature.com/ijos.

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32. Chen W, Liu F, Ling Z, Tong X, Xiang C. Human intestinal lumen and mucosa-associated microbiota in patients with colorectal cancer. PLoS One. 2012;7: e39743. 33. Rubinstein MR, et al. Fusobacterium nucleatum promotes colorectal carcinogenesis by modulating E-cadherin/beta-catenin signaling via its FadA adhesin. Cell Host Microbe. 2013;14:195–206. 34. Gur C, et al. Binding of the Fap2 protein of Fusobacterium nucleatum to human inhibitory receptor TIGIT protects tumors from immune cell attack. Immunity. 2015;42:344–55. 35. Yu T, et al. Fusobacterium nucleatum promotes chemoresistance to colorectal cancer by modulating autophagy. Cell. 2017;170:548–563.e516. 36. Zeng B, et al. Effects of age and strain on the microbiota colonization in an infant human flora-

313 associated mouse model. Curr Microbiol. 2013;67:313–21. 37. Wang AH, et al. Human colorectal mucosal microbiota correlates with its host niche physiology revealed by endomicroscopy. Sci Rep. 2016;6:21952. 38. Zhu Y, et al. Meat, dairy and plant proteins alter bacterial composition of rat gut bacteria. Sci Rep. 2015;5:15220. 39. Caporaso JG, et al. QIIME allows analysis of highthroughput community sequencing data. Nat Methods. 2010;7:335–6. 40. Ling Z, et al. Alterations in the fecal microbiota of patients with HIV-1 infection: an observational study in a Chinese population. Sci Rep. 2016;6:30673. 41. Knights D, et al. Bayesian community-wide cultureindependent microbial source tracking. Nat Methods. 2011;8:761–3.

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Mycobiome Dysbiosis in Oral Lichen Planus Yan Li, Kun Wang, Bo Zhang, Qichao Tu, Yufei Yao, Bomiao Cui, Biao Ren, Jinzhi He, Xin Shen, Joy D. VanNostrand, Jizhong Zhou, Wenyuan Shi, Liying Xiao, Changqing Lu, and Xuedong Zhou

Abstract

Electronic Supplementary Material: The online version of this chapter (https://doi.org/10.1007/978-981-15-78991_9) contains supplementary material, which is available to authorized users. Y. Li · K. Wang · B. Zhang · Y. Yao · B. Cui · B. Ren · J. He · X. Shen · L. Xiao State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Q. Tu Institute of Marine Science and Technology, Shandong University, Qingdao, China Institute for Environmental Genomics, Department of Microbiology and Plant Biology, University of Oklahoma, Norman, OK, USA J. D. VanNostrand · J. Zhou Institute for Environmental Genomics, Department of Microbiology and Plant Biology, University of Oklahoma, Norman, OK, USA W. Shi The Forsyth Institute, Cambridge, MA, USA C. Lu Department of Anatomy, West China School of Basic Medical and Forensic Medicine, Sichuan University, Chengdu, China X. Zhou (*) State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China e-mail: [email protected]

The biodiversity of the mycobiome, an important component of the oral microbial community, and the roles of fungal-bacterial and fungal-immune system interactions in the pathogenesis of oral lichen planus (OLP) remain largely uncharacterized. In this study, we sequenced the salivary mycobiome and bacteriome (Wang et al Sci Rep 6:22943, 2016) associated with OLP. First, we described the dysbiosis of the microbiome in OLP patients, which exhibits lower levels of fungi and higher levels of bacteria. Significantly higher abundances of the fungi Candida and Aspergillus in patients with reticular OLP and of Alternaria and Sclerotiniaceae_unidentified in patients with erosive OLP were observed compared to the healthy controls. Aspergillus was identified as an “OLPassociated” fungus because of its detection at a higher frequency than in the healthy controls. Second, the co-occurrence patterns of the salivary mycobiome-bacteriome demonstrated negative associations between specific fungal and bacterial taxa identified in the healthy controls, which diminished in the reticular OLP group and even became positive in the erosive OLP group. Moreover, the oral cavities of OLP patients were colonized by dysbiotic oral flora with lower ecological network complexity and decreased fungal Firmicutes and increased fungal Bacteroidetes

# Zhejiang University Press 2020 X. Zhou, Y. Li (eds.), Atlas of Oral Microbiology: From Healthy Microflora to Disease, https://doi.org/10.1007/978-981-15-7899-1_9

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sub-networks. Third, several keystone fungal genera (Bovista, Erysiphe, Psathyrella, etc.) demonstrated significant correlations with clinical scores and IL-17 levels. Thus, we established that fungal dysbiosis is associated with the aggravation of OLP. Fungal dysbiosis could alter the salivary bacteriome or may reflect a direct effect of host immunity, which participates in OLP pathogenesis. Keywords

Oral lichen planus · Network assay · Mycobiome · Bacteriome · Host immunity

9.1

Introduction

Oral lichen planus (OLP) is a chronic oral mucosal disease that occurs in approximately 0.5% to 2% of the general adult population [2], with an even higher prevalence among women. In clinical settings, OLP is classified into three subtypes (reticular, atrophic, and ulcerative) and affects the buccal mucosa in the vast majority of cases. The gingiva, tongue, and lips may also be affected. The reticular form is typically asymptomatic and is the most common type, characterized by the presence of Wickham striae. However, the atrophic and erosive types may cause different degrees of discomfort and soreness, demonstrating high risks for malignant transformation at rates of 1–2% (range of 0–12.5%) [3, 4]. The precise etiology of OLP is uncertain, which is a major obstacle in the development of new therapeutics. Various factors have been considered to be potential causes of OLP, such as infection, immunity, genetic factors, stress, and trauma [2]. However, the precise roles of these factors have been debated. Over the last decade, microbial infection has received increasing attention in the context of OLP pathogenesis. Previously, we evaluated differences in the salivary microbial communities between OLP patients and healthy individuals. We observed that the bacterial community in saliva from OLP patients

was characterized by greater variety and less bacterial specificity, comprising only Porphyromonas and Solobacterium [1], which exhibited significantly higher abundances compared with the healthy controls. Additionally, a decrease in Streptococcus and an increase in gingivitis
/periodontitis-associated bacteria were observed in OLP lesions in another study [5]. These findings implicated a link between oral bacterial dysbiosis and OLP. It is noteworthy that the oral cavity is colonized by both bacteria and fungi, the latter of which have been known to have a role in OLP for a long time. Among oral fungi, Candida species have been reported to be associated with OLP and are detected in 37–50% of OLP patients [6]. Candida albicans is the most predominant OLP-associated Candida species and is involved in the malignant transformation of OLP [7]. The carriage rate for Candida albicans in patients with erosive OLP is much higher than that observed in patients with non-erosive OLP or in healthy controls [8]. Additionally, non-Candida albicans species have been specifically isolated from OLP patients, indicating a possible association between these yeasts and OLP [9]. However, previous studies have primarily focused on fungal epidemiology, such as the carriage rate for Candida species in OLP patients, with few studies analyzing the biodiversity and composition of the entire fungal community living in symbiosis with bacteria in the oral ecosystem. The advent of the use of nextgeneration sequencing technology to evaluate microbial diversity has broadened our view of the importance of fungi. The salivary mycobiome, which primarily refers to the fungal microbiota, is an important component of the oral microbiome. Ghannoum et al. [10] and Dupuy et al. [11] evaluated the complexity of the core oral mycobiomes of healthy individuals. However, these findings did not significantly contribute to a wider understanding of the disease state. Moreover, the interaction between the mycobiome and the resident bacterial microbiome may be crucial for the progression of diseases such as inflammatory bowel disease, cystic fibrosis, and oral diseases [12– 14]. Interactions between fungi and commensal

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bacteria involve physical binding, signaling molecule communication, and metabolic exchange in oral niches [14]. Although microbial infection has been proposed to be a causative, associated or possibly worsening factor in OLP, little is known regarding the oral fungal-bacterial relationship in OLP progression. In addition, OLP is considered a T-cellmediated inflammatory disease because the infiltrating lymphocytes are primarily T cells. Recently, the Th17 subset of CD4+ T-helper cells was shown to play a crucial role in promoting immune inflammatory reactions in the defense against infection by extracellular microorganisms and in autoimmune disease. Moreover, numerous Th17 cells have been identified in OLP lesions [15], and interleukin (IL)-17 and IL-23, cytokines secreted by Th17 cells, are important components involved in the defense against pathogenic microorganisms [16]. For example, salivary IL-17 and IL-23 are significantly correlated with specific bacterial genera in OLP, such as Porphyromonas, indicating their potential roles in the pathogenic mechanism of OLP [4]. Accumulating evidence has also implicated IL-17 and IL-23 in immunity to fungal pathogens, with the IL-23/IL-17 axis being essential in the defense against Pneumocystis carinii. Conversely, this axis amplifies the inflammatory pathology in mouse models of Candida or Aspergillus infection [17]. Similar to the gut microbiome [12, 18, 19], inter-kingdom interactions between bacteria and fungi may be substantial in the oral cavity. Because the host immune system is a major stress that modulates microbial composition [14, 20– 22], perturbations in salivary IL-17 and IL-23 levels, as well as the altered oral bacteriome observed in our previous study, suggest a disequilibrium within the oral mycobiomes of OLP patients. To test this hypothesis, we evaluated the salivary fungal abundance, frequency, and diversity in OLP and explored the complex and dynamic ecological relationships between the fungal mycobiome, oral bacteria, and host immunity. Our results indicated that fungal community composition and diversity are dramatically altered among OLP patients. Thus, despite their

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numerical inferiority, the oral mycobiome may be a driving force for bacteriome shifts though the modulation of mucosal immunity, which directly or indirectly affects OLP pathogenicity.

9.2 9.2.1

Results Participant Demographics and Sequence Data

The subjects enrolled in this study included 18 healthy subjects (age 39.72  11.02 years), 17 reticular OLP patients (age 43.58  9.97 years), and 18 erosive OLP patients (age 46.72  9.80 years). There were no significant differences in the age and gender distributions among the groups (P ¼ 0.127 and P ¼ 0.815, respectively). The severity of OLP was scored using a semiquantitative scoring system based on the site, area, and clinical presence of lesions [23]. Using an Illumina MiSeq sequencing platform, 1 580 028 raw paired-end reads of ITS region amplicons were obtained. After merging the forward and reverse reads and performing quality trimming, 712 295 merged sequences with an average length of 324 bp were obtained for all 53 samples. These sequences (335 185 for the healthy control samples, 197 963 for the reticular OLP samples, and 179 147 for the erosive OLP samples) were then clustered into 4 564 OTUs after quality trimming, dereplication, clustering, and chimera removal using the UPARSE pipeline, with an OTU identity cutoff of 97%. Of the 4 564 identified OTUs, 1 990 were singletons. The taxonomic assignments made using the RDP classifier showed that 4563 OTUs were fungi, with only one OTU with 2 sequences identified as protozoan but with 20% confidence. Among the fungal ITS OTUs, 1 588 belonged to Ascomycota, 20 to Chytridiomycota, 976 to Basidiomycota, 52 to Zygomycota, and 15 to Glomeromycota. The remaining OTUs were either assigned to unidentified fungi (124 OTUs) or known fungal phyla with 0.05,

Fig. 9.1a, b). Rarefaction analyses indicated that fungal species richness and alpha diversity among the three groups gradually decreased as the disease was aggravated (Fig. 9.1c, d), which was in contrast to the tendency of the bacteriome (Table S1) [1]. The phylogenetic structure was further analyzed. Although unweighted principal coordinate analysis showed no obvious separation among the mycobiomes of the healthy subjects, reticular OLP, and erosive OLP (Fig. S1), dissimilarity tests, including MRPP, adonis, and ANOSIM, did reveal significant differences between the healthy control group and the two OLP groups (P < 0.05, Table 9.1). However, no

Fig. 9.1 Diversity analysis of the salivary fungal communities in the healthy subject (H), reticular OLP (R), and erosive OLP (E) groups. The fungal community of erosive OLP displayed significantly lower richness and α-diversity compared to the healthy controls for various

diversity measures (P < 0.05). (a) Chao1 richness. (b) Shannon index. (c) Rarefaction curves of Chao1 richness obtained by combining samples in the same group. (d) Rarefaction curves of Shannon index obtained by combining samples in the same group

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Table 9.1 Comparison of the overall fungal community structure using three nonparametric statistical methods

H vs R H vs E R vs E

MRPP δ 0.777 0.807 0.725

P 0.001 0.002 0.254

Adonis F 0.083 0.079 0.038

P 0.002 0.002 0.16

ANOSIM R 0.144 0.171 0.02

P 0.005 0.001 0.162

H healthy control, R reticular OLP, E erosive OLP

dramatic differences were detected in reticular OLP when it was compared with erosive OLP (P > 0.05, Table 9.1).

9.2.3

Taxonomic Differences Among Healthy Individuals and OLP Patients

The fungal community composition was analyzed at different taxonomic levels. At the phylum level, significantly different patterns were observed for the top two prevalent phyla: Ascomycota (59.03% in healthy individuals, 69.58% in reticular OLP, and 68.22% in erosive OLP) and Basidiomycota (15.62%, 13.46%, and 7.33%, respectively, Fig. 9.2a). Phylum Ascomycota showed higher abundance in the reticular and erosive OLP groups, whereas the abundance of Basidiomycota was lower in the OLP groups compared with the healthy controls. At the family level, there were 11 fungal families for which no significant difference was observed between the OLP patients and healthy individuals (Fig. 9.2b). At the genus level, a total of 280 genera were detected. Among them, 126 genera were only present in one individual. The abundances of several genera were significantly different among the groups (Fig. 9.2c). The relative abundances of Candida and Aspergillus were significantly increased in the reticular OLP group compared with those observed in the healthy subjects. In contrast, Ascomycota_unidentified_1_1 and Trichosporon were strikingly more abundant in the healthy subjects than in those with erosive OLP. Furthermore,

significantly higher levels of Alternaria and Sclerotiniaceae_unidentified were observed in the erosive OLP group compared with the reticular OLP group. The most frequently detected fungi (constituting the “core” mycobiome) at the genus level with an average relative abundance above 0.1% are shown in Fig. 9.3. Among them, Candida and Ascomycota_unidentified_1_1 were the two genera with the highest detectable frequencies (96%) in all three groups. In addition, the frequencies of Phoma, Trichosporon, Penicillium, Aspergillus, Fungi_unidentified_1_1, and Coniochaeta were above 50% in all of the samples. No “OLP-specific” taxa (present in either the healthy or OLP groups) were detected. However, we identified Aspergillus as an “OLPassociated” fungus, as it was present at a higher frequency in the OLP group than in healthy controls. To further investigate the key oral fungal microbiota associated with OLP, we evaluated the genera and OTUs with frequencies of at least 50% and relative abundances of 0.5%. Aspergillus was only present in the reticular OLP group, while Phoma was detected in both the healthy subject and reticular OLP groups (Fig. S2a). Although Candida and Ascomycota_unidentified_1_1 were detected in all three groups, Candida was more abundant in the reticular OLP group, and the abundance of Ascomycota_unidentified_1_1 was significantly increased in the healthy subjects (Fig. 9.2c). Additionally, in terms of OTU levels, we observed that OTU_4429 (Candida) and OTU_21 (Phoma) were only present in the two OLP groups and were absent in the healthy control group (Fig. S2b).

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Fig. 9.2 Relative abundances of fungal phyla, families, and predominant genera (P > 0.1%) among the healthy subject (H), reticular OLP (R), and erosive OLP

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(E) groups. (a) Phylum level. (b) Family level. (c) Comparison of the top 20 abundant genera. (a) H vs R; (b) H vs E; (c) R vs E. Superscript letters indicate P < 0.05

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Fig. 9.3 Frequency of fungal genera with average relative abundance above 0.1% among the healthy subject (H), reticular OLP (R), and erosive OLP (E) groups. A total of 16 genera were included in this analysis

9.2.4

Inversion of Myco-Bacteriome Co-occurrence Patterns from Antagonization to Co-prosperity

Given the observation that bacterial-fungal interactions are actively present throughout the human body and that certain fungal taxa are distinctly distributed, we hypothesized that bacteriome-mycobiome co-occurrence and co-exclusion networks differed between the OLP patients and healthy controls. The bacterial-fungal network was constructed by only including the genera detected in no fewer than eight subjects. In total, 12 fungal and 29 bacterial genera were included, as shown in Fig. 9.4. Several interesting findings were obtained from the network. First, among the healthy individuals, most of the myco-bacteriome co-occurrence interactions were negative, whereas positive co-occurrence relationships were observed in the erosive OLP group. However, in the reticular OLP group, half of the correlations disappeared because some of the enrolled fungi were not detected in this group. Second, as the predominant fungal genus, Candida exhibited 12 significant inversions (negative to positive) with bacterial genera, of which 6 genera were

identified in the reticular OLP group (Abiotrophia, Actinobacillus, Aggregatibacter, Dialister, SR1 genera incertae sedis, and Treponema), and 6 were observed in erosive the OLP group (Bacteroides, Brachymonas, Capnocytophaga, Cellulosimicrobium, Planobacterium, and Veillonella) (Table S2).

9.2.5

Distinct Network Topology Between OLP and Healthy Individuals

We also constructed co-occurrence ecological networks at the OTU level by incorporating both fungal and bacterial OTUs to predict their ecological relationships involved in OLP (Fig. S3). Strikingly, the co-occurrence or mutual exclusion patterns of the three groups were significantly different. Decreased network complexity was observed from the healthy to the erosive OLP stages. In total, 1 175 associations and 336 nodes were observed in the healthy control group network, 1241 associations and 366 nodes were observed in the reticular OLP network, and 1 175 associations and 383 nodes were observed in the erosive OLP network. The constructed healthy control group network showed an average

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Fig. 9.4 Co-occurrence relationships between abundant fungal and bacterial genera across samples. Co-occurrence and co-exclusion relationships of genera present in at least eight subjects were explored by Pearson correlation coefficient analysis. The bacterial genera are shown on the left, and the fungal genera are positioned at the top. Fungal

genera belonging to Ascomycota are marked in blue, while Basidiomycota genera are marked in gray. Rectangle frames are used to highlight the negative myco-bacteriome co-occurrence interactions in the healthy control group, which changed to positive in the erosive OLP group

connectivity of 6.994, an average geodesic distance of 5.509, a modularity of 0.76, and a centralization of connectivity value of 0.069, while the networks of reticular and erosive OLP had average connectivities of 6.781 and 6.136, average geodesic distances of 6.05 and 7.22, modularities of 0.768 and 0.777, and centralization of connectivity values of 0.061 and 0.05, respectively (Fig. 9.5, Table S3). This finding

was further confirmed via sub-networks constructed by extracting the first bacterial neighbors of the fungal nodes with the highest connectivity (Fig. 9.6). Several interesting findings were observed. The number of significant correlations involving members of the phylum Firmicutes (black nodes, the majority belonging to Streptococcus) clearly decreased in the erosive OLP network compared with the

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Fig. 9.5 Fungal-bacterial co-occurrence network analysis of the healthy subject (H), reticular OLP (R), and erosive OLP (E) groups. Various network indices were used to describe the properties of the fungal-bacterial

co-occurrence patterns, including the total number of nodes (a), total number of links (b), average connectivity (c), average geodesic distance (d), modularity (e), and centralization of connectivity (f)

healthy control network. In contrast, the involvement of OTUs from the phylum Bacteroidetes (rose red nodes, primarily Prevotella, Porphyromonas, and Capnocytophaga) in the co-occurrence network increased significantly in the erosive OLP network. For fungal genera belonging to the phylum Ascomycota, such as Candida, far fewer co-occurrence events were observed in the reticular OLP network than in the erosive OLP network.

9.2.6

Fungal Disturbance Promotes OLP Exacerbation

We also examined the relationship between fungal genera and clinical parameters based on Pearson correlation coefficient values. The salivary concentrations of IL-17 and IL-23 were measured using an enzyme-linked immunosorbent assay (ELISA) [1]. In total, 29 fungal genera were observed to have significant correlations

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Fig. 9.6 Sub-network analysis of fungal-bacterial relationships in the healthy subject (H), reticular OLP (R), and erosive OLP (E) groups. Sub-networks for the H, R, and E groups were constructed by extracting all

of the bacterial OTUs connected with the fungal OTUs. The nodes in the inner circle are fungal OTUs, and nodes in the outer circle are bacterial OTUs

Fig. 9.7 Relationship between the relative abundances of fungal genera and clinical parameters. Three clinical parameters were analyzed, including the clinical score

and IL-17 and IL-23 levels. Pearson correlation coefficient was performed. * indicates P < 0.05

with clinical parameters, including clinical scores and salivary levels of IL-17 and IL-23, and were therefore identified as keystone fungi in saliva (Fig. 9.7). Several interesting correlations were observed. First, there were significant positive correlation patterns between clinical scores and the fungal genera Erysiphe and Bovista, whereas Sordariomycetes unidentified 1 was negatively correlated with clinical scores. Second, regarding the correlation with immunologic factors involved in the inflammatory response in OLP,

several fungal genera, such as Dothiorella, Sympoventuria, and Mycosphaerella, showed significant positive correlations with salivary levels of IL-17. However, Sordariaceae unidentified, Helotiales unidentified 1, and Pestalotiopsis were negatively correlated with IL-17. Notably, no significant correlation was observed between salivary levels of IL-23 and fungal genera. Finally, among the fungal genera associated with clinical data, significant correlations with more than one parameter were determined for

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genera such as Bovista, which positively correlated with clinical scores and salivary levels of IL-17 simultaneously.

9.3

Discussion

Although numerous studies have emphasized the possible role of bacterial or viral infection in OLP [1, 4], the fungal component of the oral microbiome has not been thoroughly investigated. In the present study, we showed for the first time the structural characteristics of the core mycobiome in salivary samples from reticular and erosive OLP patients, which demonstrated lower biodiversity and an increased abundances and frequencies of the genera Candida and Aspergillus. The oral fungal community was less enriched in OLP patients compared with that observed in the healthy control group. Interestingly, the opposite pattern was observed for the bacteriome, which demonstrated significantly increased diversity in the OLP group compared to the healthy control group. The fungi-to-bacteria diversity ratio decreased sharply in the OLP group compared to the healthy control group. OLP is quite different from most other mucocutaneous diseases, such as atopic dermatitis, psoriasis, Crohn’s disease, and ulcerative colitis, which are associated with decreased diversity of the bacteriome [1, 24–26] and an increased diversity of the mycobiome [19]. This inverted mycobiome-to-bacteriome trend was similar to the results obtained by Hoarau in the gastrointestinal tract [18]. The results of a previous study showed that the Candida load negatively correlates with salivary bacterial diversity [27]. In addition, a study by Peleg study showed that anaerobic bacteria otherwise inhibit fungi [28]. Specific alterations in fungal diversity in parallel with variations in bacterial diversity implicate an oral microecological imbalance in OLP. Previous studies [10, 11] have reported that more than 100 fungal species are members of the oral flora. The results of our study further demonstrated the existence of oral mycobiota

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diversity, identifying 6 phyla, 11 families, and 280 genera of fungi. In particular, we evaluated the patterns of fungal genera associated with OLP, demonstrating an increase in opportunistic/pathogenic fungi and a decrease in symbiotic fungi. The relative abundance of Candida was higher in the reticular and erosive OLP groups (49.6% and 41.3%, respectively) than in the healthy subject group (27.1%), although a significant difference was only observed between the reticular OLP and healthy control groups. This result was in complete accordance with previous findings [29]. We propose the following possible causes of this increase in Candida abundance. First, the susceptibility of OLP patients to Candida may be increased compared with healthy controls. Second, Candida hyphae may prefer the nonlesional reticular mucosa to erosive mucosa. Third, the types of pathogenic Candida in the saliva of OLP patients may be different from those in healthy individuals, a hypothesis that is supported by our analyses at the OTU level. OTU_3662 (Candida) dominated in the saliva of the healthy control group, while the core species in the reticular and erosive OLP groups was OTU_4429 (Candida). Hoarau et al. [18] showed that C. tropicalis rather than C. albicans is the pathogen responsible for Crohn’s disease. Aspergillus, another opportunistic fungal pathogen involved in endodontic infection, cystic fibrosis [30, 31], and immunocompromised patients, may cause a spectrum of respiratory disease, wound infections, and biofilm formation on medical devices. We also observed a significantly higher abundance and frequency of Aspergillus in OLP patients than in the healthy control group. In cases of oral lesions associated with dimorphic fungi (Candida), filamentous fungi (Aspergillus spp.) have been reported to be present, but these instances typically involve severe immunosuppression and disseminated infection to extraoral sites [32]. Taking these findings into consideration, it is possible that alterations in the fungal population are driven by an expansion of Candida and Aspergillus in the oral mycobiota of OLP individuals. Similar results have revealed a higher susceptibility to Candida and Aspergillus infection in the absence

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of Toll IL1R8 (TIR8), a negative regulator of Th17 responses [33]. The overgrowth of native Candida and Aspergillus species may be positively correlated with OLP severity, suggesting a disease link. Moreover, a fungal genus associated with invasive diseases, Alternaria, was observed to have a richer abundance in individuals with erosive OLP rather than reticular OLP, indicating its potential pathogenicity with the development of OLP. Howard et al. [34] showed that asthma severity is associated with the presence of Alternaria species in the lung that may have originally been derived from the mouth. Significant differences in the abundance of Sclerotiniaceae, which has also been detected in Crohn’s disease [35], were observed between the erosive OLP group compared to reticular OLP and healthy control groups, possibly because it is a family of necrotrophic fungi. The results of the studies referenced above indicate that the oral mycobiome is involved in specific oral diseases as well as in respiratory and digestive diseases. Sixteen genera were present with frequencies greater than 20% in each group and were designated the “core” mycobiome, which exhibited substantial overlap with the core oral mycobiota described in two previous studies. Specifically, our results are in good agreement with those of Ghannoum et al. [10] and Dupuy et al. [11] with respect to the identification of Candida, Alternaria, Aspergillus, Cladosporium/Davidella, Saccharomyces, Phoma, and Malassezia. However, nine oral cavity-associated genera were uniquely identified in our study, including Ascomycota_unidentified_1_1, Trichosporon, Fungi_unidentified_1_1, and Podospora, among others. Candida species were the most prevalent in both healthy and diseased oral cavities, demonstrating a 96% carriage rate in the samples assayed in our study, higher than that observed in other studies (60–80%) [1, 10] and much higher than the culture rate of 17.7% [32]. In addition to analyzing disease-associated fungi, we further confirmed significant shifts in the salivary fungal-bacterial interactions in OLP

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patients by exploring the differences in microbial co-occurrence and co-exclusion patterns between healthy and OLP individuals. The most dominant fungal genus, Candida, was of particular interest. Candida was negatively correlated with 18 out of 29 bacterial genera in healthy individuals. In contrast, Candida was positively correlated with eight bacterial genera in reticular OLP and eight bacterial genera in erosive OLP. Some of them (Treponema, Aggregatibacter, Dialister, SR1, Bacteroides, Capnocytophaga, and Veillonella) are strict anaerobic periodontopathogenic genera. How such strict anaerobes survive in an aerobic niche such as the oral cavity may be explained by the relationship between Candida and the high level of O2 consumption that is typical of yeasts, which creates an anaerobic microniche to permit the growth and biofilm formation of these strict anaerobic bacteria under aerobic conditions [36]. Furthermore, lactic acid is the most preferred source of carbon for fungi under the hypoxic conditions created by C. albicans. Excluding metabolic interactions, Candida species also demonstrate positive physical interactions with bacteria. For example, co-aggregation promotes the growth of fungal cells in the biofilm core with bacteria around their periphery. Additionally, the Treponema flagellum forms a “bridge” between fungi and bacteria. With respect to chemical interactions, fungal ethanol secretion can enhance the growth and virulence of Acinetobacter baumannii. In contrast, bacteria may develop antibacterial tolerance by living under the protective fungal matrix umbrella [37]. Through the rapid consumption of molecular oxygen, the rapid increase in the local pH, the provision of a physical scaffold for the adhesion of oral bacteria, and the production of chemical factors that modulate oral bacteria, shifts in fungal communities may be a driving force for those that occur in bacterial communities. Mycobacterium infections have been shown to be associated with aspergillosis [38]. The abundance of Candida tropicalis has been observed to be positively correlated with the presence of Serratia marcescens and E. coli [18]. Although fungi only constitute

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approximately 0.1% of the total microbial load in the oral cavity [21], at least 10% of the biovolume compensates for the presence of these microbes. An ecological network is a representation of various biological relationships connected by pairwise links within an ecosystem [39]. By analyzing and then visualizing the spatial Pearson’s correlations between fungi and bacteria detected from saliva samples, an imbalanced microbial network was observed in patients with OLP. First, OLP patients, particularly those with erosive OLP, showed simpler co-occurrence patterns between the mycobiome and bacteriome, as evidenced by lower connectivity and higher modularity, suggesting that the fungal and bacterial nodes in the OLP networks were more sparsely connected. In addition, in the sub-networks, the correlation between Bacteroidetes and fungal species was increased, but the correlation between Firmicutes and fungal species was decreased in OLP, consistent with previous observations, such as the rapid consumption of molecular oxygen and the rapid increase in local pH. On one hand, most Bacteroidetes (including Prevotella and Porphyromonas) are strictly anaerobic bacteria, which may be favored by fungi at the expense of oxygen. Furthermore, Bacteroides excel at dominating the microbiota due to their ability to modulate surface polysaccharides in an effort to evade the host immune system [40]. On the other hand, the consumption of lactic acid by fungi causes the environment to become less acidic, which may influence the growth of most Firmicutes members (such as Lactobacillus or Streptococci). Moreover, Lactobacillus sp. stimulate the mammalian host to induce antifungal immunity in the mucosal membrane [21]. Additionally, as the most prevalent genus of the phylum Firmicutes, the abundance and networks of Streptococci were decreased in OLP patients, as was reported in our previous study [1] and in a separate study [41]. The alteration of such correlations indicates that active roles for the phyla Firmicutes and Bacteroidetes may be important for the severity and exacerbation of

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OLP. The opposite scenario has been observed with respect to obesity, inflammatory bowel diseases, and autism spectrum disorders, with increased Firmicutes and decreased Bacteroidetes observed. The phylum Firmicutes is enriched for genes encoding nutrient transporters, while the phylum Bacteroidetes enriched for genes linked to carbohydrate metabolism [42]. However, our results are in agreement with those of other studies. Sam et al. [19] observed an association between Candida and Bacteroides. Members of the genus Bacteroides are more abundant in individuals who consume a high protein diet, while the abundance of Candida is strongly associated with the recent consumption of carbohydrates. Thus, an increased connection between Bacteroides and fungi might contribute to OLP severity. Emerging evidence suggests that the entire community of microbial residents influences the balance of immune responses and microbial community dysbiosis may lead to deficient education of the host immune system followed by immunemediated diseases [40]. Furthermore, the expression of pro-inflammatory cytokines (e.g., IL-17 and IL-23) may be upregulated by the presence of pathogens and the immunomodulatory components of biofilms (e.g., fungal glucans and bacterial lipopolysaccharides), resulting in tissue damage and lesions [18, 22, 32]. In particular, IL-17, an inflammation-associated cytokine that reflects the immune dysregulation status, has emerged as a central player in the immunopathogenesis of OLP [15, 16, 43] Previously, we analyzed a potential association between the oral microbiome and IL-17 and IL-23 levels in the saliva of OLP patients [1]. In this study, we further screened oral fungal genera that are potentially associated with disease severity and immune dysfunction of OLP. In total, 23 fungal genera were analyzed, none of which were significantly associated with IL-23. A significant positive correlation was observed between IL-17 and IL-18 fungal genera, including Dothiorella, Sympoventuria, Mycosphaerella, and Psathyrella. In a previous study, the

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abundance of Psathyrella was significantly associated with Crohn’s disease, supporting the results of a previous study showing that IL-17 was essential for host defense against fungal infection [44]. Notably, the genera Bovista and Erysiphe showed significantly positive correlations with clinical scores, suggesting their involvement in the aggravation of OLP. Thus, they were defined as keystone fungal genera [37] that can modulate the host and the ecology in a manner that far outweighs their numerical representation in the community. Our results were consistent with those of a study by Wheeler et al. [45], who inoculated typically rare fungi (A. amstelodami, E. nigrum, and W. sebi) in mice and observed exaggerated immune responses, suggesting that these keystone fungi play important roles in immune homeostasis. Despite a scarcity of data, the antifungal treatment of OLP patients has been shown to improve the clinical symptoms of OLP [46]. Another study also described the involvement of fungi in the aggravation of inflammatory responses and the severity of gastrointestinal diseases [12]. Based on the correlation between the myco-bacteriome and clinical parameters observed in this study and in a previous investigation [43], we suggest that the mycobiome may interact with commensal bacteria to augment the mucosal inflammatory response. In contrast, cytokines IL-17 had been shown to influence fungal composition and are important for protecting against infections caused by fungi (Candida albicans, Aspergillus fumigatus, and Pneumocystis carinii) on mucosal surfaces [6, 15] through the release of pro-inflammatory cytokines, chemokines, and antimicrobial peptides. A functional deficiency in the Th17 cell subset is associated with a dysbiotic state characterized by Candida overgrowth [14]. Furthermore, signaling through the IL-17 receptor is crucial for protecting against candidiasis [47]. The results of these studies demonstrated that IL-17 can play a central role on influencing the composition of core fungi, such as Candida and Aspergillus. Thus, it was supposed that keystone fungi can boost the host immunity (e.g., IL-17) and shape the core fungi composition through IL-17.

Y. Li et al.

9.4 9.4.1

Materials and Methods Subject Recruitment and Sample Collection

Subjects with reticular OLP (n ¼ 17) and erosive OLP (n ¼ 18), who were diagnosed according to the clinical classification and definition of the World Health Organization, together with 18 sex- and age-matched healthy controls were recruited from the West China Hospital of Stomatology, Sichuan University. Demographic information was obtained, and an oral examination was performed. A semiquantitative scoring system [23] consistent with the site, area, and presence of OLP lesions was used to assess the clinical scores and severity of OLP. All subjects included in this study had not received treatment for OLP for at least 2 months and were asked to avoid drinking or eating for 2 h before oral sampling. Those with other oral (e.g., periodontitis or dental caries) or systemic diseases were excluded. To reflect the structural changes of the entire microbiome in the oral cavity and adopt a painless approach, approximately 5 ml of spontaneous whole unstimulated saliva (WUS) was collected in a sterile DNA-free conical tube from each subject between 8:00 and 11:00 AM following standard techniques as described previously [1]. All samples were carried to the laboratory on ice within 2 h and stored at
80  C before further processing. The methods were performed in accordance with approved guidelines.

9.4.2

Cytokine Assay

IL-17 and IL-23 levels in the saliva were measured by ELISA as described previously [43].

9.4.3

DNA Extraction

Genomic DNA was extracted from individual saliva samples using a Qiagen QIAamp® DNA Mini Kit (Qiagen, Valencia, CA, USA) according to the manufacturer’s instructions as previously

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Mycobiome Dysbiosis in Oral Lichen Planus

described [48]. Briefly, after thawing on ice, aliquots were pelleted at 5000  g for 10 min and resuspended in 600 μL sorbitol buffer (1 molL
1 sorbitol, 100 mmolL
1 EDTA, and 14 mmolL
1 ß-mercaptoethanol). After incubating with 200 U lyticase at 30  C for 30 min for cell lysis, protein digestion was achieved by adding Proteinase K and incubating the samples at 56  C for 1.5 h. The DNA was bound to a spin column filter, washed with 96–100% ethanol and then was washed with the two buffers supplied by the kit. The bound DNA was eluted from the spin column filter with 200 μL of the supplied elution buffer. DNA quality was assessed by measuring the absorbance ratios using a Nano Drop-1000 Spectrophotometer (NanoDrop Technologies Inc., Wilmington, DE, USA). DNA samples with ratios of 1.8–2.0 (for A260/280 nm) and >1.8 (for A260/230 nm) were likely to be free from contamination and were used for downstream experiments. Finally, the total DNA concentration was measured using a PicoGreen kit (Invitrogen, Carlsbad, CA, USA), and the extracts were frozen at
20  C for further analysis.

9.4.4

Illumina Sequencing

The ITS2 region was amplified from the fungal DNA using the primers gITS7F (GTGARTCATCGARTCTTTG) and ITS4R (TCCTCCGCTTATTGATATGC), the product of which is expected to be 309 bp (not including the primers) [49]. A two-step phasing amplicon sequencing approach (PAS) was performed to avoid the amplification biases introduced by long barcoded PCR primers [49, 50]. Sample libraries for sequencing were prepared according to the 500-cycle v2 MiSeq Reagent Cartridge Preparation Guide (Illumina, San Diego, CA, USA) as described previously [49]. Sequencing was performed for 251, 12, and 251 cycles for the forward, index, and reverse reads, respectively, at the Institute for Environmental Genomics, University of Oklahoma (Norman, OK, USA). The barcoded 16S rRNA amplicon sequencing was performed using an Illumina MiSeq platform

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using the primers F515 (5’-GTGCCAGCMGCCGCGG-30 ) and R806 (3’-TAATCTWTGGGVHCATCAG-50 ) at the Institute for Environmental Genomics, University of Oklahoma (Norman, OK, USA). The amplicons obtained from all of the samples were then sequenced on an Illumina MiSeq platform.

9.4.5

Data Preprocessing, OTU Clustering, and Taxonomic Classification

Data preprocessing and OTU clustering were performed as described previously [51]. Only the reads with perfectly matched barcodes were extracted and used for further data analysis. Quality trimming of raw reads was carried out using the program Btrim [52] with an average quality score cutoff of 30 and window size of 3. The paired-end reads were then joined using the program pear [53] with the default parameters. Further quality trimming and OTU clustering were carried out using the UPARSE pipeline. The joined reads were subjected to further quality control with a maximum expected error threshold of 0.5 and a length cutoff of 200. Qualifying reads were then dereplicated, sorted by size, and clustered into OTUs with 97% sequence identity. The OTU sequences were checked against the UNITE database, and potential chimeric sequences were removed. Finally, the qualifying reads were mapped to representative OTU sequences to calculate the relative abundance of each OTU. Taxonomic assignment for representative OTUs was carried out using the Ribosomal Database Project (RDP) classifier [54] trained by the UNITE database. A confidence cutoff of 50% was used for taxonomic information assignments, and a random subsampling of 2 227 reads per sample was performed for further statistical analysis.

9.4.6

Statistical Analysis

The preprocessed data were further analyzed using the following statistical methods. First, we used three different nonparametric multivariate

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analysis methods, including adonis (permutational multivariate analysis of variance using distance matrices), ANOSIM (analysis of similarities), and multi-response permutation procedure (MRPP) [1], as well as principle coordinate analysis (PCoA) to measure and visualize the overall differences in the fungal community structure between healthy and OLP individuals. Second, the fungal community diversity was assessed based on the Chao1 richness and Shannon diversity indices. Rarefaction analyses were performed using the program Mothur [55] by pooling samples within the same group. Student’s t-test was used to evaluate significant differences between healthy and OLP individuals, such as diversity indices and relative abundances of OTUs and taxonomic groups. Third, the Pearson correlation coefficient was used to construct bacterial-fungal co-occurrence patterns from the 16S rRNA gene and ITS amplicon data, which were also used for analyses of the association between fungi and clinical parameters. Bacterial and fungal OTUs present in more than 8 samples were extracted and used for correlation calculations by clustering and visualizing using the MeV package [56]. For better visualization, co-occurrence patterns with a Pearson correlation coefficient 0.6 and P-value 0.05 were extracted and plotted. Finally, OTU-level microbial co-occurrence networks were constructed and analyzed. The random matrix theory-based approach in the MENA pipeline [57] was used to construct the microbial co-occurrence networks. A Pearson correlation coefficient cutoff of 0.76 was determined by the random matrix theory approach by observing the transition point of the nearest neighbor spacing distribution of eigenvalues from Gaussian to Poisson distributions, representing two universal extreme distributions. In such networks, OTUs were represented by network nodes, while correlations were transformed into the links between them. Sub-networks representing fungal-bacterial co-occurrence networks were subsequently constructed by extracting the first neighbors of the fungal OTUs. The co-occurrence networks were then visualized using Cytoscape 3.2.1.

Y. Li et al.

9.5

Data Availability

All the ITS2 and 16S rRNA sequences were deposited at NCBI under accession number SRP067603.

9.6

Ethics Statement

Written informed consent was obtained from all of the participants in this study. All procedures were approved (WCHSIRB-ST-2015-070) by the local ethics committee of the West China Hospital of Stomatology, Sichuan University. Acknowledgments This study was supported by the National Key Research and Development Program of China (2016YFC1102700), the National Natural Science Foundation of China (Grant Nos. 81771085, 81430011, 81600858, and 81600874), and the Key Projects of Sichuan Provincial Health and Family Planning Commission (Grant No.16ZD021). The funders had no role in the study design, data collection and analysis, decision to publish, or preparation of the manuscript. The content of this chapter was modified from a paper reported by our group in Int J Oral Sci (Li Y et al. 2019). The related contents are reused with permission. Conflicts of Interest All of the authors declare no conflicts of interest. Author Contributions Conception and design of the experiments: Y.L., L. X., and X.Z.. Conducted the experiments: Y.L., B. Z., C.L., K.W., X. S., and J.V.N. Data processing and analysis: Q.T., Y.L., K.W., B.R., and J. H. Volunteer recruitment and sample collection: X. S., B. Z. B. C., and L. X. Manuscript writing: Y.L., K.W. B. Z., and C.L. Revision of the manuscript: Q.T., L. X., J.V.N., J.Z., W. S., and X.Z.

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331 innate and adaptive immunity in health and disease. Eur J Immunol. 2014;44:3166–81. 21. Bozena DK, Iwona D, Ilona K. The mycobiome – a friendly cross-talk between fungal colonizers and their host. Ann Parasitol. 2016;62:175–84. 22. Underhill DM, Iliev ID. The mycobiota: interactions between commensal fungi and the host immune system. Nat Rev Immunol. 2014;14:405–16. 23. Piboonniyom SO, Treister N, Pitiphat W, Woo SB. Scoring system for monitoring oral lichenoid lesions: a preliminary study. Oral Surg Oral Med Oral Pathol Oral Radiol Endod. 2005;99:696–703. 24. Alekseyenko AV, et al. Community differentiation of the cutaneous microbiota in psoriasis. Microbiome. 2013;1:31. 25. Lynde CW, et al. The skin microbiome in atopic dermatitis and its relationship to emollients. J Cutan Med Surg. 2016;20:21–8. 26. Pascal V, et al. A microbial signature for Crohn’s disease. Gut. 2017;66:813–22. 27. Kraneveld EA, et al. The relation between oral Candida load and bacterial microbiome profiles in Dutch older adults. PLoS One. 2012;7:e42770. 28. Peleg AY, Hogan DA, Mylonakis E. Medically important bacterial-fungal interactions. Nat Rev Microbiol. 2010;8:340–9. 29. Bokor-Bratic M, Cankovic M, Dragnic N. Unstimulated whole salivary flow rate and anxiolytics intake are independently associated with oral Candida infection in patients with oral lichen planus. Eur J Oral Sci. 2013;121:427–33. 30. Gomes CC, et al. Aspergillus in endodontic infection near the maxillary sinus. Braz J Otorhinolaryngol. 2015;81:527–32. 31. Burgel PR, Paugam A, Hubert D, Martin C. Aspergillus fumigatus in the cystic fibrosis lung: pros and cons of azole therapy. Infect Drug Resist. 2016;9:229–38. 32. Diaz PI, Hong BY, Dupuy AK, Strausbaugh LD. Mining the oral mycobiome: methods, components, and meaning. Virulence. 2017;8:313–23. 33. Mehdipour M, et al. Prevalence of Candida species in erosive oral lichen planus. J Dent Res Dent Clin Dent Prospects. 2010;4:14–6. 34. Artico G, et al. Prevalence of Candida spp., xerostomia, and hyposalivation in oral lichen planus--a controlled study. Oral Dis. 2014;20:e36–41. 35. Findley K, et al. Topographic diversity of fungal and bacterial communities in human skin. Nature. 2013;498:367–70. 36. Fox EP, et al. Anaerobic bacteria grow within Candida albicans biofilms and induce biofilm formation in suspension cultures. Curr Biol. 2014;24:2411–6. 37. Janus MM, Willems HM, Krom BP. Candida albicans in multispecies oral communities; a keystone commensal? Adv Exp Med Biol. 2016;931:13–20. 38. Amiri MRJ, et al. Invasive forms of Candida and Aspergillus in sputum samples of pulmonary tuberculosis patients attending the tuberculosis reference

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Intestinal Microbiota and Osteoporosis

10

Xin Xu, Xiaoyue Jia, Longyi Mo, Chengcheng Liu, Liwei Zheng, Quan Yuan, and Xuedong Zhou

X. Xu · X. Jia Department of Operative Dentistry and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China L. Mo State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China C. Liu Department of Periodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China L. Zheng Department of Pediatric Dentistry, West China Hospital of Stomatology, Sichuan University, Chengdu, China State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China Q. Yuan Department of Dental Implantology, West China Hospital of Stomatology, Sichuan University, Chengdu, China State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China X. Zhou (*) State Key Laboratory of Oral Diseases, National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

Abstract

Postmenopausal osteoporosis (PMO) is a prevalent metabolic bone disease characterized by bone loss and structural destruction, which increases the risk of fracture in postmenopausal women. Owing to the high morbidity and serious complications of PMO, many efforts have been devoted to its prophylaxis and treatment. The intestinal microbiota is the complex community of microorganisms colonizing the gastrointestinal tract. Probiotics, which are dietary or medical supplements consisting of beneficial intestinal bacteria, work in concert with endogenous intestinal microorganisms to maintain host health. Recent studies have revealed that bone loss in PMO is closely related to host immunity, which is influenced by the intestinal microbiota. The curative effects of probiotics on metabolic bone diseases have also been demonstrated. The effects of the intestinal microbiota on bone metabolism suggest a promising target for PMO management. This review seeks to summarize the critical effects of the intestinal microbiota and probiotics on PMO, with a focus on the molecular mechanisms underlying the pathogenic relationship between bacteria and host, and to define the possible treatment options.

Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, Chengdu, China e-mail: [email protected] # Zhejiang University Press 2020 X. Zhou, Y. Li (eds.), Atlas of Oral Microbiology: From Healthy Microflora to Disease, https://doi.org/10.1007/978-981-15-7899-1_10

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Keywords

Postmenopausal osteoporosis · Intestinal microbiota · Probiotics · Estrogen deficiency · Ovariectomy

10.1

Introduction

Postmenopausal osteoporosis (PMO) is an estrogen deficiency-induced metabolic bone disorder characterized by reduced bone strength, which increases the risk of fracture in postmenopausal women [1]. The onset of PMO is occult, without any obvious symptoms until a fracture occurs. The most prevalent complication is a fragility fracture, which often occurs in the hip, femur, or spine under nontraumatic or mildly traumatic conditions, resulting in pain, malformation, dysfunction, and even death. Studies showed that the mortality rate associated with a hip fracture was 17% in the first year [2] and approximately 12–20% within the two following years [3]. PMO is also a potential risk factor for oral bone loss and aggressive periodontitis in postmenopausal females. PMO animal models showed an equivalent bone loss in alveolar bone and femurs [4]. Compared with healthy postmenopausal women, patients afflicted with PMO also exhibited an inclination to more bone loss and lower bone mineral density (BMD) in the jaw, especially in postmenopausal females with preexisting periodontitis who suffered from accelerated alveolar bone loss under routine treatment [5–7]. In addition to bone loss and microstructural deterioration, PMO affects the osseous formation processes. Delayed osseous maturation and reduced bone regeneration during bone healing in ovariectomized (OVX) rats were reported. [8, 9] The high morbidity and serious complications of PMO have attracted major research efforts on its prophylaxis and treatment for decades. Current medications for the treatment of PMO include bisphosphonates, raloxifene, teriparatide and calcitonin, denosumab, estrogen, menopausal hormone therapy, etc. These medications can prevent bone loss and

increase bone mineral density, with a decreased risk of fractures in the vertebra, hip, or long bones [1, 10]. All of these pharmacological agents can reduce bone resorption by inhibiting osteoclasts, except teriparatide, which acts as an anabolic agent by activating or increasing osteoblast activity and prompting bone formation [1, 11]. Recent studies have demonstrated a close relationship between the intestinal microbiota and bone metabolism [12–15], providing evidence that the intestinal microbiome may serve as a potential therapeutic target for the treatment of PMO.

10.1.1

The Intestinal Microbiota and Its Regulators

The intestinal microbiota is the collection of microorganisms that colonize the gastrointestinal tract, which consists of approximately 10 trillion bacteria [16]. Obligate anaerobes such as Bacteroidetes and Firmicutes are the predominant residents of the healthy gastrointestinal tract, outnumbering aerobes and facultative anaerobes [16, 17]. Based on their roles in maintaining human health, intestinal microorganisms can be categorized into beneficial, harmful, and neutral bacteria. Both host and environmental factors can shape intestinal microbial composition and structure (Fig. 10.1). Animal experiments [18–21] and twin studies [22, 23] revealed that host genetic background had a significant impact on the abundance of the intestinal microbiota and the predisposition to the colonization of pathogens (e.g., Escherichia coli). Though still disputed, gender may be another host factor affecting intestinal microbiome species diversity [24, 25]. Environmental factors, including diet, lifestyle, hygiene, antibiotic treatment, and probiotics, also contribute to the alteration of the intestinal microbiota composition [26–31]. Notably, the effects of diet and antibiotics on the intestinal microbiota also depend on the host genetic background [32, 33]. Probiotics are defined as dietary or medical supplements consisting of live bacteria that can benefit the host if provided in adequate quantities [34–36]. Currently, approximately 20 types of

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335

Fig. 10.1 Regulators of the gut microbiota and mechanisms by which the gut microbiota regulates bone metabolism. Shaped by both host and environmental

factors, the gut microbiota regulates bone metabolism through various pathways, including the immune system, endocrine system, and influences on calcium balance

beneficial bacteria are used in probiotic supplements. They are generally classified into five categories, including lactobacilli, bifidobacteria, streptococci, yeast, and others [37]. Lactobacilli and bifidobacteria are the most commonly used probiotics. Probiotics can selectively ferment prebiotics, which contain soluble dietary fibers such as oligosaccharides and inulin, facilitating the production of beneficial products conducive to the growth of certain probiotics such as bifidobacteria [34, 38, 39]. However, it is still disputed whether probiotics can alter the gut microbiota composition. Randomized controlled trials (RCTs) in healthy adults indicated that probiotic intervention or probiotics-fermented products resulted in changes in intestinal microbiota composition or diversity [40– 43]. Although probiotics promoted the significant increase of certain bacteria, Bacteroides was the dominant genus under probiotics administration, while other bacteria such as Clostridiales were inhibited. [40, 41] In addition, the effect of probiotics on Clostridiales genera may be associated with the initial status of the intestinal

microbiome and butyrate concentrations [41]. RCTs in elder adults showed that the age-associated intestinal microbiota imbalance was restored by probiotic-based functional foods, with increased resident probiotic-related bacteria and decreased emergence of opportunistic pathogens [44, 45]. Animal experimentation also showed that probiotic administration improved the intestinal microbiota composition in hyperlipidemic rats by recovering the abundance of Bacteroidetes and Verrucomicrobia and reducing Firmicutes [46]. However, another RCT in healthy adults demonstrated that Lactobacillus rhamnosus GG (LGG) supplement induced no alteration in gut microbiota composition or diversity stability, except for a transient increased fecal excretion of probiotic-associated bacteria during the intervention [47]. Additionally, one RCT in healthy subjects and patients with irritable bowel syndrome (IBS) showed parallel, transient, and distinct increases in probiotics but limited changes in other specific bacteria in fecal samples of both healthy and IBS-afflicted subjects with Bifidobacterium infantis intervention [48].

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The Intestinal Microbiota Regulates Bone Metabolism

10.1.2.1 Involvement of the Intestinal Microbiota in Bone Metabolism The dynamic homeostasis of the gut microbiome is critical to health. Accumulating evidence has demonstrated that the gut microbiota is associated with physiological bone metabolism and a range of inflammatory or metabolic bone diseases [12– 15, 49, 50]. In animal experimentation, germ-free mice showed higher trabecular volume bone mineral density (vBMD) and improved histomorphologic indices in trabecula compared with conventionally raised (CONV-R) mice [12]. However, both trabecular BMD and cortical crosssectional area decreased when germ-free mice were recolonized by the gut microbiota, indicating that the gut microbiota is a major regulator of bone mass [12]. Microbial recolonization in germ-free mice induced an incipient acute decrease in bone mass but predominantly led to bone formation with a longer duration, leading to a new equilibrium in bone mass [14]. Furthermore, germ-free mice colonized with immature gut microbiota from donors of different ages or nutritional statuses showed varied femoral phenotypes, suggesting that the impact of the gut microbiota on bone morphologic properties is age/nutrition dependent [13]. Compromised bone biomechanical properties in mice were also induced by an altered gut microbiota resulting from immunodeficiency or long-term antibiotic intervention during growth [15]. Additionally, through post-weaning exposure to low-dose penicillin (LDP) or by introducing LDP to their mother in pregnancy, adult offspring with a perturbed gut microbiota showed altered bone mineral content (BMC) and BMD [51]. In addition to physiological condition, inflammatory or metabolic bone diseases, such as metabolic osteoarthritis, osteoporosis, and autoinflammatory osteomyelitis, are also associated with gut microbial alteration [49, 50, 52, 53]. The abundance of gut bacteria Lactobacillus spp. and Methanobrevibacter spp. was shown to have a significant relationship with the prediction of osteoarthritis

assessed by the Modified Mankin Score in rats [50]. Gut microbiota modified by diet regulated the production of IL-1β (Interleukin-1beta) and prevented the spontaneous development of osteomyelitis in Pstpip2cmo mice predisposed to autoinflammatory osteomyelitis [52, 53].

10.1.2.2 Mechanisms by Which the Gut Microbiota Regulates Bone Metabolism Gut microbiota can regulate bone metabolism, but the exact mechanisms are still unclear. Multiple approaches through which gut microbiota may regulate bone metabolism have been proposed, including actions on the immune system, endocrine system, and calcium absorption (Fig. 10.1). a. The gut microbiota regulates bone metabolism through the immune system. Recent studies have revealed a close interrelationship between the immune system and bone metabolism, leading to the development of “osteoimmunology,” which highlights the role of immune-related factors in modulating bone remodeling [54, 55]. In immune-mediated bone metabolism, the RANKL (Receptor activator NF kappa B ligand)-RANK-OPG axis and immunoreceptor tyrosine-based activation motif (ITAM) pathway play key roles in physiological bone turnover and bone diseases [54, 56]. Recently, it has been widely recognized that the gut microbiota can interact with the host immune system and further influence host health [57–59]. One study showed that altered immune status in germ-free mice (e.g., decreased pro-inflammatory cytokines, fewer CD4+ T cells, and reduced osteoclast/precursor cells in bone marrow) may account for the higher bone mass than in CONV-R mice [12]. Intestinal segmented filamentous bacteria in mice were shown to promote the production of IL-17 and IFN-γ (interferon-gamma), both of which played critical roles in the formation of osteoclasts and osteoblasts [60–62]. These studies suggest that the gut microbiota regulates bone metabolism by altering host immune status.

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b. The gut microbiota regulates bone metabolism through the endocrine system. In addition to the immune system, hormones are regarded as another important regulator of bone metabolism. As an autocrine or paracrine growth factor, insulin-like growth factor-1 (IGF-1) can promote the differentiation and growth of bone cells, including osteoblasts, osteoclasts, and chondrocytes, and enhance normal interactions among them [63–65]. Moreover, the IGF-1 signaling pathway is involved in the regulation of bone metabolism via both growth hormone and parathormone [64]. Intermittent administration of parathormone promoted bone formation by increasing local IGF-1 production and activating the IGF-1 signaling pathway in bone [64]. Growth hormone can directly or IGF-1-dependently target the growth plate to promote cartilage formation and longitudinal bone growth [66, 67]. Moreover, gonadal steroids, including estrogen and androgen, play key roles in the regulation of bone mass and turnover in bone metabolism [68–70]. Furthermore, serum neurotransmitter 5-hydroxytryptamine, namely, circulating serotonin with a hormone-like effect, can stimulate or inhibit bone formation, and dualdirectional effects may be gender/age dependent [71–74]. The gut microbiota, which is currently considered a novel “endocrine organ” of the human body, can engage in an interplay with the endocrine system (e.g., hypothalamic-pituitaryadrenal axis) and secrete hormones or hormonelike products to regulate host hormone levels, further influencing host health status [75, 76]. In animal experimentation, gut microbial colonization in germ-free mice significantly increased the serum IGF-1 level, resulting in bone growth and normalized bone mass [14]. Isoflavones, the compounds classified as phytoestrogens and structurally similar to endogenous estrogen, were converted into more estrogenic metabolite equol by specific gut microorganisms such as rod-shaped and gram-positive anaerobic bacteria in approximately 30–50% of humans [77– 81]. Polycyclic aromatic hydrocarbons— contaminants widely present in nature—can be bio-transformed into products with estrogenic

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activity by the human colon microbiota [82]. A recent study showed that the gut microbiota, especially spore-forming bacteria, can enhance the biosynthesis of serotonin by colonic enterochromaffin cells [83]. Despite the lack of direct evidence, it has been suggested that gut microbiota-bone communication likely depends on the endocrine system or hormone-like substances. c. The gut microbiota regulates bone metabolism by influencing calcium absorption. Gut microbiota can affect the absorption of skeletal development-related nutrients such as calcium and vitamin D. Calcium, the dominant mineral component in bone, is essential for bone health. Calcium absorption can be facilitated by vitamin D. Either dietary calcium deprivation or vitamin D deficiency may induce osteoporosis [84]. Sufficient calcium consumption can be a prophylactic measure against osteoporosis and relevant fracture [85]. A clinical study in adolescent girls showed decreased bone resorption in the presence of high calcium consumption (47.4 mmol/day compared to the recommended 22.5 mmol/day) [86]. Some studies showed that calcium metabolism differences among ethnic groups—in terms of dietary calcium intake, renal calcium excretion, and relevant regulatory hormone or factor—were associated with bone parameters related to osteoporosis/fracture risk [87]. In animal models, a low-calcium diet alone can lead to bone resorption, high bone turnover, and impaired bone trabecular microarchitecture in multiple bones, including the hard palate, mandible, vertebrae, femur, and proximal tibia [88–91]. Calcium is absorbed by the active transcellular pathway (ion pumps) or passive paracellular diffusion (ion channels), depending on the level of 1,25-(OH)2D (1,25-dihydroxy vitamin D) [92]. The proteins involved in the transcellular pathway consist of transient receptor potential vanilloid type 6 (TRPV6/CaT1/ECaC2), which absorbs calcium from the gut lumen into cells; calbindin-D9k, which is responsible for intracellular calcium transportation; and plasma membrane calcium-ATPase 1b (PMCA1b), which excretes calcium outside cells into the blood

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[93]. Passive paracellular calcium diffusion occurs as calcium (Ca2+) flux across the intestinal epithelium and is based on tight junction (TJ) proteins between intestinal epithelial cells [94]. Normal calcium intake rates in adults are approximately 30–35% [95, 96]; these levels can be increased by probiotics, prebiotics, and synbiotics consisting of probiotics and their favorable prebiotics [97]. Specific probiotic bacteria, such as Lactobacillus salivarius rather than Bifidobacterium infantis, stimulated calcium uptake by enterocytes in a Caco-2 cell culture model. [98] Oligosaccharides (NDO), the dietary prebiotics containing fructooligosaccharides (FOS) and inulin, significantly facilitated intestinal calcium absorption and increased skeletal calcium content in growing and adult rats [99– 102]. Prebiotic inulin produced an enhancement in calcium absorption compared to other oligosaccharides [99, 100], while the combination of both may act synergistically [101, 102]. Additionally, a study in healthy adolescent girls demonstrated that daily administration of GOS can increase calcium absorption [103]. Another clinical study reported the improvement of calcium absorption in young healthy women with long-term treatment with lactosucrose [104]. As the fermentation substrates of gut microbiota, prebiotics affect bone metabolism by producing a variety of beneficial metabolites, such as short-chain fatty acids (SCFA). The potential mechanism by which SCFA regulate bone metabolism involves direct effects on proteins associated with calcium absorption. Experiments both in vitro and in vivo using animal models showed that an SCFA supplement could increase the transcriptional levels of TRPV6 and calbindin-D9k rather than PMCA1b in cultured Caco-2 human colonic epithelium and rat colorectal mucosa [105, 106]. The TRPV6 gene was shown to contain a segment characterized by a positive response to SCFA [105]. In addition, the response of calbindinD9k to SCFA varied with time and SCFA dose [106]. The upregulation of calbindin-D9k by prebiotic diet specifically occurred in the colorectal segment regardless of dietary calcium uptake and

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serum 1,25-(OH)2D level, and it was related to the transcription factors vitamin D receptor (VDR) and cdx-2 [107–109]. The SCFA butyrate resulting from the prebiotic diet can upregulate VDR, activate the cdx-2 promoter, and facilitate cdx-2 mRNA expression [110]. Although direct evidence for the SCFA-related effect on intestinal paracellular calcium absorption is still absent, a ruminant model in which more than 50% of calcium absorption pre-intestinally occurs in the rumen manifested a dose-dependent promotion by SCFA on the ruminal calcium ion flux rate from mucosa to serum in the paracellular pathway [111, 112]. As stated above, both probiotics and prebiotics can influence intestinal epithelial permeability by regulating TJ protein expression and distribution, which possibly underlies the mechanism of prebiotic effects on paracellular calcium transport. In addition to direct action on the cellular structure involved in the calcium absorption process, prebiotics can also alter the intestinal microenvironment, thereby indirectly modulating bone metabolism. SCFA generated from prebiotics could lower the intestinal lumen pH and consequently inhibit the formation of calcium complexes, such as calcium phosphates, leading to increased calcium absorption [113].

10.1.3

Relationship Between the Intestinal Microbiota and PMO

10.1.3.1 PMO Animal Models Current data on the relationship between intestinal microbiota and PMO are primarily obtained from animal models. The most commonly used PMO animal models are rodents submitted to either surgery or medication. Ovariectomy is the most frequently used surgery to generate PMO rodent models. Bilateral ovariectomy is used to successfully set up morbid states of PMO in the proximal tibia, distal femur, and lumbar vertebra according to the guidelines for the preclinical and clinical evaluation of PMO medication issued by the US Food and Drug Administration (FDA) [114]. Gonadotropin-releasing hormone (GnRH) agonists are frequently used to induce PMO in

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rodents. The long-term or high-dose administration of GnRH agonists to rats typically housed under germ-free conditions [49] inhibits the secretion of endogenous GnRH, gonadotrophin, and estrogen [115, 116]. GnRH agonist-induced bone loss is reversible. Kurabayashi T et al. found that Sprague-Dawley (SD) rats submitted to long-term GnRH agonist treatment exhibited decreased bone mass, bone density, and bone turnover that could be partially recovered after treatment interruption [115]. Estrogen deficiency induced by either ovariectomy or GnRH agonist in murine models evidently increases bone turnover and bone loss and reduces bone mineral density and bone volume in lumbar vertebrae and long bones, thus recapitulating conditions in patients with PMO [114, 115, 117]. Animal age can affect the final experimental results, as preadolescent mice undergo rapid bone growth and high bone turnover due to the presence of growth hormones [118]. In addition, mice are likely to undergo irreversible aging symptoms [119] and potentially develop senile osteoporosis as early as 5–6 months old. Therefore, 8–20week-old rats or mice are usually used to establish PMO animal models [49, 115, 118, 120–128].

10.1.3.2 PMO Development Depends on the Intestinal Microbiota and Host Genetic Background The intestinal microbiota is indispensable to PMO development. Compared to conventionally raised (Con-R) mice, germ-free (GF) mice showed no significant alteration in either pro-inflammatory cytokines in bone marrow or femoral trabecular parameters after PMO model establishment by the administration of GnRH agonists [49]. However, similar to Con-R mice, GF mice colonized with a normal gut microbiota exhibited increased pro-inflammatory cytokines and impaired bone properties due to estrogen deficiency [49]. Accordingly, intestinal microorganisms are involved in estrogen deficiency-associated trabecular bone resorption. These microorganisms may be correlated with certain trabecular bone parameters. In particular, trabecular number (Tb.N) and trabecular spacing (Tb.Sp) are influenced by the intestinal

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microbiota, whereas trabecular thickness (Tb. Th) is not [49]. Bone resorption in PMO has also been shown to be closely related to genetic background (Fig. 10.2). Previous studies have shown that estrogen deficiency-induced bone loss varies remarkably among different mouse strains [124, 126, 127]. Genetic regulation can act on PMO bone loss through multiple mechanisms. Genetic background determines basal bone mass [1, 122] and the specific distribution of intestinal antigen-presenting cells (APCs) with different functions [129]. Intestinal APCs, especially dendritic cells (DCs), present pathogenic antigens from the gut microbiota and activate CD4+ T cells to produce pro-inflammatory cytokines such as tumor necrosis factor-α (TNF-α), which stimulates osteoclastogenesis and induces bone loss [130, 131]. In addition, host genetic background can shape the intestinal microbiota [20, 22, 23, 33, 132, 133], which can influence the development and activity of host immune systems [59, 134] and thus may indirectly regulate bone loss in PMO.

10.1.3.3 Probiotics Prevent Bone Loss in PMO Murine Models Bone loss in PMO murine models can be prevented by probiotics. Several studies have shown that bone resorption of femur and vertebra in OVX mice could be completely inhibited by the administration of probiotics such as Lactobacillus reuteri, LGG, and the commercial mixture VSL#3 [49, 118]. In addition, probiotics such as Bifidobacterium longum, Lactobacillus paracasei, and a mixture of Lactobacillus paracasei and Lactobacillus plantarum alleviated femoral bone loss and increased bone mineral density in OVX rats or mice [120, 121]. Furthermore, soy skim milk fermented by Lactobacillus paracasei subsp. paracasei NTU 101 (NTU 101F) and Lactobacillus plantarum NTU 102 (NTU 102F) mitigated bone loss and improved the trabecular microarchitecture in OVX mice [125]. The effects of probiotics on bone tissues depend on the systemic conditions of the host. McCabe LR et al. [123] showed that L. reuteri

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Fig. 10.2 Genetic background acts on PMO bone loss. Genetic regulation affects bone loss in PMO by shaping the gut microbiota and determining basal bone mass as well as the distribution of APCs

increased trabecular bone parameters of the femur and vertebra in healthy male mice (but not intact female mice), suggesting that estrogen level might affect the sensitivity of bone formation to L. reuteri in mice. L. reuteri may affect bone metabolism by activating the estrogen signaling pathway in male mice, whereas healthy adult female mice are impervious to L. reuteri due to sufficient estrogen. Notably, probiotics enhanced the trabecular bone parameters in intact female mice under inflammatory conditions after surgery [49, 135]. These results indicate that inflammatory pathways may be potential targets of probiotics to normalize bone homeostasis.

10.1.4

Host and Microbiota Interactions in the Pathogenesis and Treatment of PMO

Immune responses mediated by antigens from the intestinal microbiota play a central role in the pathogenesis of PMO. Under healthy conditions, interplays between the intestinal microbiota, the

intestinal epithelial barrier, and the host immune system maintain homeostasis, inhibiting the number of intestinal pathogens and maintaining musculoskeletal balance. If homeostasis is disturbed, intestinal pathogens intrude into the host through the epithelial barrier and provoke an immune response, ultimately promoting osteoclastic bone resorption and continual bone loss in PMO. Accordingly, probiotics ameliorate bone resorption and destruction by suppressing immune responses and restoring equilibrium between the intestinal microbiota and the host.

10.1.4.1 Intestinal Microbial Diversity in PMO Is Regulated by Estrogen and Probiotics A healthy state and sufficient estrogen levels maintain intestinal microbial diversity (Fig. 10.3a). Under these conditions, beneficial bacteria are predominant and stunt the growth of pathogenic species, preserving the stability of the intestinal microbiota composition. In postmenopausal women, the absence of estrogen alters intestinal microbial composition and structure, leading to decreased microbial diversity

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Fig. 10.3 Intestinal microbial diversity in PMO is regulated by estrogen and probiotics. Healthy status can maintain gut microbial diversity and beneficial bacteria, which can activate Tregs to sustain immune homeostasis that is resistant to pathogens (a). Estrogen deficiency

reduces gut microbial diversity and beneficial bacteria, while increased pathogens induce inflammation (b). Probiotics can prevent pathogens and increase gut microbial diversity by producing extracellular substances (c)

(Fig. 10.3b). Clinical surveys of males and postmenopausal females have shown significant correlations between biodiversity (or Clostridium abundance) in feces and urinary levels of estrogen (or estrogen metabolites) [136, 137]. Estrogen deficiency destroys intestinal microbial diversity, which is reflected as a reduction in Firmicutes populations, including Clostridium species [136–138]. Firmicutes bacteria, especially Clostridium species, possess immune-regulatory effects that boost the formation of regulatory T cells (Tregs) and enhance their function, sustaining immune homeostasis [139, 140]. Hence, estrogen deficiency undermines intestinal microbial diversity and reduces the abundance of intestinal bacteria that are conducive to immune homeostasis, consequently facilitating pathogen reproduction and initiating an immune response.

When used to treat PMO, probiotics improve intestinal microbial constitution and restore biodiversity. Probiotics halt pathogen growth and increase intestinal microbial diversity by synthesizing extracellular compounds (Fig. 10.3c). A study by Preidis GA et al. [141] showed that L. reuteri increased microbial diversity and homogeneity in the feces of mice by producing reuterin. Reuterin, an antibiotic compound, promotes oxidative stress in cells by inducing the modification of thiols on proteins or small molecules, which in turn suppress the growth of pathogens such as Bacteroides while increasing the presence of Clostridium species [118, 142]. Additionally, the Lactococcus lactis strain G50 prevented H2S-producing bacteria from growing, while strain H61 had an inhibitory effect on Staphylococcus in a mouse model of

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senile osteoporosis [119, 143]. However, it has not yet been demonstrated whether L. lactis has an equivalent role in PMO.

10.1.4.2 Intestinal Epithelial Barrier Function in PMO Is Regulated by Estrogen and Probiotics The intestinal epithelium is the first barrier to physically resist intestinal pathogens. This barrier not only absorbs water and nutrients but also limits the penetration of intestinal antigens. The ability of the barrier to function properly depends on transcellular and paracellular pathways. The fundamental paracellular pathway structure is the tight junction (TJ), the integrity, and selective permeability of which are of vital importance to intestinal epithelial barrier function. TJs are protein complexes consisting of claudin, occludin, and zo proteins, which together allow selective passage of ions and small molecules [144– 150]. TJ permeability can be represented by transepithelial electrical resistance (TER); higher TER usually indicates lower permeability [151, 152]. Both physiological and pathological stimuli can affect the production and distribution of TJ proteins, thereby modulating intestinal epithelial permeability. TJ proteins are mainly regulated by phosphorylation through protein kinase A (PKA), protein kinase C (PKC), protein kinase G (PKG), serine/threonine (Ser/Thr) kinases, Rho, mitogen-activated protein kinase (MAPK), phosphatidylinositol-3-kinase/Akt (PI3K/Akt), and myosin light chain kinase (MLCK) [144, 150]. Sufficient levels of estrogen activate the GTP-binding protein Ras and a series of kinases present in cytoplasm (Raf, MEK1/2 and ERK1/2) through estrogen receptors on the intestinal epithelium; they also maintain relatively high levels of occludin protein expression (Fig. 10.4a) [144, 153–155]. As a result of this paracellular pathway, the intestinal epithelial barrier exhibits increased TER and can prevent pathogen invasion. Estrogen deficiency weakens the effect of the aforementioned estrogen-associated pathway, leading to increased intestinal epithelial permeability [156]. Antigens from intestinal pathogens initiate inflammatory cascades across the

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epithelial barrier, leading to the production of pro-inflammatory cytokines such as tumor necrosis factor-α (TNF-α) and interferon-γ (IFN-γ). TNF-α and IFN-γ downregulate the TJ proteins occludin and zo-1 via Raf-MEK1/2-ERK1/2 or MLKs-MKK3/6-p38 in the MAPK pathway and further compromise the intestinal epithelial barrier [157]. In addition, the pro-inflammatory factor interleukin-17 (IL-17) can increase claudin-1 protein expression and reinforce the intestinal epithelial barrier through Ras-Raf-MEK1/2ERK1/2 in the MAPK pathway [158]. However, the positive action of IL-17 fails to completely compensate for the adverse effect of TNF-α and IFN-γ because TNF-α and IFN-γ may be central players in the immune responses elicited by intestinal bacteria. Hence, estrogen deficiency increases intestinal epithelial permeability (Fig. 10.4b), facilitating the intrusion of intestinal pathogens and provoking immune reactions and ultimately resulting in increased osteoclastic bone resorption and continual bone loss in PMO. When used to treat PMO, probiotics fortify the intestinal epithelial barrier to protect the host against intestinal pathogen invasion (Fig. 10.4c). Probiotics regulate the production and distribution of TJ proteins and reduce intestinal epithelial permeability by inducing changes in TJ-related gene expression. In vitro experiments have confirmed that L. plantarum can promote the production and rearrangement of claudin-1, occludin, and zo-1 proteins in the Caco-2 human colon adenocarcinoma cell line in a dose-dependent manner [159, 160]. Bifidobacteria infantis was found to increase zo-1 and occludin protein expression by inhibiting pro-inflammatory cytokines or through the secretion of polypeptide bioactive factors to augment Erk levels while decreasing p38 levels [161]. The probiotic mixture VSL#3 also promoted the expression and redistribution of occludin, zo-1, and claudin-1 proteins in a mouse model of acute colitis [162]. The potential mechanism for the probiotic regulation of TJ proteins probably involves SCFAs as fermentation products, especially butyrate, which could stimulate the reorganization of TJ proteins and promote TJ assembly by upregulating AMP-activated protein kinase

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Fig. 10.4 Intestinal epithelial barrier function in PMO is regulated by estrogen and probiotics. Sufficient estrogen can prompt the expression of TJ proteins through the Raf-MEK1/2-ERK1/2 pathway to enhance the gut epithelial barrier (a), while this active effect on TJ is weakened by estrogen deficiency (b). Under estrogen deficiency, pathogen-induced pro-inflammatory cytokines such as TNF-α and IFN-γ reduce the production of TJ proteins

through both the Raf-MEK1/2-ERK1/2 and MLKsMKK3/6-p38 pathways and compromise the gut epithelial barrier (b). The positive action of IL-17 on TJ proteins (thin green arrows in b) fails to completely compensate for the adverse effect of TNF-α and IFN-γ. Probiotics can enhance the gut epithelial barrier by regulating the production and distribution of TJ proteins and affecting the growth and movement of intestinal epithelial cells (c)

(AMPK) activity in the Caco-2 cell model, resulting in increased TER and an enhanced intestinal epithelial barrier [163]. In addition, probiotics affected the growth and movement of intestinal epithelial cells by altering gene expression related to protein synthesis, metabolism, cell adhesion, and apoptosis [162, 164]. L. reuteri substantially promoted intestinal epithelial cell migration and proliferation and increased intestinal crypt depth, ultimately improving the absorptive function of the intestinal epithelial barrier [141]. Both LGG and L. plantarum can stimulate the intestinal epithelium to produce physiological levels of reactive oxygen species (ROS), which act as a second messenger to activate the Erk/MAPK pathway and consequently lead to

intestinal epithelial proliferation [165, 166]. Probiotics also offer resistance against the toxic effects produced by intestinal pathogens on the intestinal epithelium. Bifidobacteria reduce the production of autophagy-related proteins and further prevent intestinal epithelial autophagy triggered by endotoxins from gram-negative bacteria [167].

10.1.4.3 Host Immune Responses in PMO Are Regulated by Estrogen and the Intestinal Microbiota The immune system is the final barrier to intestinal pathogen invasion and is also a critical target for PMO treatment. APCs in the intestinal lamina propria can be divided into dendritic cells (DCs)

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and macrophages [129]. Although all macrophages and DCs can induce Foxp3+ Treg cell differentiation, macrophages with a higher T cell/APC ratio are more efficient than DCs [129]. By contrast, DCs only partially induce Th17 cell differentiation [129]. Treg cells are a subset of immunocytes with inhibitory effects on the differentiation and function of Th1, Th2, and Th17 cells [130]. In addition, Treg cells can inhibit osteoclast formation by cell-to-cell contact via the cytotoxic T lymphocyte antigen (CTLA-4) or by secreting anti-inflammatory cytokines such as IL-4, IL-10, and transforming growth factor-β (TGF-β) [168–171]. Th17 cells, a subgroup of T cells, stimulate osteoclast formation and bone resorption by producing high levels of IL-17, RANKL, and TNF-α [172]. Both adequate estrogen levels and intestinal microbial diversity are needed to maintain immune homeostasis (Fig. 10.5a). Clostridium improves the aggregation, quantity, and function of Treg cells to create an environment abundant in TGF-β, which consequently prevents osteoclastogenesis [139]. Estrogen protects bone by downregulating immune responses and modulating osteoblast/osteoclast equilibrium [173]. Estrogen not only activates the apoptosis-promoting Fas/FasL pathway through direct interaction with osteoclasts [174–177] but also indirectly increases TGF-β production by Treg cells and decreases the production of TNF-a and RANKL by Th17 cells, ultimately promoting osteoclast apoptosis [131, 168, 169, 171, 178, 179]. Furthermore, estrogen exerts antiapoptotic effects on osteoblasts and osteocytes through the ERK pathway [177, 180]. Estrogen deficiency and reduced intestinal biodiversity have negative effects on bone (Fig. 10.5b). Pathogenic antigens cross the intestinal epithelium and trigger inflammatory immune responses that are mainly mediated by T cells. Estrogen deficiency boosts the antigen presentation of DCs and macrophages through multiple pathways. Upon estrogen depletion, ROS excessively accumulate in bone marrow cells [181, 182]. ROS enhance the antigenpresenting function of DCs, which further activates CD4+T cells to produce IFN-γ. The

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enhanced production of IFN-γ in turn improves the antigen-presenting ability of bone marrow macrophages (BMM) by upregulating MHC II molecules [183–187]. In addition, estrogen deficiency upregulates co-stimulator CD80 to activate bone marrow DCs [184]. Increased antigen presentation motivates CD4+ cells, including IL-17-producing Th17 cells, to mediate osteoclast formation and bone resorption [130, 188]. In addition to antigen-dependent activation, increased levels of IFN-γ and IL-7, in combination with low levels of TGF-β, indirectly activate T cells in bone marrow [130, 188, 189]. Activated T cells generate a considerable quantity of TNF-α, which acts as a key pathogenic factor in PMO development [131, 190–193]. TNF-α stimulates the production of RANKL and macrophage colony stimulatory factor (M-CSF); it also suppresses the production of osteoprotegerin (OPG) by inducing the expression of CD40L and the bone mass regulatory factor DLK1/FA-1 [130, 194, 195]. In addition, TNF-α acts either directly on osteoclast precursors to promote their maturation [196] or indirectly on TNF-α receptor p55 to augment M-CSF- and RANKL-induced osteoclastogenesis [131]. Furthermore, estrogen deficiency increases levels of Act1 adaptor protein on the surfaces of osteoblasts and subsequently activates the IL-17 signal pathway to promote bone resorption [197, 198]. These findings provide evidence that CD4+T cells (including Th17 cells) and the pro-inflammatory cytokine TNF-α are primary factors responsible for bone loss mediated by intestinal bacteria in PMO. When used for PMO treatment, probiotics also suppress bone resorption by regulating immune responses to intestinal microorganisms. Probiotics secrete small molecules to regulate the host immune response (Fig. 10.5c). Probiotics also produce SCFAs by utilizing prebiotics [30, 34, 199, 200]. SCFA receptors contain GPR41 and GPR43, the latter of which is mainly found in immunocytes such as neutrophils and monocytes [201]. SCFAs, especially butyric acid, interact with GPR43 to reduce levels of monocyte chemotactic protein-1 (MCP-1) and LPS-induced cytokines such as TNF-α and

deficiency reduces osteoblast formation; the invasion of pathogens activates CD4+T cells including TH17, which mainly produce TNF-α to promote osteoclastogenesis, leading to bone loss and microstructural destruction (b). Probiotics can regulate immune responses by secreting small molecules such as SCFAs and histamine (c)

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Fig. 10.5 Host immune responses in PMO are regulated by estrogen and intestinal microbiota. Both beneficial gut bacteria and sufficient estrogen activate Tregs, which produce TGF-β to prevent osteoclastogenesis and induce osteoclast apoptosis; estrogen prompts osteoblast formation to improve bone mass and structure (a). Estrogen

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IFN-γ. They also upregulate the expression of TGF-β1, IL-4, and IL-10, ultimately activating Treg cells [120, 201–205]. In addition, L. reuteri transforms dietary L-histidine to histamine, which inhibits the MEK1/2-ERK1/2 pathway via H2 receptors and further inhibits TNF-α production by monocytes [206]. Lactobacillus also impedes DC activation during inflammation and promotes Treg differentiation by inducing the expression of molecular ligands with inhibitory effects on pertinent DNA motifs [207].

10.1.4.4 The Intestinal Microbiota and Estrogen Orchestrate Calcium Absorption As described above, both calcium content and estrogen level are critical to bone metabolism. In postmenopausal-osteoporotic rats, combined deficiencies of dietary calcium and estrogen had a more adverse effect on bone mass and microstructure than either single deficiency, with more bone loss and more severely impaired bone properties [88, 90, 208]. Additionally, calcium balance can be regulated by estrogen. Under normal conditions, estrogen treatment can increase intestinal calcium absorption in rats [209]. Accumulating evidence suggests that estrogen deficiency could induce impaired calcium absorption, which was improved by estrogen supplementation [210–212]. The potential mechanisms of estrogen-associated regulation on calcium absorption are still disputed. Estrogen may indirectly promote vitamin D receptor (VDR) protein expression and enhance intestinal mucosal responsiveness to 1,25-(OH)2D, resulting in increased intestinal calcium absorption [213, 214]. However, estrogen deficiencyrelated calcium malabsorption may not depend on the serum 1,25-(OH)2D pathway. Estrogen reversed the reduced calcium absorption by directly interacting with estrogen receptor alpha (ER-α) on the intestine, upregulating the calcium transport protein 1 (CaT 1) of the calcium influx channel without significantly altering serum 1,25(OH)2D level. [212, 215, 216] In addition, estrogen deficiency increased the urinary fractional excretion of calcium (FECa) in OVX rats [120].

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The imbalance in calcium metabolism induced by estrogen deficiency was also redressed by the application of probiotics and prebiotics for the treatment of PMO [97]. Probiotic supplements completely inhibited the increase in FECa due to estrogen deficiency in OVX rats [120]. Oligosaccharides (NDO), dietary prebiotics such as fructooligosaccharides (FOS), galactooligosaccharides (GOS), and inulin, can significantly promote intestinal calcium absorption and skeletal calcium retention in OVX rats, resulting in suppressed bone loss [217, 218].

10.1.4.5 The Gut Microbiota Produces Estrogen-Like Metabolites with Regulatory Effects on Bone Metabolism Estrogen plays a major role in promoting osteogenesis. The role of estrogen is not limited to the direct suppression of osteoclast activity and lifespan, facilitation of osteoblast lifespan and differentiation, or reduction of mature osteoblasts apoptosis to promote osteogenesis. It also inhibits the formation of both osteoblasts and osteoclasts from bone marrow precursors to prevent bone remodeling and regulate bone turnover [69, 70]. In the absence of estrogen due to ovariectomy or post-menopause, estrogen-deficient women exhibit accelerated bone loss and increased bone turnover as well as impaired bone microarchitectural and mechanical properties [49, 177, 219]. Hormone replacement therapy (HRT), including supplementation with estrogen and progesterone, has been applied to postmenopausal women suffering from PMO and achieved favorable effects [220]. Instead of estrogen supplementation, the gut microbiota may act as another “endocrine organ” and potentiate novel access to replenish estrogen by utilizing exogenous nutrients and producing more estrogenic substances. Phytoestrogens, which are predominantly present in natural foods such as soy, are exogenous nutrients with structures and bioactivity similar to human intrinsic estrogens. Various metabolites produced from phytoestrogens by the gut microbiota, including equol, urolithins, and enterolignans, are characterized by higher

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bioavailability and respectively more estrogenic, antiestrogenic, and antioxidant bioactivities than their precursors in phytoestrogens, such as isoflavones, ellagitannins, and lignans [221]. Daidzein, the principle isoflavone in soy, has two metabolic patterns including equol and O-desmethylangolensin (O-DMA) production [222]. Equol shows much more estrogenic bioactivity or effects than O-DMA for bone metabolism in PMO [223]. Equol, which is mostly present as a glucuronide conjugate and binds to the estrogen receptor (ER), can suppress bone resorption, promote bone formation, and improve bone biomechanical and microstructural properties in subjects with PMO but has no impacts on bone in healthy early postmenopausal women [224–229]. The potential mechanism that involves equol may prevent osteoclast formation, stimulate the proliferation and differentiation of osteoblasts, and increase osteocalcin level by ER [223, 230]. Additionally, equol can inhibit the expression of relevant inflammatory cytokines in bone marrow in a dose-dependent fashion due to estrogen deficiency or LPS from intestinal pathogens [231–233]. Although produced by gut microbiota, equol may modify gut microbiota diversity and composition in turn [231]. Isoflavone metabolism can promote the growth of Clostridium clusters XIVa and IV and suppress the genera Bacteroides and Parabacteroides [234]. Nevertheless, equol production from dietary phytoestrogens has significant interpersonal variations, predominantly depending on gut microbial composition and potential correlations among the three groups of phytoestrogen metabolism as well as dietary components [77, 221, 235, 236]. At present, the key equol-producing gut bacteria have not yet been identified. Most studies target potential equol-producing bacteria by cultivation or sequence analysis of fecal samples. Two strains of Eubacterium sp. were isolated and considered the most likely equolproducing bacteria from pig feces [237]. Another intestinal bacteria, Slackia TM-30, a rod-shaped and gram-positive anaerobe isolated from healthy human feces, also proved to be highly related to equol production [78]. The sequence information

347

for fecal samples in postmenopausal women with dietary isoflavone uptake indicated obviously higher proportions of Eubacterium and Bifidobacterium in equol-producing subjects than in equol non-producers [238]. Other studies identified other bacteria that significantly increased in fecal samples of equol producers, including Collinsella, Asaccharobacter, Dorea, and Finegoldia [234, 239]. In terms of function, sulfate-reducing bacteria were suggested to be involved in equol production [235]. In addition to specific gut bacteria, equol-producing capacity may inversely correlate with O-DMA production [235]. In addition, daidzein bioavailability and the equol/O-DMA production ratio could be elevated by the combined administration of isoflavones and prebiotic oligosaccharides or probiotic bacteria such as Lactobacillus casei [240–242]. However, another study showed that the combination of soy isoflavones and fructo-oligosaccharides had no synergistic effects on bone mineral density or bone mineral content but effectively improved bone microstructural properties, including trabecular number, thickness, and separation [243]. Overall, the beneficial effects of phytoestrogen supplementation on PMO mainly depend on individual metabolisms involving both the appropriate gut microbiome and dietary composition [244].

10.1.5

Conclusion

Bone resorption in PMO is the consequence of interactions among the estrogen level, the intestinal microbiota, and the host immune system. When estrogen levels are deficient, bacteria and intestinal antigens cross the compromised intestinal epithelium barrier and initiate the immune responses associated with bone loss in PMO. Probiotics prevent bone resorption by restoring intestinal microbial diversity, enhancing the intestinal epithelial barrier, and normalizing aberrant host immune responses, as well as facilitating intestinal calcium absorption and the potential production of estrogen-like metabolites, as

348

X. Xu et al.

Table 10.1 Current probiotics with beneficial effects on estrogen deficiency-induced bone loss Probiotics Research models Lactobacillus spp. L. rhamnosus C57Bl/6 OVX mice GG C57Bl/6 OVX mice

L. reuteri

L. paracasei L. plantarum

C57Bl/6 OVX mice Balb/c OVX mice and healthy C57Bl/ 6 male mice or intact female mice with inflammation Outbred CD1 neonatal mice Outbred CD1 neonatal mice or Balb/c OVX mice C57Bl/6 OVX mice Murine and drosophila intestine or Caco-2 cell monolayers Caco-2 cell monolayers

Lactococcus SAMP6 mice lactis Bifidobacterium spp. B. longum OVX SD rat B. infantis Mixture L. paracasei and L. plantarum VSL#3a

Outcomes

References

Attenuates intestinal and BM inflammation and completely inhibits bone loss Reduces TJ destruction and gut epithelial permeability Affects enterocyte proliferation and migration Suppresses inflammation and bone loss in OVX mice and increases bone parameters in healthy male mice Increases enterocyte migration, proliferation, and crypt height Increases intestinal microbial diversity and evenness and inhibits growth of pathogens Decreases inflammatory cytokines and bone loss Induces enterocyte proliferation and modulates cellular processes, e.g., metabolism, adhesion and apoptosis Promotes production and rearrangement of TJ proteins and enhances TJ integrity Inhibits H2S-producing bacteria and Staphylococcus

[49] [49] [165] [118, 123, 135, 206] [141] [118, 141, 142] [120] [164, 166]

[159, 160] [119, 143]

Reduces bone loss and enhances bone mineral density Induces rearrangement of TJ proteins and normalizes gut permeability

[121]

C57Bl/6 OVX mice

Decreases inflammatory cytokines and bone loss

[120]

C57Bl/6 OVX mice

Attenuates intestinal and BM inflammation and completely inhibits bone loss Promotes expression and redistribution of TJ proteins and reduces intestinal epithelial permeability

[49]

IL-10-deficient mice

C57Bl/6 OVX mice or BALB/c mice in acute colitis model

[161]

[49, 162]

a The mixture VSL#3 contains Bifidobacterium breve, Bifidobacterium longum, Bifidobacterium infantis, Lactobacillus acidophilus, Lactobacillus plantarum, Lactobacillus paracasei, Lactobacillus bulgaricus, and Streptococcus thermophiles

summarized in Table 10.1. Hence, the intestinal microbiota serves as a key factor in the pathogenesis of PMO and will also serve as a new target in the treatment of PMO. The application of probiotics may be a promising adjuvant to current therapies. However, current studies on probiotics for PMO treatment are limited to animal studies. The translation from animal studies to clinical application faces many challenges, such as effective dosage and safety in humans. The safety and feasibility of

probiotics application in humans have been demonstrated by clinical studies in specific groups, such as in healthy infants [245], preterm infants, children with intractable diarrhea [246], and children and adolescents undergoing HCT [247]. However, in patients with predicted severe acute pancreatitis, significant increases in bowel ischemia and mortality were related to probiotic prophylaxis, as reported in the study by Besselink et al. [248] Hence, more studies are needed to validate the safety of probiotics and confirm the

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Intestinal Microbiota and Osteoporosis

optimal dosage and the proper time and method of delivery for probiotics in the context of PMO treatment. Acknowledgments We thank Dr. Geelsu Hwang from the University of Pennsylvania School of Dental Medicine for critical advice on the manuscript. This work was supported by the National Natural Science Foundation of China (grant numbers 81430011, 81470711, and 81670978) and the Brilliant Young Investigator Award, Sichuan University (grant number 2015SCU04A16). The content of this chapter was modified from a paper reported by our group in Bone Research (Xu X et al. 2018). The related contents are reused with permission. Disclosures The authors declare no conflicts of interest.

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Index

A Acellular microorganisms, 1, 3 Actinomyces, 28, 54, 74, 81 Adhesion, 12, 71–73, 171, 178, 207, 288, 326, 343 Aggregatibacter, 186, 227, 321, 326 Aspergillus, 282, 317, 319, 325, 326, 328 B Bacilli, 4, 15, 16, 36, 40, 81, 82, 88, 93, 97, 107, 146, 153, 154, 156, 167, 182, 291, 302 Bacterial capsule stain, 28–32 Bacteriophages, 4, 6, 18 Bacteroides, 154, 156, 159, 160, 168, 190, 201, 304, 321, 326, 327, 335, 341, 347 Bifidobacterium, 88 Binary fission, 15, 16 Biofilm, 64, 67, 71–72, 76, 136, 186, 235, 288, 325–327 C Campylobacter, 12, 32, 50, 177, 266 Candida, 33–35, 56, 63, 238, 243, 272, 276, 278, 280, 316, 319, 323, 325–328 Capnocytophaga, 55, 62, 159, 161, 165, 321, 326 Capsule, 6, 10, 22, 25, 28, 31, 32, 37, 98, 188, 217, 220, 227, 228, 230, 263, 283, 295 Cell membrane, 3, 6, 7, 9, 11, 236, 238 Cell wall, 6–7, 9–11, 13, 16, 26, 82, 100, 111, 120, 175, 202, 207, 211, 235, 238, 256 Chromosome, 2, 12, 18, 19, 21, 276 Chryseobacterium, 257 Cocci, 4, 15, 50, 51, 111, 112, 115, 126, 128, 139, 146, 153, 182, 212, 230, 253 Collection, 37–39, 68, 294, 295, 310, 328, 334 Colony forming units, 41, 68–70 Confocal laser scanning microscopy (CLSM), 57, 64, 66, 72, 291 Conjugation, 10, 19–23 Corynebacterium, 108, 110, 153, 256 Cytoplasm, 2, 6, 7, 9, 32, 238, 342 D Decline phase, 15, 66 Dental plaque, 38, 54, 64, 71, 82, 99, 101, 112, 119, 140, 153, 172, 186, 217, 227

Dilution, 38–40, 68, 294, 295 DNA, 3, 10, 13, 16–19, 21, 22, 65, 73, 74, 76, 79, 82, 85, 87, 90, 97–99, 107, 108, 112, 139, 146, 149, 154, 159, 161, 168, 193, 205, 240, 256, 260, 296, 311, 328 E Eikenella, 29, 168 Elizabethkingia, 263 Enterococcus, 146, 306 Escherichia, 4, 8, 11, 17, 20, 23, 224, 266, 309, 334 Eubacterium, 146, 149, 347 Eukaryote, 1 F Flagellar stain, 32–33 Flagellum, 6, 11, 12, 25, 32, 37, 256, 267, 296, 326 Fluctuation test, 19, 20 Fungal Structures Stain, 32–35 Fusobacterium, 54, 164, 172, 303, 306, 309 G Giemsa stain, 33, 37, 207, 236 Gram-negative, 6, 7, 9, 12, 21, 27, 29, 39, 40, 50, 51, 72, 83, 108, 110, 134, 138, 139, 153–155, 217, 224–236, 256, 267, 295, 343 Gram-positive, 6, 8, 12, 13, 16, 17, 21, 27, 28, 50, 51, 72, 81, 138, 141, 146, 173, 211–225, 230, 238, 253, 337, 347 Gram stain, 6, 9, 25–27, 32, 33, 43, 72, 83, 86, 88, 91, 94, 96, 99, 105, 109, 112, 114, 117, 121, 129, 134, 141, 149, 151, 154, 157, 163, 166, 172, 176, 181, 184, 187, 190, 193, 198, 214, 220, 226, 228, 231, 233, 236, 240, 254, 256, 263, 268, 270, 272 Growth curves, 15, 66–68 H Haemophilus, 19, 171, 186, 227, 303 Helicobacter, 176, 177 Heredity, 10, 13, 15, 17 I Identification, 25, 26, 32, 37–47, 78, 185, 220, 276, 280, 292, 295, 296

# Zhejiang University Press 2020 X. Zhou, Y. Li (eds.), Atlas of Oral Microbiology: From Healthy Microflora to Disease, https://doi.org/10.1007/978-981-15-7899-1

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360 Incubation, 20, 37–47, 68, 71, 82, 108, 124, 192, 199, 211, 238, 276, 295 Inoculation, 67, 68, 153, 294, 295 Intestinal microbiota, 306, 308, 309, 333–349 Isolation, 37–47, 119, 235, 253, 256, 257 K Klebsiella, 263 L Lactobacillus, 61, 98, 306, 327, 335, 336, 338, 339, 346–348 Lag phase, 15, 66 Leptotrichia, 134, 140 Leuconostoc, 253 Lichen planus, 293, 316–330 Logarithmic phase, 15, 66 M Moraxella, 230, 306 Mycobiome, 293, 316–330 Mycoplasma, 3, 18, 26, 29, 33, 72, 235–237 N Negative Congo red stain, 33–37 Neisseria, 230 Next-generation sequencing (NGS), 74, 316 Nuclear material, 6, 10 O Oral microbiome, 72–79, 288, 289, 294, 316, 325, 327 Osteoporosis, 334–347 P Peptidoglycan, 6, 7, 26, 111, 120, 202, 207 Peptostreptococcus, 149 Pilus, 6, 12, 21 Porphyromonas, 29, 39, 46, 51, 62, 154, 201, 205, 289, 303, 306, 309, 310, 317, 323, 327 Prevotella, 39, 55, 154, 190, 327 Prokaryote, 1, 15 Propionibacterium, 153 Protozoan Smears, 37

Index R Replica plating, 19, 21 RNA, 3, 9, 13, 17, 65 Rothia, 61 S Saccharomyces, 236–249, 326 Sanger sequencing, 73, 77–79 Scanning electron microscopy (SEM), 2, 17, 47, 60–63, 65, 72, 75, 115, 118, 122, 125, 129, 134, 139, 141, 146, 147, 150, 152, 155, 157, 159, 162, 164, 166, 169, 173, 177, 182, 184, 187, 191, 194, 196, 198, 200, 203, 206, 207, 209, 212, 217, 219, 221, 223, 226, 228, 231, 233, 238, 239, 254, 257, 259, 261, 264, 268, 270, 273, 275, 277, 279, 282, 284 Spirochete, 4, 18, 25, 31, 36, 39, 72, 205, 209, 295 Spores, 4–6, 13, 14, 25, 27–28, 30, 32, 82, 83, 92, 98, 106, 107, 128, 136, 146, 154, 155, 158, 159, 167, 168, 170, 172, 188, 201, 205, 206, 227, 228, 231, 238, 247, 263, 267, 282, 285, 295, 296 Staining bacterial spores stain, 27–30 Staphylococcus, 4, 7, 21, 38, 45, 111, 341 Stationary phase, 15, 66 Stenotrophomonas, 256 Stereomicroscopy, 47–57 Streptococcus, 6, 17, 19, 21, 22, 29, 50, 51, 56, 59, 67, 76, 112, 146, 214–225, 288, 291, 303, 306, 309, 316, 322, 348 Subgingival, 37, 38, 41, 58, 70, 88, 111, 146 Supragingival, 81–142, 186 Suspension, 32, 38–39, 71, 310 T Transduction, 10, 19, 21, 23 Transformation, 19, 309, 316 Transmission electron microscopy (TEM), 53–65, 290, 291 Transportation, 37–38, 68, 337 Treponema, 42, 205, 208, 295, 321 V Variations, 13, 15, 16, 19, 73, 282, 325, 347 Veillonella, 139, 272, 303, 321 Vibrio, 4, 12 Virus, 3, 4, 13, 18, 72, 240–250