The Reconstruction and Analysis of Oral Microbiome Composition Using Dental Calculus from the Mississippi State Asylum (1855-1935), Jackson, Ms

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Jonathan Robert Belanich

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The reconstruction and analysis of oral microbiome composition using dental calculus

from the Mississippi State Asylum (1855-1935), Jackson, MS

By TITLE PAGE Jonathan Robert Belanich

A Thesis Submitted to the Faculty of Mississippi State University in Partial Fulfillment of the Requirements for the Degree of Master of Arts in Applied Anthropology in the Department of Anthropology and Middle Eastern Cultures

Mississippi State, Mississippi

August 2016

2016

The reconstruction and analysis of oral microbiome composition using dental calculus

from the Mississippi State Asylum (1855-1935), Jackson, MS

By APPROVAL PAGE Jonathan Robert Belanich

Approved:

______Molly K. Zuckerman (Major Professor)

______Heather Jordan (Committee Member)

______Darcy Shane Miller (Committee Member)

______Nicholas P. Herrmann (Committee Member)

______David M. Hoffman (Graduate Coordinator)

______Rick Travis Interim Dean College of Arts & Sciences

Name: Jonathan Robert Belanich ABSTRACT Date of Degree: August 12, 2016

Institution: Mississippi State University

Major Field: Applied Anthropology

Major Professor: Dr. Molly K. Zuckerman

Title of Study: The reconstruction and analysis of oral microbiome composition using dental calculus from the Mississippi State Asylum (1855-1935), Jackson, MS

Pages in Study: 82

Candidate for Degree of Master of Arts

The human oral microbiome is the total amount of microbial biodiversity present

in the oral cavity and, given its relevance to human health and disease, has recently

become a foci for study. By analyzing dental calculus, and sequencing the bacterial DNA,

it is possible to reconstruct and examine the oral microbiomes of past individuals. In this

study, dental calculus was sampled from (N=4) skeletons recovered from the cemetery of

the mid 19th- 20th, century Mississippi State Asylum in Jackson, MS. Bacterial DNA

isolation and shotgun sequencing were successful, with 16S analyses yielding an average

of 96 identified species. All samples were significantly different from each other at all

taxonomic levels (p <0.0001). Targeted examinations for opportunistically pathogenic

oral bacteria were performed, but no detectable bacterial DNA was found in the samples.

This study is the first to reconstruct the oral microbiomes of a subsample of an historic

institutionalized population.

DEDICATION

This is dedicated to my fountain pens. It is additionally dedicated to my parents,

John and Donna Belanich, who were incredibly supportive throughout my undergraduate years, and remain so during my ever-increasing graduate career.

ACKNOWLEDGEMENTS

The author would like to specifically acknowledge Drs. Molly Zuckerman

(Department of Anthropology and Middle Eastern Cultures (AMEC)) and Heather Jordan

(Department of Biological Sciences) for the edits of previous drafts of the thesis proposal, and thesis, as well as Drs. Nicholas Herrmann (Texas State University) and Shane Miller

(AMEC), the additional members of this thesis committee. Thanks are also extended to

Dr. Jason Rosch (Infectious Disease Department, St. Jude) for the sequencing of the

DNA samples. Additional thanks to Amanda Lawrence (Institute for Imaging &

Analytical Technologies) and Dr. Gerald Baker (Department of Biochemistry, Molecular

Biology, Entomology, & Plant Pathology) for their assistance with the Scanning Electron

Microscopy and imaging, and Dr. Christopher Lynn and the HBERG writing group at

The University of Alabama for the comments on previous drafts of this thesis.

Additionally, Jesseca A. Paulsen is thanked for her support during the process of writing and revising.

iii

TABLE OF CONTENTS

DEDICATION ...... ii

ACKNOWLEDGEMENTS ...... iii

LIST OF TABLES ...... vi

LIST OF FIGURES ...... vii

CHAPTER

I. INTRODUCTION ...... 1

Research Questions ...... 4 Significance of Research ...... 5

II. BACKGROUND INFORMATION ...... 7

The Human Microbiome ...... 8 Bacteria and the Oral Microbiome ...... 8 Biofilms ...... 10 Dental Calculus Formation ...... 11 Ancient microbial DNA ...... 11 History of the MSA ...... 13 Individual Burials ...... 14 The Human Oral Microbiome Database ...... 15

III. MATERIALS AND METHODS ...... 16

Hypotheses ...... 16 Hypothesis 1: ...... 16 Hypothesis 2: ...... 17 Hypothesis 3: ...... 17 Hypothesis 4: ...... 18 Hypothesis 5: ...... 18 Sampling methods ...... 19 Scanning Electron Microscopy Sampling ...... 20 Microbial DNA Preparation ...... 21 DNA Isolation Kits ...... 21 Targeted Polymerase Chain Reaction and gel electrophoresis ...... 22 iv

Metagenomic Sequencing ...... 23 Illumina BaseSpace Analysis ...... 23 Scanning Electron Microscopy ...... 23 Human Oral Microbiome Database Biodiversity ...... 24 Statistical Analyses ...... 25

IV. RESULTS ...... 26

DNA Isolation Results ...... 26 16S Metagenomic Analysis ...... 27 Burials ...... 28 Negative Control ...... 28 Scanning Electron Microscopy ...... 29 Chi-Square Test of Independence ...... 34 Microbes in Burials ...... 35 Specific Pathogenic Bacteria ...... 35 Targeted Polymerase Chain Reaction ...... 35 Phyla Comparison against Modern HOMD ...... 36 Environmental Contaminants ...... 38

V. DISCUSSION AND CONCLUSION...... 40

Future Work ...... 45 Conclusion ...... 46

REFERENCES ...... 48

APPENDIX

A. GENUS AND SPECIES LISTS FOR SEQUENCED DENTAL CALCULUS ...... 55

B. HIGH-RESOLUTION TAXONOMIC CHARACTERIZATIONS OF THE RECONSTRUCTED ORAL MICROBIOMES ...... 76

LIST OF TABLES

2.1 Summary of Osteoware Burial Records ...... 15

3.1 Summation of Dental calculus sample removed for DNA isolation ...... 20

3.2 Summation of Dental calculus samples removed for SEM ...... 21

3.3 Table of bacterial specific primers for targeted PCR...... 22

3.4 Scanning Electron Microscopy Dental Calculus Samples ...... 24

4.1 Concentration of microbial DNA (ng/µL) in isolated samples ...... 27

4.2 16S Metagenomics Aggregate Results ...... 27

4.3 Natural environments of potential bacterial contaminants ...... 39

A.1 Burial 1 Genus and species list ...... 56

A.2 Burial 3 Genus and species list ...... 58

A.3 Burial 4 Genus and species list ...... 64

A.4 Burial 24 Genus and species list ...... 69

A.5 Negative control Genus and species list ...... 75

LIST OF FIGURES

1.1 Example of dental calculus present on the lingual side of a 3rd mandibular molar...... 4

4.1 Photo of bacteria on the surface of dental calculus ...... 30

4.2 Photo of bacteria on dental calculus ...... 31

4.3 High magnification photo of bacteria on dental calculus ...... 32

4.4 Photo of fungal hyphae on dental calculus ...... 33

4.5 Photo of mineral element in dental calculus ...... 34

4.6 Phyla comparison of the four individual burials and the data acquired from the HOMD...... 37

4.7 Phyla Dendrogram ...... 38

B.1 Kingdom Comparison ...... 77

B.2 Phyla Comparison ...... 78

B.3 Class Comparison ...... 79

B.4 Order Comparison ...... 80

B.5 Family Comparison ...... 81

B.6 Genus Comparison ...... 82

vii

CHAPTER I

INTRODUCTION

Analyses of human oral cavities and dental pathologies are useful tools that can

reveal substantial amounts of information about health (DeWitte and Bekvalac 2010),

diet (Warinner et al. 2014a), and diseases present in individuals and their communities

within the archaeological record. These have become foci for recent bioarchaeological

and paleopathological research, since the oral cavity, and the teeth in particular, are

constantly exposed to ingested food and are directly affected by nutrition. This has

resulted in a large body of literature within bioarchaeology about teeth and the oral

cavity. Typically this literature has focused on reconstructing diet (Buckley et al. 2014;

Lustmann et al. 1976), exploring the relationship between diet and disease (Hillson 1979;

Lukacs 2012), and examining evidence of antemortem stress (Armelagos et al. 2009;

Goodman et al. 1980; Hutchinson and Larsen 1988), and tooth wear (Eshed et al. 2006).

Recently, novel research methods involving the use of dental calculus have been implemented, and results from these studies demonstrate that unique chemical and

biological information can be gleaned from analysis of preserved human dental calculus.

Dental calculus (Figure 1.1) is the calcified form of dental plaque, a naturally occurring biofilm that forms on surfaces of teeth due to the growth and proliferation of the oral bacteria (White 1997; Zijnge et al. 2010).

Within the last ten years, modern microbial sequencing techniques have been applied to samples of dental calculus, which has enabled analysis and characterization of ancient oral microbiomes. The human oral microbiome is the ecological community of microorganisms that exist in the oral cavity during life (Dewhirst et al. 2010; Lederberg

and McCray 2001). These microorganisms comprise one of the most diverse subsections

of the human microbiome (Aas et al. 2005; Chen et al. 2010; Wade 2013). The human

microbiome is the total amount of bacterial diversity that is present in, and on the human

body (Turnbaugh et al. 2007). These bacteria have direct effects on human health,

digestion, and immunity (Bengmark 1999).

Through 16S and metagenomic sequencing techniques, oral microbiomes from

skeletal material dating to as early as 12,000 BCE and as recently as the 19th century CE have been examined for overall microbial diversity as well as more targeted investigation of dietary composition (Adler et al. 2013; Warinner et al. 2014a; Warinner et al. 2014b).

In contrast to this time depth, a literature review reveals that no published works have

analyzed more recent historical dental calculus and associated oral microbiomes.

However, this time period—c. the 19th to the mid 20th century—is integral for

understanding how the human oral microbiome has altered in composition over time,

with particular relevance to the major economic, behavioral, and subsistence shifts of the

19th and 20th centuries, specifically, industrialization, urbanization, and the emergence of

western diets. Subsistence shifts indirectly altered the microbiome; the change in ingested

diet affected the metabolizable nutrients that the oral bacteria can utilize, therefore

changing the oral environment.

The data gathered from this study allowed for the examination of oral microbiomes with relation to shifts in biodiversity over time. This work will add to the body of knowledge concerning the importance of the human microbiome. Additionally, other analyses have used dental calculus recovered from various archaeological contexts, including cemeteries, but no samples from known populations of institutionalized persons have been sequenced or analyzed. This will allow for the examination of a potentially unique subset of the human population.

Samples of preserved dental calculus (N=24) were removed from individuals recovered from the cemetery of the Mississippi State Asylum (MSA) (1855 – 1935 CE), a mental institution in Jackson, MS. DNA was isolated from the dental calculus samples, and the samples with a detectable amount of microbial DNA (N=4) were then sequenced and analyzed. During active years, this asylum housed and treated nearly 35,000 individuals diagnosed with a range of mental conditions, along with infectious, chronic, and degenerative diseases. The four individuals that were sequenced in this study date to the 1920’s based on dendrochronology dates.

Living conditions in the MSA were variable, but poor overall; admission and death records from the asylum indicate that pellagra, tuberculosis, epilepsy, and maniacal exhaustion were the leading causes of death (MSA records). Pellagra itself is of interest, since it is caused by a niacin deficiency, and is indicative of a high corn and low protein diet (Brenton 2000; Etheridge 1972). This analysis reconstructs a subset of historic oral microbiomes in order to investigate their place in the changing biodiversity of the human oral microbiome. This allows for the examination of the evolution of the oral microbiome, with relevance to pathogenicity, virulence, and antibiotic resistance, as well

as its connection to oral health, and health histories. Therefore, the primary research question for this project is “What was the total microbial composition of the oral

microbiome of marginalized individuals who lived and died in an institutionalized

environment in the early 20th century mid-South?”

Figure 1.1 Example of dental calculus present on the lingual side of a 3rd mandibular molar.

Research Questions

To address this primary question, several secondary research questions were

asked. (1) What is the level of microbial diversity between individuals in the sequenced

sample (N=4)? (2) What is the amount and diversity of opportunistically pathogenic oral

bacteria such as Streptococcus mutans or Porphyromonas gingivalis in these samples? (3)

Given the high prevalence of acute and chronic infectious conditions in the MSA, are

there known non-commensal pathogenic bacteria in these samples such as Streptococcus

pneumoniae or Mycobacterium tuberculosis? And (4) Does the microbial composition of these samples differ from the composition of modern oral microbiomes? To address these 4

questions, high-resolution taxonomic characterizations (species-level) were performed on

microbial DNA extracted from the samples of sequenced dental calculus (N=4). The

Human Oral Microbiome Database (HOMD) served as a modern comparison point to

examine if these reconstructed oral microbiomes were similar, or different from those of

modern individuals (Chen et al. 2010; Dewhirst et al. 2010). The HOMD is a

collaborative effort between scientists to provide information on the total biodiversity of

bacterial species present in the modern oral microbiome.

Significance of Research

Based on a review of the published literature, this research is the first to examine

the oral microbiome composition of individuals belonging to a marginalized and

institutionalized population. In addition, this is the first study to have performed a

taxonomic reconstruction of an historic skeletal sample from the past century, providing a

novel glimpse into the recent evolutionary history of the oral microbiome. These samples

of calculus provide novel information on the oral microbiome of individuals living in situations with dietary and nutritional insufficiencies. This research further demonstrates

the applicability of dental calculus studies in aiding research into how health and disease

are affected by diet, by directly analyzing the oral microbiome which is modified and

affected by dietary composition. Continuing research into the metagenome of sequenced

dental calculus will provide evidence for the evolution of pathogenesis and antimicrobial

resistance of the oral microbiome. This project, and future research, provides small, but useful datasets that can be used to examine the coexistence of humans and their bacteria.

Furthermore, a review of the published literature indicates that this is the first

study examining dental calculus conducted in a non-ancient DNA dedicated laboratory. 5

Although strict, and proper, contamination controls are still required and were employed, the low level of bacterial contamination detected in this study indicates that this type of

research need not be restricted to aDNA laboratories. Hopefully, this will allow for a proliferation of research examining ancient oral microbiomes.

CHAPTER II

BACKGROUND INFORMATION

The human microbiome, and in particular the oral microbiome, is quite diverse,

and its composition has great relevance to health and disease. The microbiome is an

important adaptation within human evolution. These microbes maintain the homeostasis

of the oral microbiome, defend against pathogenic bacteria, and play a key role in the

creation and regulation of human immune system (Bengmark 1999; Maslowski and

Mackay 2011; Rook and Brunet 2005). With the advent of high-throughput DNA sequencing (Sanger et al. 1977) and metagenomic sequencing, research began to examine

entire communities of bacteria at once. Utilizing this methodology, microbiome composition and biodiversity can be established down to the species level, and the

abundance of each taxon can be quantified. Additionally, specific bacterial genes can be

targeted and individually analyzed. This type of research is now relevant to the fields of

bioarchaeology and paleopathology, as the oral microbiomes of archaic populations can

be recreated and examined through the analysis of dental calculus. The recovered

microbial data allows for inferences about past diets and diseases to be made (Adler et al.

2013; Warinner et al. 2014a; Warinner et al. 2014b). By examining modern and ancient

human microbiomes, biologists and anthropologists can examine differences between

groups and explore how diet affects our health, and show how changes in subsistence

patterns made thousands of years ago can affect us today (Adler et al. 2013). Through

these analyses, researchers gain a clearer picture of how complex our bodies and immune systems really are.

The Human Microbiome

The human microbiome is composed of the microbiota present in gastrointestinal tracts, the oral cavity, the lungs, the vaginal cavity, and on the skin (Human Microbiome

Project 2012; Lazarevic et al. 2010; Nasidze et al. 2009; Ravel et al. 2011; Sommer et al.

2009; Turnbaugh et al. 2007). Together, these microbiomes comprise up to 3% of total body mass, and can outnumber human cells almost 10 to 1 (Human Microbiome Project

2012). These microbiomes are necessary for the proper functioning of the human digestive tract (De Filippo et al. 2010; Faith et al. 2013; Rook and Brunet 2005), and additionally form an integral part of the immune system by preventing infection by pathogenic bacteria (Tosh and McDonald 2011; Wade 2013).

Bacteria and the Oral Microbiome

The oral microbiome is one of the more, if not most, biologically diverse

microbiomes in the human body, and it is heavily colonized by different taxa of

microorganisms (Aas et al. 2005; Bik et al. 2010; Dewhirst et al. 2010; Wade 2013).

There are somewhere between 500 and 800 (Aas et al. 2005; Chen et al. 2010; Dewhirst

et al. 2010; Zarco et al. 2012) separate species of bacteria present in a given oral

microbiome at any one time. The oral microbiota are maintained through the constant supply of nutrients available from ingested food. In modern oral microbiomes, the major phyla that are represented are Firmicutes, Tenericutes, Actinobacteria, Fusobacteria,

Bacteroidetes, Proteobacteria and Spirochaetes (Chen et al. 2010; Dewhirst et al. 2010;

Wade 2013). The bacteria that are normally found in the oral cavity are mostly

commensal, and play an important role in the maintenance of health (Wade 2013). The

colonization of the oral cavity by commensal bacteria prevents external, potentially

pathogenic bacteria from establishing a presence. When disruptions occur to the normal

oral flora, opportunistic pathogens can quickly colonize, and lead to infection (Dewhirst

et al. 2010; Wade 2013). Therefore, the normal commensal diversity of this microbiome plays a role in maintaining health and preventing disease.

The oral cavity is not only diverse in biological assortment, but in microbial habitats as well. There are several distinct microenvironments present, such as the surface of the teeth, the gingival sulcus, the supragingival and subgingival areas, the hard and soft palates, the tongue, the cheeks, and the lips (Dewhirst et al. 2010; Zarco et al. 2012).

Although the mouth forms a distinct microbiome, the oral cavity is contiguous with several other microbial areas in the head and upper thoracic cavity such as the tonsils, the pharynx, the esophagus, the trachea, the nasal passages, and the sinuses (Dewhirst et al.

2010). Each distinct environment forms its own niche, and the microorganisms present in

the oral cavity differentially colonize surfaces (Aas et al. 2005). The differences in both

the substrate and the bacterial adhesins cause each microenvironment in the oral cavity to

harbor different types of bacteria (Gibbons 1989; Zarco et al. 2012). The adhesins bind to

both saccharide receptors, as well as acidic proline-rich proteins (PRPs), which are

present on the surface of the teeth. These binding sites on the proteins assist in the

attachment of biofilms and plaque forming bacteria (Gibbons 1989).

Biofilms

Biofilms are groups of bacteria that stick together to form a colony on a living surface (Hall-Stoodley et al. 2004). By binding together through the formation of an

extracellular matrix of extracellular polymeric substances (EPS) (Flemming and

Wingender 2010), communities of bacteria within biofilms not only increase their

defense against antibiotics, invasive bacteria, phagocytes, and changes in pH levels, but

against physical cleaning processes as well (Jefferson 2004). The biofilm also aids in the

creation of a favorable habitat for the bacteria, and potentially allows for additional

bacterial growth, and the transport of metabolizable products (Flemming and Wingender

2010). Within the oral cavity, the swishing and washing action of saliva is unable to shear

biofilms from their substrates, whereas planktonic bacteria are washed down the

esophagus and dissolved in the stomach acid (Gibbons 1989; Jefferson 2004). This

confers an evolutionary advantage to bacteria that are able to colonize oral surfaces and

form biofilms.

Biofilms that form on the surface of the teeth are known as dental plaque, and are

naturally found in all humans, although to varying degrees (Gibbons 1989; Rosan and

Lamont 2000). The process starts by the binding of adhesins to PRPs on the surface of

the teeth, which allows for the first set of bacteria to start a colony (Gibbons 1989). After

bacterial replication and growth, secondary colonization occurs through inter-bacterial

adhesion and co-aggregation (Rosan and Lamont 2000). This creates irreversible changes

in the surface of the microbes, effectively binding them together, which creates a biofilm.

Over time, bacteria in the mouth bind to the dental plaque, and effectively become stuck

as a semi-permanent feature. These bacteria then form commensal relationships and

extracellular matrices for the transport of nutrients and molecules are built (Rosan and

Lamont 2000).

Dental Calculus Formation

Over time, as the plaque becomes denser it comes into contact with more and

more minerals, such as calcium phosphate, through interaction with the tongue and

saliva. By constant exposure to these minerals, the dental plaque becomes calcified and

the bacteria are eventually locked into a matrix, which is similar to bone (Adler et al.

2013; Warinner et al. 2015). This is a continual process; when the dental calculus has

solidified, more bacteria colonize the surface and start over again as a sort of biological

palimpsest. In this manner, the bacteria that are present are preserved in the dental plaque

once it has calcified into dental calculus. Dental calculus is one of the only ways that

preserved bacteria can be retrieved from historic and archaic humans (Adler et al. 2013).

Due to their density and composition, teeth tend to be relatively well preserved in the

archaeological record (Adler et al. 2013; Devault et al. 2014; Hillson 1996; Hillson

2005). Therefore, if dental calculus is present, it tends to preserve as well. This

combination of archaeological and biological preservation makes teeth a useful avenue for bioarchaeological research examining oral microbiomes, dietary composition, and oral health in the past.

Ancient microbial DNA

Through the sequencing and analysis of the bacterial DNA that are trapped in the

dental calculus, it is possible to reconstruct both modern and ancient oral microbiomes.

16S and metagenomic sequencing have allowed researchers to examine ancient oral

microbiome composition. Recent studies examining dental calculus are present at the forefront of an exciting new frontier in bioarchaeological and paleopathological research

(Warinner et al. 2015). Dental calculus analysis has already allowed for the examination of dietary shifts (Adler et al. 2013), targeted examination of pathogens (Warinner et al.

2014b), and examination of dietary composition (Warinner et al. 2014a).

In 2013, a study was published by Adler et al. that examined and sequenced the

oral microbiome of prehistoric, historic, and modern samples, and compared against each

other. This was performed in order to examine the differences in oral microbiome

composition from before and after major subsistence pattern changes, such as the

Neolithic and Industrial revolutions. They used individuals from the Mesolithic in

northern Europe to the Late Medieval period in England. Adler et al. (2013) were able to

see large taxonomic shifts in the composition of the oral microbiome that correlated with

a temporal transect. Effectively, they found that, within their samples, the overall diversity of the oral microbiome decreased over time, and the prevalence of pathogenic bacteria increased. Their samples of modern day, oral microbiomes were the least phylogenetically diverse, and contained high levels of caries-causing bacteria. The samples that were from after the Industrial revolution were found to have the lowest amount of biodiversity of all the samples.

Warinner et al. (2014b) analyzed dental calculus and focused specifically on reconstructing the pathogenic components of the oral microbiome at the genus and species level. Their samples of dental calculus came from four adult skeletons with periodontal disease from Germany dated to 950-1200 CE, and from nine clinical samples from modern individuals with known dental histories. They were able to successfully

reconstruct the oral microbiome and accurately show the diseased state of the historic samples, and the authors found that there was a particularly high abundance of cariogenic, periodontal and other opportunistic pathogens in the diseased samples. From these samples Warinner et al. (2014b) also attempted to examine virulence and antibiotic resistance, and although they were able to detect proteomic evidence of the mechanisms for horizontal gene transfer and low-level antibiotic resistance, they were unable to demonstrate the degree at which it occurred, or if it even occurred at all. This demonstrates that metagenomic research, as well as 16S can be performed on dental calculus.

Also in 2014, Warinner et al. examined dental calculus for a specific biomarker that is involved in dairy consumption to determine direct evidence of milk consumption.

They used dental calculus samples from northern and central Europe, as well as southwest Asia, that dated from the Bronze though the 19th century. The authors were

able to determine the presence of the biomarker in dental calculus, and were able to

positively identify it in specimens that dated to the Bronze Age (Warinner et al. 2014a).

They also examined a Norse colony from Greenland, and found that the biomarker

decreased until the ultimate abandonment of the colony in the 15th century. They believe

that the diminishing presence of the biomarker was evidence that one of their major food

sources was collapsing. They were able to not only reconstruct diet, but to answer

specific questions on how specific food resources affected individuals in the past.

History of the MSA

The Mississippi State Asylum was located in Jackson MS, and opened in 1855 for

individuals who had chronic disabilities, or diseases. The conditions were inadequate; the 13

asylum was overcrowded, there were epidemics of yellow fever, tuberculosis, and influenza, as well as poor nutrition as evidenced by the high prevalence of death by pellagra (Goldberger et al. 1915). The average length of institutionalization in the MSA

was thirteen months. Additionally, from work conducted at the MSA, Goldberger (1914)

noted that asylum populations were fed cornmeal and canned foods, but were lacking any fresh foods or meat.

The diet from the Mount Vernon Hospital for the insane, in Tuscaloosa, Al, provides a glimpse into the diet of institutionalized individuals from the same era (Searcy

1907). Patients were primarily fed spoiled cornmeal in the form of grits and cornbread.

From these sources and records, we can infer that the diet of the individuals

institutionalized at the MSA was not diverse: their diets consisted primarily of corn.

Individual Burials

In total, 66 burials were recovered from the site at the Mississippi State Asylum,

but this oral microbiome reconstruction will focus on four individual burials, 1, 3, 4, and

24, that were sequenced. Burials 1, 3, 4, 24, and 47 were sampled since they had the

highest amount of macroscopically visible dental calculus of all of the recovered burials.

The summaries of these burials are in Table 2.1. These were recovered from an area that

was dated to c. 1920, and so these burials are a sample from the later years that the MSA

was active. Unfortunately, there is no information pertaining to the length of stay, or the

locations for these specific individuals.

Table 2.1 Summary of Osteoware Burial Records

Burial Sex Age Range Notes 1 Female 20-35 years Some dental calculus 3 Probable Male 35-39 Extensive caries, dental calculus, and Linear Enamel Hypoplasias (LEH) 4 Probable Female 40-44 Significant dental calculus 24 Female 20-35 Significant dental calculus 47* Unknown 22+ Highly fragmentary, some dental calculus These notes are copied verbatim from the Osteoware records. The scoring was performed by was previously scored by Amber Plemons following a modified version of Buikstra and Ubelaker’s methodology (1994). Burial 47 was not sequenced due to an insufficient amount of microbial DNA extracted from the sample (Table 4.1), although additional calculus was sampled for SEM (Table 3.2).

The Human Oral Microbiome Database

This database was created to provide the scientific community with

comprehensive information concerning the total prokaryote biodiversity that is present in

the modern human oral microbiome (Chen et al. 2010; Dewhirst et al. 2010). These have

been recorded by previously published studies. The HOMD serves as a modern

comparison point for the MSA samples, in order to determine if their biodiversity is

similar to modern oral microbial diversity.

CHAPTER III

MATERIALS AND METHODS

To evaluate the research questions, samples of dental calculus were removed from four individuals previously recovered from the cemetery of the Mississippi State Asylum

(MSA). The skeletal remains are currently curated at Mississippi State University. A total of 24 samples were removed from the burials. These samples were divided into two

subsets: the first subset (N=11) was removed for bacterial DNA isolation and sequencing,

and the second subset (N=13) was removed for Scanning Electron Microscopic (SEM)

imaging. SEM imaging was performed to visualize the structure of the dental calculus,

and to identify the presence of preserved bacteria. From the first subset, four samples of

dental calculus had a detectable amount of bacterial DNA. These samples (N=4) were

then sequenced.

Hypotheses

Hypothesis 1:

The samples of sequenced dental calculus (N=4) will generate a detectable

amount of microbial DNA, which will allow for the generation of a high-resolution

taxonomic characterization of the oral microbiome.

This hypothesis enabled evaluation of the first through fifth research questions.

The metagenomically sequenced microbial DNA was run through 16S metagenomic sequencing in Illumina BaseSpace, which generated full quantitative and qualitative 16

taxonomic trees to the species level. The data generated from the sequencing, and the taxonomic trees was used in answering the additional research questions.

Hypothesis 2:

When compared, the overall biological diversity of the samples (N=4) will be

statistically different, despite similar living conditions and food.

This hypothesis was used to answer the second research question. The number of

individual species and their quantitative presence was recorded from the 16S sequencing.

The established taxonomic trees were used to calculate the individual biodiversity and the

biodiversity of each taxonomic level. Statistics and analytical software were used to perform chi square tests of independence to compare the quantitative biodiversity of each sample. In addition, the Shannon diversity index was calculated for each sample.

Hypothesis 3:

There will be a presence of opportunistically pathogenic oral bacteria, such as S.

mutans, in the samples (N=4) due to the lack of adequate nutrition during time spent in the asylum.

This hypothesis was tested using the taxonomic trees in order to answer the third research question. In addition, targeted Polymerase Chain Reaction (PCR) and gel electrophoresis were performed using specific primers to identify known opportunistically pathogenic oral bacteria. When the oral microbiome is not disrupted by diminished nutrition or decreased hygiene, the presence of the commensal bacteria aids

in protection against pathogenic bacteria (Wade 2013). It is possible that in the asylum,

due to the lack of proper nutrition, and decreased hygienic standards, the normal flora of

these four individuals was disrupted. If disrupted, there might be a presence of pathogenic bacteria that opportunistically colonized the oral cavity, and surfaces of the teeth.

Hypothesis 4:

There will be non-commensal, non-oral pathogenic bacteria, such S. pneumoniae,

in these samples due to the unsanitary conditions of the Mississippi State Asylum, and

the prevalence of bacterial epidemics, prior to death.

This hypothesis was tested using the established taxonomic trees in order to

answer the fourth research question. Additionally, targeted sets of PCR and gel

electrophoresis were performed using specific primers to try and identify known

pathogenic bacteria. Due to the close and unsanitary conditions of institutions during that

time, contagious disease was rampant, and mortality from these types of diseases was

high. Therefore, there may be potential for the identification of these types of bacteria in

the samples of dental calculus.

Hypothesis 5:

The samples from the Mississippi State Asylum will differ in their biodiversity

when compared to modern oral microbiomes when compared against the Human Oral

Microbiome Database (HOMD).

This hypothesis was tested using the established phyla to visually compare the

diversity of the samples from the MSA (N=4) and the HOMD (Chen et al. 2010).

Statistics and analytical software were used to perform chi square tests of independence

to compare the qualitative biodiversity of each sample.

Sampling methods

The deposits of dental calculus present on the total number of individuals

recovered from the MSA cemetery were screened by Molly Zuckerman to assess which

were likely large enough to yield at least 0.05 mg of dental calculus. Dental calculus on

each of these individuals was previously scored by Amber Plemons following a modified

version of Buikstra and Ubelaker’s (1994) standards. The dental calculus was

preferentially taken from the first and second molar maxillary molars, subsequently

followed by the third molars, and then by the first and second mandibular molars. If no

samples were removed, calculus was then removed from the maxillary premolars,

followed by the mandibular premolars, and subsequently by the canines and incisors.

This was performed by Molly Zuckerman following previously established standards for

the treatment and extraction of DNA from dental calculus (Warinner et al. 2014b). The

samples of dental calculus (N=11) were removed using a dental scaler which was

sterilized between every sample using isopropanol to remove contaminating live bacteria

and DNA, and prevent cross contamination (Table 3.1). The samples were placed individually into sterile 2.0 mL eppendorf centrifuge tubes, and wrapped with parafilm to prevent opening and contamination.

Table 3.1 Summation of Dental calculus sample removed for DNA isolation

Sample Burial number Side Maxilla or Mandible Tooth 1 1 Left Maxilla 3rd Molar 2 3 Left Maxilla 2nd Molar 3 4 Left Mandible 1st Premolar 4 4 Left Mandible 2nd Incisor 5 4 Left Mandible 1st Incisor 6 4 Right Mandible 1st Incisor 7 4 Right Mandible 2nd Premolar 8 24 Right Maxilla 2nd Molar 9 24 Right Maxilla 1st Incisor 10 24 Left Maxilla 2nd Molar 11 47 Left Maxilla 1st Molar Table 3.1 is a summation of samples of dental calculus removed by Dr. Molly Zuckerman.

Scanning Electron Microscopy Sampling

Samples for SEM imaging were removed from burials 24 and 47, due to the presence of additional dental calculus. This was performed by Jonathan Belanich in

accordance with previously established methods (Warinner et al. 2014b). The samples

and the tooth locations are recorded in Table 3.2. Dental calculus samples were removed using a dental scaler, which was sterilized between every sample using 70% molecular grade isopropanol. Once removed, the samples (N=13) were stored individually in sterile

2.0 mL eppendorf centrifuge tubes, and were sealed with a parafilm wrap to ensure a total seal.

Table 3.2 Summation of Dental calculus samples removed for SEM

Sample Burial number Side Maxilla or Mandible Tooth 1 24 Right Maxilla Molar 2 24 Left Maxilla 1st Premolar 3 24 Left Maxilla 2nd Premolar 4 24 Right Mandible Molar 5 24 Right Mandible Incisor 6 24 Left Mandible Molar 7 24 Left Mandible Incisor 8 24 Left Mandible Premolar 9 47 Left Maxilla Premolar 10 47 Right Mandible Molar 11 47 Right Mandible Molar 12 47 Right Mandible Premolar 13 47 Right Mandible Incisor Table 3.2 is a summation of the samples of dental calculus removed by Jonathan Belanich, prior to SEM. For SEM, presence of calculus was more important than the size of the sample removed.

Microbial DNA Preparation

The microbial DNA was isolated in the Laboratory of Dr. Heather Jordan, Harned

Hall, Mississippi State University. Negative controls were utilized to assess the level of

bacterial contamination from the laboratory samples. The laboratory and equipment used

in this experiment, while not being a dedicated aDNA laboratory, is a Biosafety Level 2

(BSL-2) laboratory that has dedicated spaces for DNA/RNA isolation. In addition, the

laboratory follows EPA standards of workflow, sample preparation and decontamination

for quality control of analyses (Keya et al. 2008).

DNA Isolation Kits

A PowerBiofilm® DNA Isolation Kit from MO BIO Laboratories, Inc, was used

on the samples (N=11) of dental calculus and the controls (Manual). Bead beating was

used to lyse the microbial cells, extract the DNA, and elute it in solution (Probst et al.

2013). A Qubit® 2.0 Fluorometer was then used to determine the concentration of the

DNA (ng/µL) in each sample prior to sequencing. The concentrations are recorded in

Table 4.1. This allowed for the identification of the four samples with the highest

concentrations of DNA for each individual burial. These samples (N=4) and the negative

control were sent to Dr. Jason Rosch, Department of Infectious Disease at St. Jude

Hospital for metagenomic sequencing. The negative control, lacking a sample template

was used to assess the amount, if any, of laboratory or kit reagent contamination.

Targeted Polymerase Chain Reaction and gel electrophoresis

Six primers were used to try and identify specific bacteria from the microbial

DNA of the four samples. Table 3.3 contains a list of the primers and the diseases that

they cause. These primers were used to test for opportunistic and obligate pathogenic

bacteria.

Table 3.3 Table of bacterial specific primers for targeted PCR.

Primer Bacteria Causative agent of 1* Porphyromonas gingivalis Periodontal disease (Red Complex) 2* Tannerella forsythia Periodontal disease (Red Complex) 3* Treponema denticola Periodontal disease (Red Complex) 4 Aggregatibacter comitans Periodontal disease 5 Enterococcus aggregate Endodontic infection 6 Streptococcus pneumoniae Bacterial pneumonia Primers that are marked with an (*) were a part of a single set. These three bacteria comprise the “red complex” and when found in conjunction, typically cause intensive periodontitis (Holt and Ebersole 2005). With the exception of the S. pneumonia, all of these bacteria are opportunistic pathogens and can survive in the oral microbiome in low concentrations.

Metagenomic Sequencing

DNA was purified using the MOBIO DNA Purification kit to achieve highly purified samples. DNA libraries were prepared and adapter and unique index (for purposes of multiplexing) ligation was achieved using the TruSeq sample preparation kit

(Epicentre). Multiplexed transcripts were sequenced using Illumina HiSeq next- generation pair-end sequencing technology to describe the bacterial communities.

This sequencing approach allowed up to multiplexing of 384 samples per run with

2 x 100 bp paired-end reads. The raw fastq files barcoded Illumina reads were assembled, and quality-filtered where reads were discarded if they had a quality score < Q20, or contained ambiguous base calls or barcode/primer errors.

Illumina BaseSpace Analysis

The data that was received from Dr. Rosch was uploaded into Illumina

BaseSpace. 16S Metagenomics | Version: 1.0.1 was used to perform the taxonomic

analysis.

Scanning Electron Microscopy

The samples (N=13) that were used in the scanning electron microscope were

processed at the Institute for Imaging & Analytical Technologies (I2AT) in the Clay Lyle

Building at Mississippi State University. Samples were removed from their vials and

mounted onto individual pegs using double sided adhesive carbon tape. Samples were

placed either dorsally or ventrally, so that images were taken of both the side in contact

with the tooth (ventral) and the side exposed to the oral cavity (dorsal). Samples 9 and 11

were fractured in half using sterile forceps and were placed both dorsally and ventrally.

The determination of dorsal or ventral was performed using a light-dissecting microscope. The sample placement is recorded in Table 3.4. All samples were then

coated with a single coating of 15 nanometers of platinum (99.99% pure) using a Polaron

E5100 sputter coater. Samples were viewed using a Carl Zeiss EVO50VP Variable

Pressure Scanning Electron Microscope, and the images taken were stored digitally.

These methods were modified from previous studies examining dental calculus (Brady et

al. 1989; Lustmann et al. 1976; Schroeder and Shanley 1969) to use the modern

equipment that is available at the Institute for Imaging & Analytical Technologies at

Mississippi State University.

Table 3.4 Scanning Electron Microscopy Dental Calculus Samples

Sample Tooth Mounted Side 1 Molar Dorsal 2 1st Premolar Ventral 3 2nd Premolar Dorsal 4 Molar Unknown 5 Incisor Ventral 6 Molar Dorsal 7 Incisor Dorsal 8 Premolar Unknown 9 Premolar Dorsal and Ventral 10 Molar Unknown 11 Molar Dorsal and Ventral 12 Premolar Unknown 13 Incisor Ventral

Human Oral Microbiome Database Biodiversity

The data about the biodiversity of the modern human oral microbiomes was retrieved from the open source online database (Chen et al. 2010). The taxonomic

diversity was recorded and coded in the same as the data from the MSA to allow for comparative analysis.

Statistical Analyses

All statistical analyses and tests were performed using Microsoft Excel and IBM

SPSS Statistics V.23. Chi-square tests of independence (Pearson 1900) were performed

on the microbial data at the kingdom, phylum, class, and order taxonomic levels.

Phylogenetically related groups of bacteria were clustered together when necessary to

achieve the minimum cell count to perform these tests. Comparisons with the HOMD

were checked with Cramer’s V (Cramér 1946). The histogram was performed using an

agglomeration hierarchical cluster analysis, utilizing square Euclidian distance (Steinbach et al. 2000).

CHAPTER IV

RESULTS

The methods described in Chapter III were performed, and the microbial DNA

from the samples of dental calculus (N=4) were successfully isolated, sequenced, and

analyzed.

DNA Isolation Results

The isolation of the microbial DNA was successful, and yielded enough bacterial material to warrant sequencing (Table 4.1). Burial 1 and burial 3 provided enough DNA from a single piece of dental calculus that they could be sequenced individually. Burial 4 did not provide enough microbial DNA from a single sample, and so the samples 3-7, were combined into a single burial 4 sample for sequencing (Table 4.1). This combination yielded a significantly increased amount of DNA. Additionally, burial 24 did not provide enough microbial DNA from a single sample, and so the samples 8-10 were again combined into a single sample (Table 4.1). Burial 47 did not have a detectable concentration of microbial DNA in the provided sample of dental calculus, and was

therefore not sequenced. Furthermore, the negative control sample did not have a

detectable amount of DNA according to Qubit quantification, but was sequenced to

determine the potential level of contamination.

Table 4.1 Concentration of microbial DNA (ng/µL) in isolated samples

Sample number Sequencing sample Burial Number Total Concentration 1 HJ001 1 9.0 ng 2 HJ002 3 9.0 ng 3 HJ004 4 58.8 ng 4 - 4 - 5 - 4 - 6 - 4 - 7 - 4 - 8 HJ005 24 35.82 ng 9 - 24 - 10 - 24 - 11 - 47 Concentration undetectable Negative Control HJ006 - Concentration undetectable

Table 4.1 refers to the total microbial concentration of the samples of dental calculus, as tested with the Qubit® 2.0 Fluorometer. Samples 3-7 were combined into sample HJ004, and samples 8-10 were combined into sample HJ005.

16S Metagenomic Analysis

The bacterial DNA extracted from the dental calculus was successfully analyzed

by using the 16S Metagenomics application on BaseSpace. These data are shown in

Table 4.2.

Table 4.2 16S Metagenomics Aggregate Results

Sample Burial Number of Percent Reads Number of Shannon Number Number Reads Classified to Species Diversity Genus Identified Index 1 (HJ001) 1 133,734 0.14% 51 22.1% 2 (HJ002) 3 377,864 0.14% 147 25.9% 3 (HJ004) 4 838,175 0.05% 114 27.1% 4 (HJ005) 24 241,002 0.06% 70 24.9% 5 (HJ006) Negative 2,199 0.05% 1 1.4% Control

Data acquired from BaseSpace 16S Metagenomics Aggregate Analysis. The Shannon Diversity Index data was calculated manually.

Burials

The 16S metagenomic analyses resulted in positive identification of bacterial

species, with a mean of 95.5 ± 43.3 identified species (Table 4.2). This refers only to the

DNA sequences that could be identified fully to the species level; certain sequences could

only be identified to the genus, or other taxonomic levels. The full lists of bacterial

genera and species are reported in Appendix A (Tables A.1-A.4), and the visual

representation of the bacterial diversity at the kingdom-genus taxa are reported in

Appendix B. The low percentage of the reads classified to genus is due to the DNA being

sequenced for metagenomic analysis. The non-classified reads are additional genetic

material that references genes, rather than bacteria, and so would not be classified when

sequenced using standard 16S databases (Shah et al. 2011). The Shannon Diversity index,

which is a measure of statistical diversity, was calculated manually. This is a measure of

the abundance and evenness of species present in ecological samples.

Negative Control

The negative control was performed in the same manner as the other samples, and

was subject to the same protocols and methods. This negative control resulted in only

2,199 reads, and in one positive bacterial match (Table 4.2). Furthermore, this bacterial

species was not identified within any of burial samples. This indicates that there was little

contamination of the samples during DNA extraction, isolation, and processing. The

single bacterium that was identified was Frankia alni (Table A.5) a nitrogen-fixing

environmental bacteria that exists symbiotically with tree root systems (Schwencke and

Carú 2001).

Scanning Electron Microscopy

SEM was performed on the samples of dental calculus, and individual bacteria, as well as large groups of bacteria were clearly visible (Figures 4.1-4.4). Bacteria were found on both dorsal and ventral sides of the dental calculus, and were clearly visible to around 20,000 times magnification. Also visible were fungal hyphae (Figure 4.4) and mineralized elements (Figure 4.5). Sample 7 (Table 3.4) was the most photographed due to the abundance of bacteria on the surfaces of the photos, and the relative evenness of platinum coating (Figures 4.1-4.3). Sample 5 was photographed for the high concentration of fungal hyphae on the surface, and present in the crevasses in the calculus

(Figure 4.4). Figure 4.5 was included since it shows a very clear mineralized element.

10 µm

Figure 4.1 Photo of bacteria on the surface of dental calculus

This photo taken at 4,460 times magnification clearly shows the abundance of bacteria on the surface of dental calculus from the dorsal side of sample 7 (Table 3.4)

2 µm

Figure 4.2 Photo of bacteria on dental calculus

This photo taken at 4,930 times magnification further demonstrates the abundant presence of bacteria on the dental calculus from the dorsal side of sample 7 (Table 3.4)

1 µm

Figure 4.3 High magnification photo of bacteria on dental calculus

This photo taken at a magnification of 17,150 times shows the surface structure of bacteria on the outside of dental calculus from the dorsal side of sample 7 (Table 3.4)

10 µm

Figure 4.4 Photo of fungal hyphae on dental calculus

This photo magnified at 2,520 times shows bacteria trapped within a matrix of microscopic fungal hyphae from the ventral side of sample 5 (Table 3.4)

20 µm

Figure 4.5 Photo of mineral element in dental calculus

This photo which was taken at 1,960 times shows a mineral element present on the surface of the dental calculus from the dorsal side of sample 6 (Table 3.4)

Chi-Square Test of Independence

The data were evaluated statistically to determine the similarity or dissimilarity,

of the reconstructed microbiomes. The samples (N=4) were statistically significant at each level, indicating that they were statistically different with the p-values decreasing at

each level: Kingdom (X2 = 32.43; df = 3; p < 0.0001); Phyla (X2 = 331.42; df = 9; p <

0.0001); Class (X2 = 453.03; df = 12; p < 0.0001); and Order (X2 = 562.81; df =15; p <

0.0001).

Microbes in Burials

From these samples (N=4) a total of 778 bacteria were identified, representing

303 individual species. This includes commensal and pathogenic bacteria. The frequency of the biodiversity is visualized in figures B.1-B.6 (Appendix B).

Specific Pathogenic Bacteria

In these samples pathogenic bacteria were positively identified at the species level. Staphylococcus aureus, Streptococcus pneumoniae, Streptococcus

pseudopneumoniae, Haemophilus parainfluenzae, Streptococcus bovis, Listeria

monocytogenes, Clostridium tetani, and Clostridium botulinum, among others were

identified. Additional taxons were identified, but not at the genus or species levels. It is

possible that the DNA was degraded, and did not allow for positive identification of a

specific species.

Targeted Polymerase Chain Reaction

PCR was performed on the samples in order to try and identify specific

pathogenic bacteria. The bacterial primers used are summarized in table 3.3. These PCR

analyses were all negative, and none of the bacteria tested for were identifiable though

this method. The species level 16S metagenomic analysis did not concur with the

negative PCR tests; one of the pathogenic bacteria was found in the samples of dental

calculus. Streptococcus pneumoniae was identified through DNA sequencing, but not

through PCR. Potentially, this is due to the low amount of S. pneumoniae in the isolated

DNA, giving a false negative result from the PCR and gel electrophoresis.

Phyla Comparison against Modern HOMD

The comparison to modern oral microbiomes was tested using chi square goodness of fit (Pearson 1900), and checked with Cramer’s V (Cramér 1946). This was not tested using chi square tests of independence, since the large amount of observations contributed to edge effects. To ensure that the minimum cell count was met, all phyla that represented less than 10% of the total diversity was combined into an “other” category

(Figure 4.6). When compared against the HOMD, the microbial compositions of burials from the MSA are not very similar. Each of the four burials was statistically different from the HOMD: buria1 1 (X2 = 65.11; df = 3; p < 0.0001; V = 0.37), burial 3 (X2 =

60.43; df = 3; p < 0.0001; V = 0.13), burial 4 (X2 = 66.42; df = 3; p < 0.0001; V = 0.17),

and burial 24 (X2 = 25.83; df = 3; p < 0.0001; V = 0.15). The divergence of the burials

from the HOMD can be visually seen in Figure 4.7.

Three distinct phyla are similar between the two sets of data, Actinobacteria,

Firmicutes, and Proteobacteria. Actinobacteria are the most similar with 12.6% of the

total phyla for the HOMD and 18.6% for the MSA samples. Firmicutes are less similar,

comprising 33.7% of the total phyla for the HOMD and 41.7% for the MSA samples.

Proteobacteria comprise 16.3% of the HOMD, but make up 24.4% of the MSA sample

diversity. Most different were the Bacterodetes and the Spirochaetes. Bacterodetes make

up 17.3% of the HOMD, but only 0.8% of the MSA samples, while Spirochaetes represent 7.0% of the HOMD, but only 0.3% of the MSA oral microbiomes.

Figure 4.6 Phyla comparison of the four individual burials and the data acquired from the HOMD.

Phyla that were represented less than 10% were grouped into the other category. This was performed visually demonstrate the microbial diversity used in the statistical tests.

Figure 4.7 Phyla Dendrogram

This figure is a visualization of a cluster analysis performed on the phyla from the burials and the HOMD. This dendrogram does not include identified environmental contaminants (Table 4.3). This demonstrates the divergence of the burials from each other, and from the HOMD.

Environmental Contaminants

Several of the bacterial groups that were identified might be possible taphonomic contaminants from the soil from which the burials were extracted. The phyla

Acidobacteria, Caldiserica, Caldithrix, Crenarchaeota, Cyanobacteria, Deferribacteres,

Gemmatimonadetes, Nitrospirae, Planctomycetes, Thermi, Thermodesulfobacteria,

Thermotogae, Verrucomicrobia are not found within the taxonomic hierarchy of the

HOMD (Chen et al. 2010), and do not seem to be natural oral bacteria (Table 4.3). Many

of these phyla are newly recognized, and in some cases, Acidobacteria are typically found in soils and function as a part of the ecosystem (Rawat et al. 2012; Ward et al.

2009), but it is interesting to note that this phylum has previously been found to be a contaminant of bacterial isolation kit reagents, and it is possible that this contamination was from the kit itself (Salter et al. 2014). Additionally, Alteromondales are an order of

Gram-negative, marine bacteria, and would not normally be found within the oral microbiome (Ivanova et al. 2004). Holophagales are another Gram-negative order of marine bacteria, not typically found in the oral cavity (Fukunaga and Ichikawa 2014).

Table 4.3 Natural environments of potential bacterial contaminants

Phyla Environment Citation Acidobacteria Soils (Rawat et al. 2012) Caldiserica Unknown (Mori et al. 2009) (Miroshnichenko et al. 2003) Caldithrix Marine hydrothermal vents Crenarchaeota Soils (Jurgens et al. 1997) (De Los Ríos et al. 2007; Cyanobacteria Soils, marine Proctor and Fuhrman 1990) Deferribacteres Unknown (Jumas-Bilak et al. 2009) Gemmatimonadetes Soils (Fawaz 2013) Nitrospirae Marine (Watson et al. 1986) Planctomycetes Marine (Fuerst 1995) Thermi Soils (Griffiths and Gupta 2010) Thermodesulfobacteria Freshwater hot springs (Vick et al. 2010) Thermotogae Soils, oil reservoirs (Nesbø et al. 2010) Verrucomicrobia Soils, freshwater (Cho et al. 2004)

CHAPTER V

DISCUSSION AND CONCLUSION

This study has allowed for the successful reconstruction and the examination of oral microbiomes from four individuals, which has provided a glimpse into the oral biodiversity of these individuals, allowing for meaningful analysis. The results of this

work clearly allow for the hypotheses and the overarching research question to be

answered. The data supported the first hypothesis, in that there was an adequate amount

of microbial DNA to generate the high-resolution taxonomic characterizations for each of

the samples (Table 4.1-2; Appendix B), and this allowed for the subsequent hypotheses to be tested. The negative laboratory control sample indicates the low level of bacterial contamination that occurred during the DNA isolation process. The only species

identified in the negative control was Frankia alni (Table B.4), and this bacterium was

not found in any of the other four samples. These results demonstrate the viability of

historic dental calculus research to be performed outside of an ancient DNA laboratory, while still maintaining proper anti-contamination equipment and methodologies.

Additional experiments, utilizing the methodologies from this study, analyzing samples

of ancient DNA are required before any definitive conclusions can be drawn regarding the feasibility of performing aDNA research in this type of laboratory.

The data generated from the taxonomic hierarchies allowed for the statistical

comparison of the biodiversity of the four microbiomes. Both qualitative and quantitative

data were generated, allowing for testing of overall composition and the presence of specific bacteria. Chi-square tests demonstrated that the statistical difference increased by

orders as the taxonomic differences increased, indicating that the oral microbiomes of

each of the individuals were statistically unique. Even though the four individuals were in

the same asylum, their oral microbiomes were composed of different combinations of

bacteria (Figures B.1-6).

Through comparisons, it can clearly be seen that the samples from the MSA were quite different from the biodiversity present in modern human oral microbiomes, as indicated by the HOMD (Chen et al. 2010). The MSA microbiomes had a lower diversity than the HOMD (Figure 4.6), but were more similar to each other than to the HOMD

(Figure 4.7). The composition of the microbiomes of the individuals was not only

dissimilar to modern microbiomes, but statistically different. Many phyla seen in the

HOMD were not represented in the MSA samples, and of the three main phyla that were

present, Actinobacteria, Firmicutes, and Proteobacteria, there were significant differences

in the composition. These results do tend to indicate that individuals from the Mississippi

State Asylum had significantly decreased biodiversity in their oral microbiomes. It is

possible that the recovered dental calculus was altered taphonomically by environmental

bacteria, but this would have to be examined through the inclusion of additional samples,

and the sequencing of associated soil samples.

Although the taxonomic reconstruction was successful, there was no positive

identification of any opportunistic oral pathogens such as Streptococcus mutans,

Porphyromonas gingivalis, Tannerella forsythia, or Treponema denticola. S. mutans in

particular is a commonly occurring oral bacteria that is typically associated with the

consumption of processed sugars (Burt and Pai 2001; Ikeda et al. 1978), and forms biofilm matrices when exposed to glucose (Shemesh et al. 2007). In turn, this biofilm

formation aids in the attachment of additional pathogenic oral bacteria, including P.

gingivalis (Kolenbrander et al. 2010; McNab et al. 2003). The lack of S. mutans and other

opportunistic oral pathogens could indicate that individuals in the MSA did not have

access to adequate nutrition; potentially there was no access to processed foods, and

individuals were fed complex, non-processed carbohydrates such as starch, deriving from

corn or maize. From Searcy (1907), it was seen that this was the case in a similar asylum;

individuals were primarily fed an insufficient diet of unprocessed corn meal. This lack of

diversity in the diet would have had significant effects on the composition of the oral

microbiome. It is noted that S. mutans does not require tryptophan or niacin for bacterial

growth (Terleckyj and Shockman 1975), and so its absence is not due to the lack of a

specific required nutrient, but a lack of metabolizable products.

Since the normal commensal flora defend against pathogenic colonization, the

lack of processed sugars could have afforded the commensal bacteria an advantage, in

that there were fewer metabolizable products for opportunistic pathogens to utilize. These

colonizing bacteria present in the oral cavity would have been the hardiest members of

the oral microbiome, effectively, the only members able to survive in a nutrient deficient environment. Their colonization could have blocked opportunistic pathogens from

establishing colonies, or if bacteria such as S. mutans were able to attach, the lack of

simple metabolizable compounds could have effectively caused starvation. This further

indicates that the individuals in this asylum did not receive adequate nutrition, or a

requisite amount of food. Since opportunistically pathogenic bacteria require simple

carbohydrates, the lack of processed nutrition might have cultivated a “healthy” oral microbiome, while potentially causing other diseases such as pellagra. Furthermore, any dental caries that were present on the teeth may have formed prior to institutionalization.

Additionally, since the average stay in the asylum was thirteen months, the composition

of the dental calculus reflects the diets of Mississippi during this time period, as much as

it does the diet of the MSA.

Opportunistically pathogenic oral bacteria have been previously associated with a

higher intake of sugars (Ikeda et al. 1978; Lukacs 2012; Wade 2013), and Adler et al.

(2013) found that modern oral microbiomes contained a high amount of pathogenic

bacteria, and decreased biodiversity when compared to archaic samples These modern

oral microbiomes relatively low in biodiversity, and high in pathogenic bacteria due to

the consumption of a fully processed diet. An industrialized diet contains an abundance

of simple sugars and easily metabolizable products, allowing for the proliferation of

pathogenic bacteria (Adler et al. 2013; Wade 2013). The samples from the MSA

therefore seem contrary to this established paradigm, since they are from a period after

industrialization. However, these results do not contradict the findings of Adler et al.

(2013). Since individuals in the MSA were fed such an inadequate and unvaried diet,

their food did not contain enough processed sugars to allow for the growth of

opportunistic oral pathogens. Even though these individuals were from a post-

industrialized era, they were not being fed an industrialized diet. Therefore, these

reconstructed oral microbiomes reflect a unique, and marginalized, subset of the human

population of Mississippi at this time.

With respect to the oral pathologies present in the burials (Table 2.1), the only burial that had evidence of oral pathologies besides dental calculus was burial 3. This burial had evidence of dental caries, but no evidence of any opportunistic oral pathogens.

Therefore, these dental caries may have formed at a time prior to admittance to the MSA.

In addition, since there was no presence of periodontal disease, it was unsurprising that there was a lack of periodontal causing bacteria. Furthermore, due to the rough consistency of the diet, it may be possible the coarseness of the food abraded the teeth, and knocked off fragments of dental calculus, leading to the low prevalence of visible dental calculus in this sample.

Additionally, there was very little presence of infectious, pathogenic bacteria. The sample from Burial 1 contained Listeria monocytogenes, and the samples from Burial 24

contained three separate pathogenic species: Streptococcus pneumoniae, Streptococcus

pseudopneumoniae, and Haemophilus parainfluenzae. L. monocytogenes is a food-borne

bacterium which is the causative agent of listeriosis, and the two Streptococci are

causative agents of bacterial pneumonia, whereas Haemophilus is a causative agent of

bacterial endocarditis (Murray et al. 2015). Although Clostridium tetani, and Clostridium

botulinum were positively identified in the MSA samples, it is likely that these were

taphonomic contaminants since they typically exist in soils (Murray et al. 2015).

Most of the other bacteria that were identified were not associated with human

diseases. Any additional pathogenic bacteria that were present in the MSA were not

preserved in these samples of dental calculus. Or, it is possible that these individuals were not exposed to them. Since many of these pathogenic bacteria are airborne, and not static biofilm producers, they might not have maintained in the oral cavity, which would

explain why that they did not preserve in the dental calculus. Interestingly,

Mycobacterium tuberculosis was not detected, even though it was known to be present at

the MSA, and was one of the leading killers of individuals in the MSA (MSA records).

Additionally, since the microbes that were identified from these samples of dental calculus were extant 100 years ago, evolutionary effects might have played a role. Over time, the genome of the microbes might have changed through mutations and natural

selection. When the metagenomic data was analyzed, the recovered microbial DNA

might be different enough to create a false negative in the results. Even if a bacterial

species was present in the dental calculus, the modern genome might different enough to

cause misidentification. Bacterial sequences were identified at higher taxonomic levels,

but evolution might have made certain bacteria unidentifiable at the species level.

Future Work

Future work involving this data will analyze the metagenome of the sequenced samples. This will include additional targeted searching for pathogenic bacteria, identification of genes for virulence and pathogenicity, and identification of pathogenic viruses. Over 99% of the successfully sequenced DNA was not identified as bacterial species using 16S RNA databases, and therefore there is a plethora of data that are yet to be analyzed. Additional work will include the sequencing of contemporaneous non-

institutionalized samples to establish context. Furthermore, future dental calculus work

will be performed examining the effects of the first epidemiologic transition on the

biodiversity and composition of the human oral microbiome in the Southeastern United

States.

Conclusion

This study successfully reconstructed oral microbiomes through the extraction of

DNA from dental calculus. These samples of preserved dental calculus provided enough microbial DNA to not only reconstruct the composition of the oral microbiome, but to statistically compare the compositions. It was found that these samples from the MSA differed greatly in their composition from each other, as well as modern human oral

microbiomes.

One of the most surprising conclusions from this study is the marked absence of

common opportunistically pathogenic oral bacteria. The fact that there was no evidence

of these bacteria seems almost contradictory. It is possible however, that this might stem

from a relatively simple cause, a dearth of simple sugars. If the individuals housed in this

asylum were fed an insufficient diet comprised of unprocessed corn, they would have had

a primarily starch based diet. Starch, being a large and relatively complex carbohydrate,

would not have been efficiently utilized by certain opportunistic oral pathogens such as S. mutans. These bacteria prefer easily metabolized simple carbohydrates, which are found in processed foods, to which institutionalized individuals would not have had access. This

might have lead to the paradoxical result in which individuals in this study from the

Mississippi State Asylum had oral microbiomes with no detectable presence of

opportunistic oral pathogens, in spite of deficient nutrition.

This research provides novel information on the oral microbiome of a subset of a

population of marginalized individuals who lived within an institutionalized context,

furthering the applicability of dental calculus studies within the fields of archaeology and

anthropology. Future work on these and other samples will reveal more about the

evolution of the human oral microbiome, and metagenomic work will provide genomic evidence for the evolution of bacteria. This collaboration between the fields of microbiology, paleopathology and archaeology will, and should continue, as combined methodologies allow for the examination of new sources of material and the creation of new and informative types of research.

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APPENDIX A

GENUS AND SPECIES LISTS FOR SEQUENCED DENTAL CALCULUS

Table A.1 Burial 1 Genus and species list

Genus Species Number of hits Acetobacterium tundrae 4 Alkaliphilus 2 Alkaliphilus peptidifermentans 1 Alteromonas genovensis 1 Amaricoccus macauensis 1 Ammoniphilus oxalivorans 1 Anaerococcus octavius 1 Aneurinibacillus 2 Bacillus 3 Bacillus ginsenggisoli 2 Bacillus aryabhattai 1 Bacillus ginsengihumi 1 Bacillus alcaliinulinus 1 Bacillus horneckiae 1 Bacillus thioparans 1 Bartonella phoceensis 1 Brachybacterium arcticum 1 Brevibacillus thermoruber 1 Brochothrix 1 Caloramator uzoniensis 1 Caloramator mitchellensis 1 Caloramator indicus 1 Carnobacterium gallinarum 1 Chryseobacterium 1 Clostridium 30 Clostridium alkalicellulosi 10 Clostridium thermocellum 8 Clostridium straminisolvens 7 Clostridium thermosuccinogenes 2 Clostridium tetani 2 Clostridium aldrichii 1 Clostridium subterminale 1 Clostridium thermoalcaliphilum 1 Clostridium acetireducens 1 Clostridium clariflavum 1 Cohnella 2 Dehalobacterium 2 Desulfosporosinus 1 Desulfotomaculum australicum 1

Table A.1 (continued)

Genus Species Number of hits Devosia 1 Dietzia alimentaria 1 Faecalibacterium 3 Heliorestis 2 Kurthia zopfii 3 Lachnospira pectinoschiza 1 Listeria 1 Listeria monocytogenes 1 Methanobrevibacter acididurans 1 Natroniella 1 Natronincola peptidivorans 6 Negativicoccus succinicivorans 1 Nitrosococcus watsoni 1 Pedomicrobium 1 Pelobacter acetylenicus 1 Pelotomaculum 3 Peptoniphilus methioninivorax 3 Peptoniphilus coxii 3 Polynucleobacter 1 Propionibacterium acnes 3 Rhodobacter 1 Rhodococcus rhodochrous 1 Sedimentibacter 6 Sediminibacillus halophilus 1 Selenomonas infelix 1 Soehngenia saccharolytica 3 Sporanaerobacter 8 Sporanaerobacter acetigenes 3 Streptococcus bovis 1 Streptomyces roseosporus 5 Sulfurospirillum 1 Syntrophomonas 6 Syntrophomonas palmitatica 1 Tepidanaerobacter 1 Tepidimicrobium ferriphilum 6 Tepidimicrobium 1 Thermoanaerobacter 2

Table A.2 Burial 3 Genus and species list

Genus Species Number of hits Acetobacterium tundrae 1 Acidovorax 1 Actinoallomurus 5 Actinoallomurus yoronensis 2 Actinomadura vinacea 1 Actinomadura 1 Actinomadura chibensis 1 Actinomadura cremea 1 Aeromicrobium marinum 1 Aeromicrobium 1 Agromyces salentinus 1 Alteromonas addita 1 Amycolatopsis decaplanina 1 Anaerolinea thermolimosa 2 Anaerolinea thermophila 1 Anaerolinea 1 Anaeromyxobacter 1 Anoxybacillus 3 Anoxybacillus amylolyticus 1 Arthrobacter psychrochitiniphilus 2 Arthrobacter psychrolactophilus 2 Arthrobacter 2 Atopobium fossor 2 Azospirillum palatum 1 Bacillus simplex 61 Bacillus 41 Bacillus thioparans 19 Bacillus arbutinivorans 16 Bacillus foraminis 9 Bacillus niacini 9 Bacillus methanolicus 7 Bacillus fumarioli 6 Bacillus soli 6 Bacillus horneckiae 6 Bacillus oryzae 4 Bacillus aryabhattai 3 Bacillus asahii 2 Bacillus marisflavi 2 Bacillus cohnii 2

Table A.2 (continued)

Genus Species Number of hits Bacillus thermoamylovorans 2 Bacillus muralis 2 Bacillus longiquaesitum 2 Bacillus senegalensis 2 Bacillus selenatarsenatis 2 Bacillus litoralis 2 Bacillus circulans 2 Bacillus ginsenggisoli 1 Bacillus boroniphilus 1 Bacillus velezensis 1 Bacillus coagulans 1 Bacillus alcalophilus 1 Bacillus aerophilus 1 Bacillus butanolivorans 1 Bacillus humi 1 Balneimonas 2 Bartonella rochalimae 1 Bdellovibrio 1 Brevibacillus 3 Brevibacillus ginsengisoli 1 Brevibacillus formosus 1 Bulleidia extructa 1 Burkholderia ubonensis 1 Butyrivibrio proteoclasticus 1 Caldicellulosiruptor bescii 1 Caldilinea tarbellica 2 Caldisericum exile 1 Caldithrix 1 Caloramator mitchellensis 3 Caloramator viterbiensis 2 Caloramator 1 Calothrix parietina 1 Campylobacter 1 Candidatus Koribacter 1 Candidatus Methylacidiphilum 1 Candidatus Nitrososphaera 1 Candidatus Rhabdochlamydia crassificans 1 Candidatus Scalindua brodae 1 Carboxydocella ferrireducens 1

Table A.2 (continued)

Genus Species Number of hits Chondromyces pediculatus 1 Chthoniobacter flavus 1 Clostridium 6 Clostridium hveragerdense 3 Clostridium thermosuccinogenes 2 Clostridium acetireducens 2 Clostridium kluyveri 1 Clostridium sulfidigenes 1 Clostridium botulinum 1 Clostridium thermocellum 1 Clostridium aldrichii 1 Clostridium haemolyticum 1 Cohnella soli 3 Conexibacter 1 Curvibacter lanceolatus 1 Cystobacter armeniaca 1 Dactylosporangium maewongense 2 Dechloromonas 1 Deefgea rivuli 1 Deferribacter autotrophicus 1 Dehalobacterium 1 Dehalogenimonas lykanthroporepellens 3 Desulfosporosinus 1 Desulfotomaculum halophilum 2 Desulfovibrio brasiliensis 1 Desulfuromonas svalbardensis 1 Diaphorobacter 1 Euzebya tangerina 1 Flavisolibacter 1 Gemella cunicula 1 Gemmata obscuriglobus 1 Gemmatimonas aurantiaca 1 Geobacter sulfurreducens 1 Giesbergeria 1 Heliorestis 1 Herbaspirillum chlorophenolicum 1 Herbaspirillum rubrisubalbicans 1 Hydrogenophaga pseudoflava 1 Iamia 5

Table A.2 (continued)

Genus Species Number of hits Janthinobacterium 5 Janthinobacterium agaricidamnosum 1 Kaistobacter 3 Kineosporia 2 Kribbella koreensis 1 Kurthia zopfii 1 Lactobacillus 2 Lautropia mirabilis 1 Legionella 1 Leucobacter chironomi 1 Limnohabitans 3 Listeria 1 Longilinea arvoryzae 5 Macrococcus 1 Macrococcus brunensis 1 Marinobacterium sediminicola 1 Mesorhizobium septentrionale 1 Mesorhizobium loti 1 Methanosaeta concilii 1 Methanosaeta 1 Micrococcus luteus 1 Moorella 1 Mycobacterium 1 Natronincola peptidivorans 2 Negativicoccus succinicivorans 1 Niabella soli 1 Nocardioides 8 Nocardioides plantarum 4 Nocardioides islandensis 2 Nocardioides lentus 1 Nonomuraea 1 Novosphingobium 1 Oceanobacillus sojae 2 Oerskovia 3 Opitutus 1 Oxalobacter vibrioformis 8 Paenibacillus 1 Paucibacter 1 Pectinatus cerevisiiphilus 1

Table A.2 (continued)

Genus Species Number of hits Pedosphaera 1 Pelotomaculum 1 Peptoniphilus methioninivorax 2 Peptoniphilus gorbachii 1 Phenylobacterium 3 Pilimelia 2 Pirellula 1 Planococcus 6 Planococcus maritimus 2 Polynucleobacter rarus 1 Propionibacterium acnes 1 Propionispora hippei 1 Pseudomonas 1 Pullulanibacillus naganoensis 1 Ralstonia detusculanense 1 Ramlibacter 5 Ramlibacter tataouinensis 1 Rhizobium 1 Rhodococcus percolatus 1 Rhodoferax 1 Rhodoferax ferrireducens 1 Rhodoplanes 2 Rhodovibrio sodomensis 1 Saccharopolyspora 1 Schlegelella aquatica 1 Sebaldella termitidis 2 Sedimentibacter 2 Serinicoccus marinus 1 Soehngenia saccharolytica 2 Solibacillus silvestris 2 Sphaerochaeta pleomorpha 1 Sphingomonas suberifaciens 1 Sphingomonas wittichii 1 Sporanaerobacter 6 Sporanaerobacter acetigenes 3 Sporichthya polymorpha 1 Sporosarcina luteola 1 Sporosarcina contaminans 1 Sporosarcina ginsengi 1

Table A.2 (continued)

Genus Species Number of hits Staphylococcus kloosii 1 Streptococcus 1 Streptomyces roseosporus 6 Streptomyces luteireticuli 1 Streptomyces griseoaurantiacus 1 Sulfurospirillum 1 Syntrophomonas 3 Syntrophomonas curvata 1 Syntrophus 3 Tepidanaerobacter 2 Tepidimicrobium ferriphilum 2 Terracoccus 1 Thermoanaerobacterium islandicum 1 Thermobaculum 1 Thermobaculum terrenum 1 Thermobifida 1 Thermogemmatispora 2 Thermogemmatispora onikobensis 1 Thermosipho africanus 1 Thiobacillus denitrificans 4 Thiobacillus thioparus 3 Thiobacillus 2 Variovorax 1

Table A.3 Burial 4 Genus and species list

Genus Species Number of hits Acetobacter tropicalis 1 Acetobacterium tundrae 1 Acidimicrobium 1 Acidithiobacillus cuprithermicus 2 Acinetobacter 1 Actinomadura yumaensis 2 Actinomadura 2 Actinomadura miaoliensis 2 Actinomadura glauciflava 1 Actinomadura vinacea 1 Actinomyces cardiffensis 1 Actinomycetospora 1 Actinopolymorpha alba 2 Alkaliphilus crotonatoxidans 3 Aminiphilus circumscriptus 2 Ammoniphilus oxalivorans 1 Anaerofustis 2 Anaerolinea 12 Anaerolinea thermolimosa 8 Anaerolinea thermophila 1 Anaeroplasma abactoclasticum 1 Anaerostipes 1 Ancylobacter polymorphus 6 Arcobacter skirrowii 1 Arthrobacter 1 Arthrobacter psychrolactophilus 1 Arthrobacter psychrochitiniphilus 1 Bacillus horneckiae 2 Bacillus oryzae 1 Bacillus 1 Balneimonas 1 Beijerinckia mobilis 1 Bellilinea caldifistulae 4 Bellilinea 1 Blastochloris sulfoviridis 1 Caldilinea tarbellica 3 Caldithrix 1 Caloramator mitchellensis 3 Caloramator 1

Table A.3 (continued)

Genus Species Number of hits Candidatus Koribacter versatilis 2 Candidatus Liberibacter 3 Candidatus Microthrix parvicella 1 Candidatus Nitrososphaera gargensis 1 Candidatus Phytoplasma phoenicium 1 Candidatus Phytoplasma 1 Candidatus Solibacter 3 Candidatus Tammella 1 Clostridium 9 Clostridium thermocellum 5 Clostridium alkalicellulosi 4 Clostridium straminisolvens 2 Clostridium thermosuccinogenes 1 Clostridium histolyticum 1 Cohnella soli 1 Conexibacter woesei 3 Conexibacter 2 Dactylosporangium maewongense 1 Dactylosporangium aurantiacum 1 Dactylosporangium roseum 1 Dechloromonas aromatica 1 Dehalogenimonas lykanthroporepellens 5 Desulfobacter 1 Desulfomonile tiedjei 1 Desulfonatronum thiosulfatophilum 1 Desulfosporosinus auripigmenti 2 Desulfosporosinus acidiphilus 1 Desulfotomaculum putei 1 Desulfovibrio profundus 2 Desulfovibrio intestinalis 1 Desulfovibrio 1 Dokdonella fugitiva 1 Erysipelothrix inopinata 1 Gallionella capsiferriformans 1 Gordonia hirsuta 1 Granulicella tundricola 1 Halanaerobacter chitinivorans 1 Halomonas almeriensis 3 Halomonas johnsoniae 1

Table A.3 (continued)

Genus Species Number of hits Halorhodospira halochloris 3 Hydrogenophilus 3 Hyphomicrobium vulgare 1 Iamia 1 Ignavibacterium 1 Kibdelosporangium 1 Kitasatospora 1 Knoellia subterranea 1 Knoellia 1 Leucothrix mucor 2 Longilinea arvoryzae 16 Longilinea 1 Marinomonas mediterranea 1 Marinomonas 1 Megasphaera hominis 1 Methanobacterium curvum 1 Methanobacterium uliginosum 1 Methanobrevibacter smithii 11 Methanobrevibacter 6 Methanobrevibacter gottschalkii 1 Methylophaga 4 Micrococcus flavus 1 Micromonospora 2 Mogibacterium 2 Mycoplasma verecundum 1 Myroides odoratus 2 Natronincola peptidivorans 3 Natronincola 2 Negativicoccus succinicivorans 8 Nitratireductor kimnyeongensis 1 Nitriliruptor 1 Nitrosococcus watsoni 12 Nocardia novocastrensa 1 Novosphingobium yangbajingensis 1 Parascardovia 9 Patulibacter americanus 1 Pedomicrobium 4 Pelobacter acetylenicus 1 Peptoniphilus olsenii 1

Table A.3 (continued)

Genus Species Number of hits Petrotoga miotherma 1 Phascolarctobacterium succinatutens 2 Phenylobacterium 2 Pilimelia 2 Pirellula staleyi 1 Planctomyces limnophilus 1 Planifilum fimeticola 1 Polaromonas 7 Polynucleobacter 2 Propionicimonas 1 Propionispora hippei 1 Pseudoalteromonas 1 Pseudonocardia asaccharolytica 2 Rhodoplanes 6 Rhodospirillum 1 Rickettsia 1 Ruegeria lacuscaerulensis 1 Saccharopolyspora 3 Saccharopolyspora erythraea 1 Saccharothrix 1 Sedimentibacter 3 Sharpea azabuensis 1 Soehngenia saccharolytica 4 Solirubrobacter soli 2 Solirubrobacter 1 Sporanaerobacter acetigenes 1 Staphylococcus aureus 1 Steroidobacter 1 Streptacidiphilus 2 Streptococcus bovis 1 Streptococcus 1 Streptomyces roseosporus 13 Streptomyces 7 Streptomyces pseudogriseolus 2 Streptomyces poonensis 2 Streptomyces humiferus 1 Streptomyces nanchangensis 1 Streptomyces thermodiastaticus 1 Streptomyces viridosporus 1

Table A.3 (continued)

Genus Species Number of hits Streptomyces radiopugnans 1 Streptomyces thermospinosisporus 1 Streptomyces atrovirens 1 Syntrophobacter 1 Syntrophomonas palmitatica 2 Tepidanaerobacter syntrophicus 1 Tepidimicrobium ferriphilum 7 Tepidimicrobium 1 Terracoccus 2 Thermacetogenium phaeum 1 Thermobaculum 6 Thermobaculum terrenum 2 Thermobifida 1 Thermococcus 2 Thermodesulfatator atlanticus 1 Thermodesulfovibrio thiophilus 2 Thermogemmatispora onikobensis 1 Thiobacillus denitrificans 1 Thiobacillus 1 Tsukamurella 1

Table A.4 Burial 24 Genus and species list

Genus Species Number of hits Acetobacterium 1 Acetobacterium tundrae 1 Acidisphaera 2 Acidovorax 1 Acinetobacter bouvetii 1 Actinobacillus 1 Actinomadura vinacea 2 Actinomyces odontolyticus 1 Aggregatibacter segnis 1 Alteromonas genovensis 2 Alteromonas addita 1 Ammoniphilus oxalivorans 1 Anaerolinea thermophila 1 Anaerolinea 1 Anaeromyxobacter dehalogenans 3 Anaeromyxobacter 1 Anaerovibrio lipolyticus 1 Aneurinibacillus 4 Aneurinibacillus thermoaerophilus 1 Arcanobacterium 1 Azospirillum palatum 1 Bacillus niacini 1 Bacillus horneckiae 1 Bacillus longiquaesitum 1 Bacillus siralis 1 Bacillus methanolicus 1 Bacillus 1 Bacillus ginsenggisoli 1 Bacillus boroniphilus 1 Bacillus velezensis 1 Bacillus coagulans 1 Bacillus alcalophilus 1 Bacillus aerophilus 1 Bacillus butanolivorans 1 Bacillus humi 1 Balneimonas 2 Bartonella rochalimae 1 Bartonella rochalimae 1 Bdellovibrio 1

Table A.4 (continued)

Genus Species Number of hits Borrelia 1 Bradyrhizobium liaoningense 1 Brevibacillus ginsengisoli 1 Brevibacillus formosus 1 Brevibacterium 1 Bulleidia extructa 1 Burkholderia ubonensis 1 Butyrivibrio proteoclasticus 1 Caldicellulosiruptor bescii 1 Caloramator 1 Campylobacter 1 Candidatus Koribacter 3 Candidatus Koribacter versatilis 1 Candidatus Methylacidiphilum 1 Candidatus Protochlamydia amoebophila 1 Candidatus Rhabdochlamydia 1 Candidatus Scalindua brodae 1 Candidatus Solibacter 2 Carboxydocella ferrireducens 1 Chondromyces pediculatus 1 Chthoniobacter flavus 1 Clostridium 5 Clostridium fallax 2 Clostridium novyi 1 Clostridium sulfidigenes 1 Clostridium botulinum 1 Clostridium thermocellum 1 Clostridium aldrichii 1 Clostridium haemolyticum 1 Cohnella 1 Curvibacter lanceolatus 1 Cystobacter armeniaca 1 Dechloromonas 1 Deefgea rivuli 1 Dehalobacterium 1 Desulfobacter 1 Desulfosporomusa polytropa 2 Desulfosporosinus 1 Desulfotomaculum australicum 1 70

Table A.4 (continued)

Genus Species Number of hits Desulfovibrio brasiliensis 1 Desulfurispora thermophila 1 Desulfuromonas svalbardensis 1 Diaphorobacter 1 Euzebya tangerina 1 Exiguobacterium 1 Gallionella capsiferriformans 2 Gallionella 1 Gemella cunicula 1 Gemmata obscuriglobus 1 Gemmatimonas aurantiaca 1 Geobacillus 1 Geobacter bremensis 1 Geobacter sulfurreducens 1 Georgenia 1 Geothrix fermentans 2 Giesbergeria 1 Granulicatella 1 Haemophilus parainfluenzae 1 Helicobacter suncus 1 Heliorestis 1 Herbaspirillum chlorophenolicum 1 Herbaspirillum rubrisubalbicans 1 Hydrogenophaga pseudoflava 1 Hydrogenophilus 1 Hyphomicrobium aestuarii 1 Janthinobacterium 5 Janthinobacterium agaricidamnosum 1 Kaistobacter 3 Kurthia zopfii 1 Lactobacillus 1 Lactobacillus equi 1 Lautropia mirabilis 1 Lautropia mirabilis 1 Legionella 1 Limnohabitans 3 Listeria 1 Longilinea arvoryzae 2 Macrococcus 1

Table A.4 (continued)

Genus Species Number of hits Macrococcus brunensis 1 Marinobacterium sediminicola 1 Mesorhizobium septentrionale 1 Mesorhizobium loti 1 Methylobacillus 1 Moorella 1 Nannocystis 1 Natronincola 2 Negativicoccus succinicivorans 1 Nitrosococcus watsoni 2 Nitrospira 1 Novosphingobium 1 Opitutus 1 Oxalobacter vibrioformis 8 Paenibacillus 1 Paenibacillus 1 Parascardovia 1 Paucibacter 1 Pectinatus cerevisiiphilus 1 Pediococcus argentinicus 1 Pediococcus cellicola 1 Pedobacter 1 Pedosphaera 1 Pelotomaculum 1 Pelotomaculum 1 Peptococcus niger 1 Peptoniphilus gorbachii 1 Phenylobacterium 3 Pirellula 1 Planctomyces 1 Planctomyces limnophilus 1 Planomicrobium psychrophilum 1 Polynucleobacter 1 Polynucleobacter rarus 1 Polynucleobacter rarus 1 Propionibacterium acnes 1 Propionibacterium humerusii 1 Propionispora hippei 3 Propionispora hippei 1

Table A.4 (continued)

Genus Species Number of hits Pseudomonas 1 Pseudonocardia kongjuensis 1 Pullulanibacillus naganoensis 1 Ralstonia detusculanense 1 Ramlibacter 5 Ramlibacter tataouinensis 1 Rhizobium 1 Rhodoferax 1 Rhodoferax ferrireducens 1 Rhodoplanes 1 Rhodoplanes 2 Rhodovibrio sodomensis 1 Rothia mucilaginosa 3 Schlegelella aquatica 1 Schlegelella aquatica 1 Sebaldella termitidis 2 Sebaldella termitidis 2 Sediminibacillus halophilus 1 Solirubrobacter soli 1 Sphaerochaeta pleomorpha 1 Sphingobium 1 Sphingomonas suberifaciens 1 Sphingomonas wittichii 1 Sporanaerobacter 1 Sporosarcina luteola 1 Sporosarcina contaminans 1 Sporosarcina ginsengi 1 Sporotomaculum syntrophicum 1 Staphylococcus caprae 1 Staphylococcus kloosii 1 Streptococcus 5 Streptococcus vestibularis 2 Streptococcus pseudopneumoniae 2 Streptococcus bovis 2 Streptococcus intermedius 1 Streptococcus parasanguinis 1 Streptococcus infantis 1 Streptococcus fryi 1 Streptococcus pneumoniae 1

Table A.4 (continued)

Genus Species Number of hits Streptococcus 1 Streptomyces roseosporus 2 Streptomyces rubrogriseus 1 Streptomyces viridosporus 1 Streptomyces chromofuscus 1 Streptomyces danangensis 1 Sulfurospirillum 1 Syntrophomonas sapovorans 1 Syntrophomonas curvata 1 Syntrophus 3 Tepidimonas ignava 1 Thermoanaerobacterium islandicum 1 Thermobacillus xylanilyticus 1 Thermococcus 1 Thermosinus 3 Thermosipho africanus 1 Thiobacillus denitrificans 4 Thiobacillus thioparus 3 Thiobacillus 2 Trichococcus 1 Variovorax 1 Vulcanithermus mediatlanticus 1

Table A.5 Negative control Genus and species list

Genus Species Number of hits Frankia alni 1

APPENDIX B

HIGH-RESOLUTION TAXONOMIC CHARACTERIZATIONS OF THE

RECONSTRUCTED ORAL MICROBIOMES

Figure B.1 Kingdom Comparison

Figure B.2 Phyla Comparison

Figure B.3 Class Comparison

Classes totaling less than 1% of total biodiversity were grouped into the “other” category.

Figure B.4 Order Comparison

Orders totaling less than 1% of total biodiversity were grouped into the “other” category.

Figure B.5 Family Comparison

Families totaling less than 1% of total biodiversity were grouped into the “other” category.

Figure B.6 Genus Comparison

Genera totaling less than 1% of total biodiversity were grouped into the “other” category. en test to allow template to find last page in document