Genotyping and Phenotypic Analysis of Streptococcus mutans

Isolates From Children With and Without Dental Caries

by

Aaron Howard Bottner

A thesis submitted in conformity with the requirements

for the degree of Master of Science Orthodontics

Faculty of Dentistry

University of Toronto

© Copyright by Aaron Howard Bottner 2018

ii

Genotyping and Phenotypic Analysis of Streptococcus mutans

Isolates From Children With and Without Dental Caries

Aaron Howard Bottner

Master of Science Orthodontics

Faculty of Dentistry

University of Toronto

2018

Abstract

Within dental plaque, Streptococcus mutans produces acids from fermentable carbohydrates,

decreasing the biofilm pH, causing enamel demineralization. S. mutans produces persisters,

dormant variants of cells that survive lethal antibiotic concentrations without developing antibiotic

resistance. Persisters have been associated with infection chronicity. We hypothesized that

phenotypic heterogeneity of S. mutans is directly associated with cariogenicity. S. mutans were

isolated from plaque collected from caries-free (CF) children and children with Severe Early

Childhood Caries (S-ECC). For each identified genotype, an acid tolerance analysis was

performed and persisters were quantified. In S-ECC patients with >1 strain, the dominant strain

exhibited higher cell survival following exposure to pH 3.2, whereas in CF patients, survival was

similar. S. mutans isolates from S-ECC patients produced ~15x more persisters than isolates from

CF patients. S. mutans exhibits phenotypic diversity. The ability of S. mutans to produce high

levels of persisters may be a hallmark of cariogenicity.

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Acknowledgments

Thank you to my supervisors, Dr. Celine Lévesque and Dr. Siew-Ging Gong, for your support

and guidance toward the completion of my thesis. Your vision and abilities to push both me and

this project further have been crucial.

Thank you to Dr. Hashim Nainar for your direction and expertise in the clinical aspects of this

project. Your experience and insight were critical in making this project a reality.

To Delphine, thank you so much for all your help in lab and for educating me on all things

S. mutans and microbiology. Your passion has truly been inspiring and we could not have

accomplished the scope of this project without your unwavering help.

To the Lévesque lab and summer students: Richard, Myra, and Andrea. This project would be

nothing but frozen plaque samples without your dedication and hard work. Thank you so much

for all your help in the lab. We really could not have done it without you all.

To my parents, Jack and Michelle, and my siblings, Daniel and Leah: Thank you so much for

your help and support over the past three years and for encouraging me to go back to school to

become an orthodontist. We have enough dentists, but finally another Masters in our family!

To my wife, Avra: Thank you, thank you, thank you for everything. Your support and

encouragement have been invaluable over these past three years through all the ups and downs.

Thank you for continually pushing me and helping me at every step along the way. I cannot wait

for the next chapter!

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Table of Contents

Acknowledgments ........................................................................................................................ iii

Table of Contents ........................................................................................................................... iv

List of Abbreviations ..................................................................................................................... vi

List of Tables ................................................................................................................................ vii

List of Figures .............................................................................................................................. viii

1. Introduction ........................................................................................................................... 1

2. Review of the Literature ....................................................................................................... 2

2.1 Epidemiology of Dental Caries in Children ........................................................................ 2

2.2 Classification of Dental Caries in Young Children ............................................................ 2

2.3 Sequelae of Caries Infection in Children ............................................................................ 3

2.3.1 Sequelae of ECC – Costs Associated with ECC ............................................................. 3

2.3.2 Sequelae of ECC – Family-Associated Morbidity .......................................................... 4

2.3.3 Sequelae of ECC – Hospital Costs.................................................................................. 4

2.3.4 Sequelae of ECC – Death ............................................................................................... 4

2.4 Etiology of Dental Caries .................................................................................................... 5

2.4.1 Role of Bacteria in Dental Caries ................................................................................... 6

2.5 Biofilms and Dental Plaque ................................................................................................ 7

2.6 Streptococcus mutans .......................................................................................................... 8

2.7 Role of S. mutans in Dental Caries ..................................................................................... 9

2.8 Genotypic Diversity of S. mutans ..................................................................................... 10

2.9 S. mutans Virulence Factors ............................................................................................. 12

2.10 Acid Tolerance (Acidurance) of S. mutans ....................................................................... 13

2.11 Bacterial Dormancy and Persistence ................................................................................ 15

3. Statement of the Problem ................................................................................................... 18

3.1 Objective ........................................................................................................................... 18

3.2 Aims .................................................................................................................................. 18

3.3 Rationale ........................................................................................................................... 19

3.4 Hypothesis......................................................................................................................... 19

4. Materials and Methods ....................................................................................................... 20

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4.1 Sample Population ............................................................................................................ 20

4.2 Subject Recruitment .......................................................................................................... 20

4.3 Ethics Approval ................................................................................................................ 21

4.4 Plaque Sampling ............................................................................................................... 21

4.5 Mutans streptococci percentage determination in plaque samples ................................... 21

4.6 Identification of S. mutans by PCR ................................................................................... 22

4.7 Genotyping of S. mutans isolates by Arbitrarily-Primed Polymerase Chain Reaction .... 23

4.8 Acid Tolerance Response of S. mutans clinical isolates ................................................... 23

4.9 Persister cell formation by S. mutans clinical isolates ...................................................... 24

4.10 Statistical analysis ............................................................................................................. 24

5. Results .................................................................................................................................. 25

5.1 Subject data ....................................................................................................................... 25

5.2 Mutans streptococci percentage determination in plaque samples ................................... 26

5.3 Identification of S. mutans and genotyping ...................................................................... 28

5.4 Acid Tolerance Response of S. mutans clinical isolates ................................................... 31

5.5 Persister cell formation by S. mutans clinical isolates ...................................................... 33

5.6 Correlations between measures of caries severity, mutans streptococci percentage, acid

tolerance, and persister cell survival ............................................................................................. 35

5.7 Summary of Results .......................................................................................................... 35

6. Discussion ............................................................................................................................ 36

6.1 Sample population data ..................................................................................................... 36

6.2 Mutans streptococci percentage ........................................................................................ 38

6.3 Identification of S. mutans ................................................................................................ 39

6.4 Genotyping S. mutans ....................................................................................................... 40

6.5 Acid Tolerance Response of S. mutans isolates ................................................................ 41

6.6 Persister cell formation of S. mutans isolates ................................................................... 42

7. Conclusions and Future Studies ........................................................................................ 45

8. Appendices ........................................................................................................................... 46

9. References ............................................................................................................................ 57

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List of Abbreviations

AAPD American Academy of Pediatric Dentistry AP-PCR Arbitrarily Primed-Polymerase Chain Reaction ATR Acid Tolerance Response BHI Brain Heart Infusion CA Caries-Active CF Caries-Free CFU Colony Forming Units dmfs Decayed, missing or filled surfaces (primary teeth) dmft Decayed, missing or filled teeth (primary teeth) ECC Early Childhood Caries h hour MS Mutans streptococci MSB Mitis Salivarius Bacitracin NHANES National Health And Nutritional Examination

Survey PBS Phosphate-Buffered Saline PCR Polymerase Chain Reaction s second S-ECC Severe Early Childhood Caries S. mutans Streptococcus mutans TAE Tris-acetate-EDTA THYE Todd Hewitt Yeast Extract

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List of Tables

Table 1. Subject age data.…………………………………………….……….……………… 25 Table 2. Subject caries data for S-ECC group......………………………….………………… 26 Table 3. %MS in individual subjects (top) with mean %MS for all subjects, subjects greater than 3 years of age, and subjects with positive S. mutans identification (bottom).….….….… 27 Table 4. Number of genotypes identified per subject……………………………………...… 30 Table 5. Mean Acid Tolerance Response survival percentage from S-ECC and CF isolates... 32 Table 6. Mean persister cell survival percentages from S-ECC and CF isolates…………….. 33

viii

List of Figures

Figure 1. Mean %MS with standard deviations for S-ECC and CF subjects..………………… 28 Figure 2. Gel showing the amplified PCR product of 479bp using the htrA S. mutans gene-specific primers of MS isolates from Subject #194 (S-ECC group).…………………………... 29 Figure 3. AP-PCR gel demonstrating multiple S. mutans isolates from subject #194.…..….… 30 Figure 4. AP-PCR gel demonstrating the 13 different S. mutans genotypes identified from the 10 S-ECC subjects analyzed..…………………….………….…………...……..………….. 31 Figure 5. AP-PCR gel demonstrating the 9 different S. mutans genotypes identified from the 6 CF subjects analyzed…….………….………….………….………….………….….………... 31 Figure 6. Mean Acid Tolerance Response with standard deviations between S-ECC versus CF S. mutans isolates……………………………………………………………….…….…….. 32 Figure 7. Mean Acid Tolerance Response of isolates with standard deviations indicating survival percentage at non-adaptive pH 3.2.….….…..…………………………………...….…. 33 Figure 8. Mean persister cell survival with standard deviation in S-ECC and CF isolates........ 34 Figure 9. Mean persister cell survival percentages with standard deviations of all isolates…... 34

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1. Introduction

Early Childhood Caries (ECC) is a disease that affects millions of children and their families in

North America and worldwide. The disease process of dental caries is multifactorial, but is

dependent on the metabolism of available sugars in the oral cavity by bacteria, which produce

acids capable of demineralization of enamel and dentin. While several different bacterial species

have been shown to cause dental caries, the most studied and well-recognized of them is

Streptococcus mutans (S. mutans). S. mutans’ cariogenicity is related to several factors including

its ability to produce acids from a variety of carbohydrates (acidogenicity) and its ability to

survive in the acidic environment (acidurance) that is subsequently created. Furthermore, its

ability to form biofilms on tooth surfaces in the form of dental plaque, wherein cell-signalling

can occur, confers upon the bacterial community an increased tolerance for environmental

stressors. One of the mechanisms used by S. mutans colonies to survive in the face of

environmental stressors may be through the formation of persister cells. Persister cells are

physiologically dormant variants of regular non-mutated cells that survive lethal concentrations

of drugs without expressing antibiotic resistance. This state of metabolic dormancy protects

persister cells from environmental stressors, including high doses of antibiotics, until the stressor

is removed, allowing these cells to again become metabolically active and repopulate the colony.

If differences exist in the genotypic and phenotypic characteristics of S. mutans collected from

the plaque of children with and without ECC, then perhaps these differences can be targeted in

such a manner that would eliminate or significantly reduce S. mutans’ cariogenicity. By further

examining the role of acid tolerance and persister cell formation, we can learn more about the

mechanisms by which S. mutans can survive in the harsh environment of the oral cavity, and

ultimately learn to eradicate its role in the propagation of dental caries.

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2. Review of the Literature 2.1 Epidemiology of Dental Caries in Children The U.S. Surgeon General has highlighted that “dental caries is the most common chronic

disease in children: it is about five times as common as asthma and seven times as common as

hay fever” (Benjamin, 2010). The overall prevalence of ECC among Canadian children is less

than 5%, however in high-risk populations, such as First Nations communities, 50–80% of

children are affected (Harrison et al., 1997; Albert et al., 1988; Peressini et al., 2004). National

Health And Nutritional Examination Survey (NHANES) (1999–2002) data showed the high

prevalence of ECC among American children at 2 (10.9%), 3 (20.9%), 4 (34.4%), and 5 (44.3%)

years of age (Centers for Disease Control and Prevention, 2002). More recent data from

NHANES (2011-12) showed that dental caries prevalence in 2–5-year-old children was nearly

23%, with 10% going untreated (Centers for Disease Control and Prevention, 2012; Dye et al.,

2015). In children aged 6-8 years, the prevalence dramatically increased to 56%, with twice as

many (20%) going untreated (Dye et al., 2015). In the United States, children of recent

immigrant backgrounds have three times higher prevalence of caries than non-immigrants (Nunn

et al., 2009). In addition, there is an inverse relationship between socioeconomic status and

caries prevalence in children under age six years of age (Vargas et al., 1998). In general, ethnic

minority children, children from low income families, and children whose parents have less than

a high school education are more likely to experience dental disease and, moreover, are less

likely to receive dental treatment (Henry, 1997).

2.2 Classification of Dental Caries in Young Children The American Academy of Pediatric Dentistry (AAPD) defines the disease of ECC as “the

presence of one or more decayed (noncavitated or cavitated lesions), missing (due to caries), or

filled tooth surface in any primary tooth in a child under the age of six.” ECC can be further sub-

categorized into Severe Early Childhood Caries (S-ECC), which the AAPD defines as “any sign

of smooth-surface caries in a child younger than three years of age, and from ages three through

five, one or more cavitated, missing (due to caries), or filled smooth surfaces in primary

maxillary anterior teeth or a decayed, missing, or filled score of greater than or equal to four

3

(age 3), greater than or equal to five (age 4), or greater than or equal to six (age 5).” (Drury et al.,

1999).

2.3 Sequelae of Caries Infection in Children Casamassimo et al. (2009) constructed a morbidity and mortality pyramid to represent the

sequelae associated with ECC. The pyramid is broken down into four tiers that categorize the

effects of ECC on patients, their families, and society. The hierarchy of the pyramid places the

most severe sequelae on the top and those with descending severity below. From the bottom, the

tiers are: 1) costs associated with ECC, including morbidity associated with treatment, chewing

of the lip or cheek following local anaesthesia, and inappropriate use of pain medications; 2)

family associated morbidity, referring to family stress, loss of work time, loss of school hours

and academic performance, travel and child care costs, eating dysfunction, sleeping dysfunction,

and pain perception; 3) hospital costs, including morbidity resulting from general anaesthesia,

costs of hospital admission, costs of antibiotics and medications, and misuse of emergency

department resources; and 4) death, including mortality related to infection and use of sedation.

2.3.1 Sequelae of ECC – Costs Associated with ECC According to Torabinejad (2009), bacterial infection of the dental pulp, or pulpitis, may

ultimately require endodontic (root canal) treatment or extraction of the tooth. If left untreated,

pulpal necrosis will develop, followed by a dental abscess, leading to destruction of alveolar

bone, with the possible introduction of infectious bacteria into the bloodstream. Furthermore,

facial cellulitis may develop and spread through facial planes to deeper spaces. If unchecked,

infection may progress to involve the orbit, cavernous sinus and, if the mandible is involved,

Ludwig’s angina may develop, compromising the airway (Henry, 1997). Dental intervention in a

hospital setting is typically limited to management of pain and infection, often leaving the source

of the infection untreated with significant cost to the patient’s family (Graham et al., 2000).

Many admissions become prolonged hospitalizations for management of facial cellulitis, which

may last up to five days or longer. (Lin and Lu, 2006) In 2000, the average cost of care across

five children's hospitals for a single admission for odontogenic infection was $3,223 (Ettelbrick,

2000).

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2.3.2 Sequelae of ECC – Family-Associated Morbidity ECC also negatively affects child development and well-being. Reports of parents of children

seeking emergency dental care have shown that children experienced interference with play,

school, sleeping and eating due to dental disease (Edelstein, 2006). Body measurements and

blood test results indicative of malnourishment have been strongly linked with severe ECC and

suggest iron-deficiency anemia (Clarke, 2006). Intermittent pain originating from dental caries

is a consistent finding, affecting up to 20 percent of preschoolers (Edelstein, 2006; Clarke, 2006).

Studies have identified associations between poor systemic and oral health with poor academic

performance at school (Vargas et al., 2005; Blumenshine, 2008).

2.3.3 Sequelae of ECC – Hospital Costs Treatment of paediatric patients under general anesthesia for dental rehabilitation is a costly

consequence of ECC. Dental pain is a leading pediatric admission symptom in many hospitals

emergency departments (Ettelbrick, 2000). Tens of thousands of young children in Canada and

the United States undergo restoration and extraction of teeth under general anaesthesia annually.

The Canadian Institute for Health Information’s Report on Dental Caries demonstrated that ECC

was the leading cause of hospital day surgery for children aged 1-5 years, accounting for 31% of

such procedures. The report concluded that 12.5 ECC-related day surgeries were performed per

1000 children aged 1-5 years, of which 99.6% received general anaesthesia. Furthermore, the

average costs of these procedures ranged from $1,271 to $1,963, depending on the province, with

the annual hospital-related costs attributable to ECC totaling $21.2 million (Canadian Institute

for Health Information Report on Dental Caries, 2013). Extrapolating these costs across the tens

of thousands of children who receive general anesthetic services annually in Canada and the

United States reflects an expenditure of millions of dollars toward treatment of a largely

preventable disease.

2.3.4 Sequelae of ECC – Death Beyond the financial costs, the paediatric population is at the highest risk and has the lowest

tolerance for error for general anaesthetic and sedation procedures (Cravero et al., 2006). While

few have been reported, there are many cases of deaths as a complication of treatment of ECC

5

(Hines, 2008). The absolute mortality associated with ECC will never be fully known on

account of inadequate surveillance, lack of an ECC registry, confidentiality issues, the terms of

legal settlements, and missing or incorrect diagnoses (Casamassimo et al., 2009). Among brain

abscesses, 15% result from infections of unknown source, some of which would include those of

dental origin (Mathisen and Johnson, 1997). In a study attempting to identify paediatric deaths

associated with sedation during an almost 30-year period, Coté et al. (2000) commented that

their study sample demonstrated gross underreporting and that dental specialists were

disproportionately represented among paediatric health care providers. Acetaminophen toxicity

is another concern in pediatric emergency medical care, caused by excessive administration of

the drug by parents for management of pain (Squires, 2006). Toxic doses of acetaminophen can

accumulate rapidly, inducing liver damage in small children (Amar and Schiff, 2007).

2.4 Etiology of Dental Caries Dental caries is a disease process propagated by the bacteria colonizing the oral cavity (Mandel,

1979). Bacteria form complex communities that adhere to tooth surfaces in the form of biofilms,

or dental plaque (Mandel, 1979). Within dental plaque, acidogenic bacteria, such as S. mutans,

produce organic acids from fermentable carbohydrates, which result in a decrease in pH of the

biofilm and saliva and demineralization of enamel (Lingström et al., 2000). The resultant

increase in acidity also favors the growth of bacteria that are able to thrive in acidic conditions

(aciduric bacteria) over other innocuous bacteria that cannot adapt as well to the change in pH

(Selwitz et al., 2007). Acidic dissolution of enamel occurs below the critical pH of 5.5,

removing calcium ions from the crystalline structure of hydroxyapatite (Dong et al., 1999).

However, if the source of the local acid production is removed and the pH in the vicinity of the

demineralized tooth structure increases, remineralization of enamel can spontaneously occur

(Zero, 1999). According to Iijima et al. (1999), remineralization occurs when calcium and

phosphate ions in the saliva crystalize to repair damaged hydroxyapatite crystals. If fluoride is

present during the remineralization process, fluoride ions will become incorporated into the

crystalline structure of hydroxyapatite replacing its hydroxyl groups resulting in a stronger, less

porous, and more acid-insoluble form called fluorapatite (Ca10(PO4)6F2) (Zero, 1999; Iijima et

al., 1999). Caries develops through interrupted and imbalanced episodes of demineralization and

remineralization (Seltwitz, 2007). Caries development is therefore directly dependent upon the

6

presence of acidogenic bacteria on the tooth surface, available fermentable carbohydrates, and

the ability of salivary factors to remineralize the tooth structure following acidic

demineralization (Kidd, 2004).

At the structural level, caries risk can be influenced by several different factors. Featherstone

(2006) illustrated the relative contribution of pathological and protective factors through the

concept of caries balance. Pathological factors include the presence of cariogenic bacteria,

presence of fermentable carbohydrates, and salivary dysfunction. Protective factors include

saliva components and flow, extrinsic fluoride, antibacterial therapy, and oral hygiene. A

reduction in pathological factors and an increase in protective factors have been shown to reduce

relative caries risk (Featherstone, 2006). Studies in genetic twins have shown a genetic

component to the development of dental caries despite being raised in different locations and

environments (Conry et al., 1993). Genetic contribution has been long recognized for

involvement in pH of saliva (Turner et al., 1953), serum levels of immunoglobulins and

antibodies (Allansmith et al., 1969), and activity of salivary amylases (Goodman et al., 1959).

2.4.1 Role of Bacteria in Dental Caries Analyses within cariogenic dental plaque have demonstrated the presence of many bacterial

species including Streptococcus, Lactobacillus, Actinomyces, Veillonella, Granulicatella,

Leptotrichia, Thiomonas, Bifidobacterium, and Scardovia (Aas et al., 2008; Becker, 2002).

Since all the bacteria that have been associated with caries belong to the normal microflora of the

oral cavity, dental caries has been described as an endogenous infection (Fejerskov and Nyvad,

2003). When the normal homeostatic balance of the biofilm is disturbed, endogenous infections

can occur when certain bacteria within the flora have a selective advantage over other species,

suggesting an ecological hypothesis to the process of caries development (Marsh and Martin,

1999). Dental plaque is a dynamic microbial ecosystem in which non-mutans bacteria, such as

Actinomyces and Streptococcus other than S. mutans, maintain dynamic stability and produce

acids from sugary foods capable of demineralizing enamel. However, the resultant temporary

decreases in pH, shown to be a part of the normal pH cycle, are easily returned to neutral level

by homeostatic mechanisms in the plaque and occur numerous times daily in supragingival

plaque (dynamic stability stage) (Marsh and Martin, 1999; Takahashi and Nyvad, 2008).

7

However, with frequent supply of sugar or when salivary secretion is limited to neutralize the

acids produced, the pH decrease in the plaque may enhance the acidogenicity and acidurance of

the non-mutans bacteria adaptively. Under such conditions the populations of the ‘low-pH’ non-

mutans streptococci and Actinomyces increase via acid selection, leading to a microbial shift to a

more acidogenic microflora. These changes in the genotype and phenotype of the microflora may

shift the demineralization/remineralization balance from ‘net mineral gain’ to ‘net mineral loss’

and initiate lesion development (acidogenic stage). At this early stage, lesion development could

be arrested with de-adaptation of the microflora, provided that the mineral balance is restored to

a ‘net mineral gain’ by reduced environmental acidification, such as with sugar restriction and

oral hygiene measures. In these environments, more aciduric bacteria such as S. mutans and

lactobacilli may replace the ‘low-pH’ non-mutans bacteria and further accelerate the caries

process (aciduric stage). Even at this highly aciduric stage, the mineral balance and composition

of the microflora may be reversible through modification of the acidic environment

(de Stoppelaar et al., 1970).

2.5 Biofilms and Dental Plaque Biofilms are microbial communities attached to surfaces and encased in an extracellular matrix

of microbial origin (Costerton et al., 1978). Biofilms represent a way in which bacteria are able

to survive in the face of stressful, hostile environments (Hall-Stoodley et al., 2004) and are a

major cause of many chronic human infections, including pneumonia, endocarditis, skin

infections, chronic otitis media, periodontal disease, dental caries, and numerous others

(Costerton et al., 1999; Marsh, 2006). Extensive clinical studies have indicated that the oral

microbial flora, primarily in the form of dental plaque, is responsible for two major biofilm-

related diseases: dental caries and periodontal disease (Kuramitsu et al., 2007). Dental plaque is

known to be composed of more than 700 different bacterial species and is among the most

researched biofilms (Marsh, 2006). The precursor to dental plaque, the acquired enamel pellicle,

forms within minutes after a professional cleaning (Skjørland et al., 1995). The acquired enamel

pellicle is an integument composed almost entirely of oral, fluid-derived proteins, which

originate from saliva and the gingival crevicular fluid (Dawes et al., 1963; Heller et al., 2017).

The biological role of the pellicle is its protective effect on tooth surfaces against acid insults and

abrasion, and its role in guiding the attachment of early microbial colonizers (Kolenbrander and

8

London, 1993). The proteins that compose the pellicle provide the necessary binding receptors

for the initial colonizers, most of which are beneficial to the oral health of the host, including

primarily Streptococci such as S. oralis, S. gordonii, and S. sanguinis and, with a lesser

frequency, Actinomyces (Socransky et al., 1998; Li et al., 2004; Heller et al., 2017). The bacteria

comprising the dental plaque biofilm are not randomly distributed and display a high degree of

organization formed by sequential and ordered colonization of multiple species of bacteria

(Marsh, 2006; Marsh, 1994; Socransky et al., 1998; Kolenbrander et al., 2006). Middle

colonizers, such as S. mutans, coaggregate with early colonizers, followed by late colonizers,

which then bind to already attached early and middle colonizers (Kolenbrander, 2011). This

increasing bacterial adhesion culminates into microbial succession in species diversity and the

maturation of the complex biofilm community (Periasamy and Kolenbrander, 2009).

2.6 Streptococcus mutans S. mutans was first isolated and identified from carious lesions in human subjects by Clarke in

1924, who used the term mutans to describe the oval-shaped cells he observed microscopically

as mutant forms of streptococci (Clarke, 1924). The Streptococcus genus as a whole is

considered one of the most invasive groups of bacteria (Krzyściak et al., 2013). The genus

consists of greater than 50 species and subspecies, many of which have been identified as

causative agents in human infections (Facklam, 2002). Streptococci commonly found in the oral

cavity include the anginosus, mitis, salivarius, and mutans groups, with some additional species

not fitting the criteria of the aforementioned groupings (Whiley and Beighton, 1998). The

mutans group includes S. mutans along with S. sobrinus, S. criceti, and S. ratti, all of which have

been found in humans, while other members, such as S. macacae, S. downei, and S. ferus, have

not (Whiley and Beighton, 1998). Oral streptococci can be alpha-hemolytic or gamma-

hemolytic (Facklam, 2002). S. mutans is a facultative anaerobic, Gram-positive, coccus-shaped

bacterium, whose genome was first sequence by Ajdić et al. in 2002 (Ajdić et al., 2002). They

determined that the UA159 strain of S. mutans is composed of 2,030,936 base pairs and 1,963

genes (Ajdić et al., 2002), which is approximately half the size of the genome of Escherichia

coli. More recent techniques used by Cornejo et al. (2012) obtained the genome sequences of 57

clinical isolates of S. mutans, who estimated the S. mutans core genome to contain close to 1,500

genes, with each genome containing, on average, 1636 genes (Cornejo et al., 2012). S. mutans

9

colonies grown on 5% sucrose agar frequently present as white and occasionally yellow, rough,

heaped, and detachable from the agar surface (de Stoppelar et al., 1967). The species in the

mutans group are capable of producing acids from a wide range of carbohydrates, including N-

acetylglucosamine, esculin, amygdalin, arbutin, cellobiose, galactose, inulin, lactose, maltose,

mannitol, melibiose, raffinose, salicin, sorbitol, tagatose, and trehalose (Whiley and Beighton,

1998). While most commonly studied for its role in the pathogenesis of dental caries, S. mutans

has also been demonstrated as a cause of infective endocarditis, particularly the serotype f and k

strains (Sato et al., 2004; Nomura et al., 2012).

2.7 Role of S. mutans in Dental Caries S. mutans is thought to play a critical role in the pathogenesis of dental caries and is considered

the most notorious bacterial species involved in caries development (Caufield et al., 1993;

Becker et al., 2002; Beighton et al., 2004; Ge et al., 2008). It is found in a high proportions of

subjects with dental caries and has been extensively characterized for its role in caries

development and formation (Van Houte, 1980; Loesche, 1986; Ge et al., 2008; Palmer et al.,

2010; Mitrakul et al., 2016). Marchant (2001) found that in children with ECC, S. mutans

represented a significantly greater proportion of carious lesion flora, whereas S. oralis, S.

sanguinis and S. gordonii formed a significantly greater proportion of the plaque flora from

caries-free (CA) tooth surfaces. A three-year cohort study in children older than 2.5 years of age

demonstrated a significant correlation between the clinical caries score and presence of S. mutans

in plaque or saliva (Roeters, 1995). When plaque was removed from single occlusal fissures,

71% of' the carious fissures had S. mutans accounting for more than 10% of' the viable flora,

whereas 70% of' the CF fissures had no detectable S. mutans (Loesche et al., 1975). Multiple

bacteriologic studies have demonstrated that in children with ECC, S. mutans frequently

exceeded 30% of the cultivable plaque flora (Van Houte et al., 1982; Berkowitz, 1984; Milnes

and Bowden, 1985). Direct inoculation of hamsters and gnotobiotic rats in research conducted in

the 1950s and 60s established S. mutans as a causative agent of dental caries (Fitzgerald &

Keyes, 1960; Orland et al., 1955; Zinner et al., 1965). S. mutans has also been shown to directly

cause caries in germ-free and specific pathogen-free rat models (Michalek and McGhee, 1977).

Multiple studies have shown that the relative levels of S. mutans and other bacterial species may

10

be important in caries development, with S. sanguinis, S. gordonii, and other bacteria playing

protective roles (Li et al., 2007; Ge et al., 2008; Kreth et al., 2008).

2.8 Genotypic Diversity of S. mutans Within the S. mutans species, a considerable amount of genotypic diversity exists, which was

first highlighted by Kulkarni et al. (1989) via restriction fragment length polymorphism analysis

from S. mutans strains isolated from different individuals. Since then, microarray-based

comparative genomic hybridization techniques have shown that regardless of the geographical,

temporal or familial context considered, between 15 % and 20 % of the total open reading frames

identified in the S. mutans UA159 genome were absent or divergent in other strains (Waterhouse

et al., 2007; Zhang et al., 2009). Further sequencing of additional S. mutans genomes have

supported this genotypic divergence with the finding that the more genomic sequences released,

the greater the divergence observed (Maruyama et al., 2009; Cornejo et al., 2012; Song et al.,

2013; Argimon et al., 2014). As of this writing, 183 S. mutans genome sequences have been

released with an average genome size of 1.95 Mb and an average of 1,820 protein-coding genes

(Meng et al., 2017). Comparison of the protein-coding genes of all 183 S. mutans strains

highlights the presence of a pan-genome (a full set of different genes discovered in the species)

composed of more than 4000 genes and a core-genome (genes shared by all strains of the

species) composed of only 1083 genes (Meng et al., 2017). Consequently, 75% of the S. mutans

genes identified so far are strain-specific, emphasizing the vast genotypic diversity of this

bacterial species (Meng et al., 2017). Other faster and less expensive techniques have also been

used to highlight the genotypic diversity of S. mutans, including the arbitrarily primed-

polymerase chain reaction (AP-PCR) (Jiang et al., 2012; Arthur et al., 2007; Gilbert et al., 2014).

AP-PCR studies have also revealed that a single individual may carry more than one S. mutans

genotype (Jiang et al., 2012; Gamboa et al., 2010; Valdez et al., 2017). While some studies have

found a positive correlation between the number of bacterial genotypes and the presence of

dental caries (Napimoga et al., 2004; Alaluusua et al., 1996; Pieralisi et al., 2010), others do not

corroborate this finding and have found no correlation between genotypic diversity and caries

experience (Kreulen et al., 1997; Lembo et al., 2007; Damle et al., 2016). This observed

correlation between cariogenicity and genotypic diversity may be attributed to the simultaneous

11

action of several strains with differing cariogenic potential with complimentary virulence factors,

however this is speculative (Alaluusua et al., 1996; Pieralisi et al., 2010).

Alaluusua et al (1996) first demonstrated that genotypic differences between strains of S. mutans

were related to disparities in their cariogenicity. This genotypic diversity has led researchers to

search for genotypes or genetic loci specific to S. mutans strains isolated from CA versus CF

teeth, but as of yet, no clear correlation has been established (Jiang et al., 2012; Meng et al.,

2017). Mitchell et al. (2009) proposed that over time, the oral cavity of individuals with S-ECC

undergoes a continuous and progressive re-distribution of S. mutans strains, influenced by

several risk factors, including caregivers containing high S. mutans bacterial numbers or

untreated carious lesions, frequent ingestion of sucrose-containing food or drink, and poor oral

hygiene. The product of these factors can lead to earlier S. mutans colonization and increased

S. mutans strain diversity at higher concentrations in children with S-ECC as compared to CF

children (Napimoga et al., 2004; Napimoga et al., 2005; Lembo et al., 2007; Gilbert et al., 2014).

The S. mutans genotypes found in CA individuals may be influenced by environmental factors,

such as increased acidity due to the metabolism of frequently consumed sugary foods, allowing

for the survival of the strains that are capable to tolerate such conditions (Lembo et al., 2007).

Saxena et al. (2008) demonstrated differences in the genomic composition of S. mutans strains

associated with S-ECC compared to CF controls and employed an artificial intelligence

algorithm to predict which combinations of genetic fragments would best predict the presence of

caries. The presence of specific DNA fragments in 90-100% of the 26 S-ECC isolates tested in

the study, suggests their involvement in the pathogenesis of S. mutans associated with dental

caries and their potential for use as biomarkers with predictive power for disease (Saxena et al.,

2008). The genes involved in these biomarkers are responsible for different cellular processes,

such as sucrose modulation (Macrina et al., 1991), mutacin production (Kamiya et al., 2005),

serotypic antigens (Shibata et al., 2003), and adhesion (Yamashita et al., 1992). Gilbert et al.

(2014) used AP-PCR to identify specific S. mutans strains present from carious lesions, white

spot lesions, and non-carious enamel surfaces in S-ECC subjects, and observed primary

S. mutans strains isolated from carious lesions that were distinct from dominant strains found on

enamel. The putative virulence genes of S. mutans are broadly distributed, both in strains

isolated from caries-active (CA) and CF individuals, however their relative expression may

12

differ (Meng et al., 2017). The genotypic diversity and the absence of definitive caries-specific

genetic markers make S. mutans extremely difficult to target, suggesting that other factors, both

environmental and host-related, are contributing to the cariogenicity of the species (Meng et al.,

2017).

2.9 S. mutans Virulence Factors The virulence and cariogenicity of S. mutans is multifactorial, but is largely related to its ability

to form and thrive in biofilms (Hamada et al., 1984; Napimoga et al., 2005; Zhu et al., 2006). In

order to form biofilms, adhesion of bacteria to the acquired enamel pellicle and to one another is

required, which S. mutans can accomplish via both sucrose-dependent and sucrose-independent

mechanisms in vitro (Zhu et al., 2006). The sucrose-dependent method involves the production

of extracellular polysaccharides (glucans) from the glucose moiety of sucrose, through the

enzymatic activity of three glucosyltransferases (GtfB, -C and -D). Glucans allow S. mutans to

firmly adhere to smooth surfaces of teeth (Kuramitsu, 1993; Yamashita et al., 1993) and the

glucan-binding proteins (i.e. GbpA, -B, -C and -D) are thought to play important roles in

subsequent cell–cell aggregation and biofilm development (Douglas & Russell, 1982; Shah &

Russell, 2004; Smith & Taubman, 1996). The sucrose-independent mechanism is less

understood, but involves the function of the wall-associated protein A (WapA) (Levesque et al.,

2005; Zhu et al., 2006). Saliva and plaque samples of S. mutans from CA children have been

shown to have an increased biofilm-forming capacity compared to those sampled from CF

children, suggesting the importance of WapA in cariogenic strains (de Camargo et al., 2018;

Valdez et al., 2017). Another important protein involved in mediating binding of S. mutans to

salivary components on tooth surfaces is the cell surface protein antigen c (PAc), also known as

antigen I/II, B, IF, P1, SR, or MSL-1 (Yu et al., 1997; Koga et al., 1990). The significance of

PAc in allowing S. mutans to colonize tooth surfaces is such that its disruption has been

considered as a target for a possible anti-caries vaccine (Russell et al., 2004). Acidogenicity, the

ability to produce organic acids by fermentation of dietary carbohydrates, is a critical virulence

factor of S. mutans as it causes the direct demineralization of tooth structure (Banas, 2004).

S. mutans is capable of producing lactate, formate, acetate, and ethanol as fermentation products

(Ajdic et al., 2002) depending on growth conditions, with lactate being the major product when

glucose is abundant (Dashper and Reynolds, 1996). Strains with reduced L+ lactate

13

dehydrogenase production, the enzyme responsible for the conversion of pyruvate to lactic acid,

display reduced cariogenicity and their use has been suggested for replacement therapy (Johnson

et al., 1980; Fitzgerald et al., 1989; Hillman, 2002). As with other virulence factors, the relative

acidogenicity of S. mutans can vary from one strain to another, however correlations between

acidogencity and caries experience have not been demonstrated (Köhler et al., 1995; Napimoga

et al., 2004; Mattos-Graner et al., 2000; Valdez et al., 2017). Regardless, the acidogenicity of

S. mutans is thought to lead to ecological changes in the plaque flora involving a decrease in pH,

thereby increasing the proportions of S. mutans and other acid-tolerant species (Banas, 2004).

S. mutans is also known to produce relatively high quantities of bacteriocins (or mutacins) and

bacteriocin-like inhibitory substances (Balakrishnan et al., 2002; Hale et al., 2005). Bacteriocins

are peptides or protein products that are bactericidal for other bacteria of the same or closely

related species, and help to establish an ecological advantage in diverse bacterial communities

such as dental plaque (Balakrishnan et al., 2002). In addition to the virulence factors described

above, S. mutans cariogenicity is also related to its acidurance and ability to form persister cells,

which will be described in greater detail subsequently. The relationship between S. mutans’

cariogenicity and its link to persister cell formation has yet to be studied until this investigation.

2.10 Acid Tolerance (Acidurance) of S. mutans S. mutans and other bacteria in the oral cavity experience a wide range of environmental stresses,

particularly the intermittent ingestion of food by the host, which results in sudden dramatic

changes in pH, nutrient availability, oxygen tension and osmolality (Lemos et al., 2005). The

decrease in pH to below 5.5 following sugar intake requires acidurance: the ability of bacteria to

develop mechanisms to survive when exposed to these harsh acidic conditions (Palmer et al.,

2013). S. mutans and other cariogenic bacteria, such as S. sobrinus and Lactobacillus

acidophilus have been suggested to proportionally increase in dental plaque of patients, in large

part due to their acidurance (Marsh, 1991; Bowden and Hamilton, 1987). Within the

mechanisms utilized to adapt to a variety of sudden environmental changes is the close

connection between responses to environmental stressors and increased biofilm formation

(Lemos and Burne, 2008). This suggests that the stress regulon of S. mutans controls responses

to a variety of stressors via a well-coordinated, community-driven approach (Lemos and Burne,

2008). McNeill and Hamilton (2003) found that biofilms of S. mutans exhibited improved acid

14

resistance as compared to planktonic cells, and were capable of a strong physiological response

to acid stress. This difference in acid tolerance of the biofilm as compared to dispersed cells

reflects the value in the physical structure of the biofilm in enhancing cellular systems for growth

and pH homeostasis. S. mutans utilizes multiple mechanisms of acid tolerance, such as activity

of the membrane-bound enzyme F-ATPase (Bender et al., 1986; Lemos et al., 2005; Sheng and

Marquis, 2007), the agmatine deiminase system (Griswold et al., 2006), and malolactic

fermentation (Sheng and Marquis, 2007). Many of the stresses encountered by oral bacteria

induce damage to DNA and other macromolecules (Lemos and Burne, 2008). As a result,

various proteins and gene pathways appear to be essential for protection and repair of

macromolecules that can be damaged due to the stressors of the oral cavity, which include

GroEL and DnaK chaperones (Lemos et al., 2007), RopA (Wen et al., 2005), repression of the

surface-associated protease HtrA or the cytoplasmic ClpP peptidase (Ahn et al., 2005; Biswas &

Biswas, 2005; Deng et al., 2007), among several others. S mutans also responds to

environmental acidic stress via cell envelope alterations, including changes in proportions of

membrane fatty acids (Fozo and Quivey, 2004), translocation and assembly of membrane

proteins (Hasona et al., 2005), and cell wall biogenesis (Wen et al., 2006). The ffh gene also

appears to have a role in the acidurance of S. mutans, due to its involvement in the maintenance

of a functional membrane protein composition during adaptation to changing environmental

conditions (Gutierrez et al., 1999).

Some researchers have postulated that acidurance may be a critical phenotypic difference

between S. mutans strains present in CA versus CF individuals due to the more acidic

environment brought on by bacterial metabolism of ingested sugars (Harper and Loesche, 1984;

Takahashi and Nyvad, 2008). Greater lactic acid concentrations, the primary acidic metabolic

end product in dental plaque exposed to excess of sugars, have been demonstrated in CA than

caries-inactive subjects (Geddes, 1974; Gao et al., 2001; Margolis et al., 1994). Caries activity

has also been associated with a faster pH decrease and a lower minimal pH in dental plaque (Van

Houte et al., 1991; Georgios et al., 2015). Therefore, S. mutans strains with greater acid

tolerance, brought on by changes to the aforementioned acidurance-related genes, would be

expected to have an improved ability to survive the oral cavities of CA individuals, causing

further development and progression of caries (Lembo et al., 2007). Acid Tolerance Response

15

(ATR) is an adaptive response wherein bacteria that are exposed to initial sub-lethal acidic

conditions enhance their survival at a lower “lethal” pH (Welin-Neilands and Svensäter, 2007).

Under acidic conditions, S. mutans alters its physiology in a variety of ways in order to survive

and prepare for the next acid challenge (Welin-Neilands and Svensäter, 2007). S. mutans adapts

by the synthesis of stress-responsive proteins, increasing glycolytic activity and increasing the

activity of the proton-translocating ATPase regulating intracellular pH (Bender et al., 1986;

Hamilton and Ellwood, 1978; Hamilton and Buckley, 1991; Svensäter et al., 2000). The initial

exposure to a sub-lethal pH is critical to this process, as previous studies have shown that very

few cells can survive sudden drops in pH (i.e. from 7.5 to 3), however survival is enhanced upon

an initial exposure to the sub-lethal pH 5.5 (Svensäter et al., 1997; Welin et al., 2003). Previous

studies have also indicated that the ATR is significantly enhanced in biofilm cells as compared to

planktonic cells with increased acid-tolerant protein expression, indicating that bacteria are more

resistant to acid stress when part of a biofilm community (Welin et al., 2003; McNeill and

Hamilton, 2003; Svensäter et al., 2001). Utilizing an ATR protocol, Valdez et al. (2017) found

that S. mutans isolates from S-ECC subjects demonstrated a significantly higher percentage of

bacterial survival at pH 2.8 as compared to the isolates from CF subjects. Jiang et al. (2017)

found that an S. mutans strain isolated clinically from a CA adult exhibited greater acidurance

compared to strains isolated from a CF adult and a reference strain. Lembo et al. (2007) also

found that genotypes with high acid tolerance were more common among CA than CF isolates.

These studies all point to greater acidurance in S. mutans strains isolated from individuals with

dental caries, suggesting the importance of this virulence factor in cariogenic bacteria.

2.11 Bacterial Dormancy and Persistence Bacterial persistence refers to a phenomenon where within a bacterial population, a small

subpopulation of cells, called persisters, enters into a state of dormancy (Bigger, 1944; Lewis,

2010; Jayaraman, 2008). Persister cell formation can occur via both stochastic and deterministic

mechanisms (Jayaraman, 2008). Stochastic formation of persister cells may arise as a result of

random intrinsic fluctuations of protein levels (Yamaguchi et al., 2011) and spontaneous gene

expression during transcription and translation (Kint et al., 2012). Persister cells are also formed

deterministically when bacterial populations are exposed to environmental stressors (chemical

and physical) (Vega et al., 2012; Lewis 2008). The mechanism to form persisters (or persister

16

cells) is different from antibiotic resistance in that these cells are not mutants, but are phenotypic

variants of the wild-type strain that arise in a clonal population of genetically identical cells

(Levin and Rozen, 2006). Persisters are dormant cells that neither grow nor die in the presence of

high doses of bactericidal antibiotics (Lewis, 2008). By entering into a growth-arrested

physiological state, persisters are shutting down the activity of essential cellular processes

targeted by antibiotics allowing them to survive without expressing a drug resistance mechanism

(Lewis, 2010; Wood et al., 2013). Persistence has been observed in multiple bacterial species,

such as Escherichia coli, Staphylococcus aureus, Mycobacterium tuberculosis, Pseudomonas

aeruginosa, and S. mutans. Persisters arise in biofilm and in planktonic cells, and are among the

main factors responsible for the tolerance of pathogens to antibiotics (Spoering and Lewis,

2001). Persisters play a crucial role in biofilm resilience, as the survival of just a few persisters

can repopulate the biofilm, causing the infection to relapse (Dufour et al., 2012; Lewis, 2008;

Balaban et al., 2013). Persister formation can be induced following exposure to stress or by

quorum sensing molecules, such as phenazine pyocyanin and acyl-homoserine lactone in

P. aeruginosa, indole in E. coli and Salmonella typhimurium, and CSP pheromone in S. mutans

(Maisonneuve and Gerdes, 2014). Quorum sensing, a mechanism for intra- and inter-species

communication between bacteria, serves to prime the bacterial population’s cellular response to

stress and ensure survival by inducing the formation of antibiotic-tolerant persisters

(Maisonneuve and Gerdes, 2014). The prevention of persister formation by interfering with the

quorum sensing system may represent a possible drug targets for the development of effective

antimicrobial strategies (Leung and Levesque, 2012). Several algae and terrestrial plants

produce compounds able to interfere with bacterial quorum sensing, whose further research may

lead to promising pharmaceutical agents (Defoirdt et al., 2010).

S. mutans has been shown to produce substantial numbers of persisters when growing in an in-

vitro biofilm (Leung and Levesque, 2012; Leung et al., 2015). Older biofilms (72-hour-old)

biofilms produced more persisters than younger biofilms (6-hour-old or 24-hour-old), suggesting

that more persisters could be formed due to gradually limiting conditions prevailing in the

biofilm (Leung and Levesque, 2012). Studies of S. mutans persisters have identified multiple

genes that have been implicated in bacterial persistence, including toxin-antitoxin systems, genes

involved in transcription/replication, sugar metabolism, cell wall synthesis, and energy

17

metabolism (Leung et al., 2015; Koyanagi et al., 2013). Oral bacteria are constantly exposed to a

wide range of stresses (e.g., constant cycles of famine and feast, fluctuations in pH, temperature

shock, oxidative stress) (Marsh et al., 2006; Vega et al., 2012). In S. mutans, the onset of

environmental stresses encountered in the oral cavity also induce the formation of persisters

(Leung and Levesque, 2012). Jiang et al. (2017) induced persister cell formation in S. mutans by

exposing colonies to different antibiotics and to an antibacterial quaternary ammonium

compound (dimethylaminododecyl methacrylate). They monitored several metabolism-related

genes and found that these genes were downregulated as compared to the control cells. This

metabolically dormant state is a hallmark of persister cells, and they suggested that activation of

their metabolism via a carbon source may be able to “awaken” the persister cells, making them

susceptible to antibacterial agents (Jiang et al., 2017). While it has been shown that S. mutans

does indeed produce persister cells when faced with environmental stressors, the exact

mechanisms under which this phenomenon occurs are still not well understood (Leung et al.,

2015). Targeting persister cell formation could prove to be a critical strategy in the fight against

dental caries, as it would combat the ability of S. mutans to repopulate following exposure to

antimicrobials, thereby hindering the recurrence of the infection (Leung et al., 2015).

Combatting persistence would be particularly useful if it could be shown to occur more in high-

risk populations, such as children with ECC, where recurrence of carious lesions is particularly

common (Amin et al., 2010).

18

3. Statement of the Problem

Dental caries is a chronic infectious disease with many associated morbidities and costs to

patients, their families, and society. S. mutans is consistently identified as one of the major

etiological bacteria involved in the propagation of this disease. It may be postulated that the

recurrent nature of caries, particularly within high-risk populations, is in part due to the highly

aciduric nature of S. mutans and its ability to form persister cells. If an increase in S. mutans

persisters is identified in children with S-ECC, it may provide a new target in the management

and prevention of carious lesions in high-risk populations.

3.1 Objective To investigate genotypic and phenotypic differences that exist between the S. mutans sampled

from children with and without S-ECC.

3.2 Aims

1. To acquire dental plaque from children 71 months of age and under who are classified as

having S-ECC and those who are CF.

2. To determine the percentage of mutans streptococci from the total cell count in the subjects’

plaque samples.

3. To confirm the presence of S. mutans and to isolate S. mutans from the subjects’ plaque

samples.

4. To determine the genotypic diversity in S. mutans isolated from the subjects’ plaque samples.

5. To compare the ATR of the S. mutans isolated from subjects with S-ECC and CF children.

6. To compare the ability of S. mutans strains isolated from the plaque of children with S-ECC

and CF children to form persister cells.

19

3.3 Rationale The significance of this study is to determine if differences exist in the phenotypic traits of

acidurance and persistence between S. mutans strains isolated from the plaque of children with

S-ECC and those who are CF. This study is novel in that it is the first to evaluate the ability of

S. mutans to form persister cells in strains isolated from children with S-ECC and CF children.

3.4 Hypothesis

S. mutans strains isolated from children with S-ECC will demonstrate increased acidurance and

higher proportions of persister cells than S. mutans strains isolated from CF children.

20

4. Materials and Methods 4.1 Sample Population Inclusion Criteria for S-ECC group:

- Younger than six years of age (≤71 months)

- Subjects in primary dentition, prior to the eruption of permanent teeth

- Healthy children with non-contributory medical history

- Undergoing treatment at the Paediatric Clinic at University of Toronto, Faculty of

Dentistry

- Must meet the AAPD’s definition of S-ECC:

o In children younger than three years of age, any sign of smooth-surface caries

o From ages three through five, one or more cavitated, missing (due to caries), or

filled smooth surfaces in primary maxillary anterior teeth or a decayed, missing,

or filled score of greater than or equal to four (age 3), greater than or equal to five

(age 4), or greater than or equal to six (age 5)

Inclusion criteria for CF group:

- Younger than six years of age (≤71 months)

- Subjects in primary dentition, prior to the eruption of permanent teeth

- Healthy children with non-contributory medical history

- Zero dmft

- Bitewing radiographs taken, if necessary, within the past year to confirm absence of

carious lesions

4.2 Subject Recruitment Subjects were recruited from the Graduate Paediatric Dentistry Clinic at the University of

Toronto, Faculty of Dentistry. Subjects were examined by the paediatric dentistry resident and

A. Bottner to ensure they met the inclusion criteria for one of the groups in the study.

21

4.3 Ethics Approval This study obtained approval from the University of Toronto Research Ethics Board on March

21, 2016 (Protocol Reference Number 32740).

4.4 Plaque Sampling A consent form was reviewed and signed by each subject’s parent/guardian, which outlined the

study’s protocol, and indicated that their participation in this study will have not effect on their

treatment or standing in the waitlist for treatment at the Faculty. It also indicated that no

additional care was being provided because of participation in this study. A short questionnaire

collected patient information including the patient’s age in months, gender, whether they were

born prematurely, and whether they were born via vaginal birth or Caesarian section. In

addition, an odontogram was completed to indicate decayed, missing and filled surfaces. This

information was collected and analyzed in accordance with the University of Toronto, Faculty of

Dentistry’s Privacy Policies.

Plaque was collected from facial and lingual smooth surfaces of primary maxillary incisors,

using a sterile toothpick and placed in a capped microcentrifuge tube. Each tube was labelled

with a participant number assigned at the time of participant selection. The samples were

immediately stored in a secured −80°C freezer until analyzed.

4.5 Mutans streptococci percentage determination in plaque samples Plaque samples were thawed and resuspended in 300µL of sterile Phosphate-Buffered Saline

(PBS, pH 7.2). Tubes were vortexed for 30 s for content homogenization. Each sample was

serially diluted in PBS and plated on both Brain Heart Infusion (BHI) agar, to quantify the total

bacterial cell count, and Mitis Salivarius Bacitracin (MSB) agar, to quantify the number of

mutans streptococci (MS) cells, using an automated spiral plater for colony forming unit (CFU)

determination. Plates were then incubated at 37°C in air with 5% CO2 for 48 h for BHI and 72 h

for MSB and CFU were determined. The percentage of mutans streptococci (%MS) from each

sample was determined by the following calculation:

22

4.6 Identification of S. mutans by PCR Colonies of S. mutans grown on MSB agar were isolated and re-patched on BHI plates with a

5x5 grid in order to establish 25 isolates per subject. Colony PCR was performed using a set of

primers specific to S. mutans, targeting the intergenic region between the htrA gene and

SMU.2165 (CMT-1036: 5' TGC CGA AAA AGA TAA ACA AAC A 3'; CMT-1037: 5' GCC

CCT TCA CAG TTG GTT AG 3'). The htrA gene encodes a heat-shock induced surface

protease that is essential for stress tolerance and is a regulator involved in cellular growth, stress

tolerance, biofilm formation, competence development, and genetic transformation (Ahn et al.,

2005). The reactions were processed in 20µL mixtures containing 1x Taq buffer, 0.2mM dNTP,

1µM of each of CMT-1036 and CMT-1037 primer, 2mM MgCl2 and 0.02 units/µL of Taq DNA

polymerase. Reactions were performed with one initial cycle of denaturation at 94°C for 5 min,

followed by 30 cycles of 94°C for 30 s (denaturation), 56°C for 30 s (annealing) and 72°C for 1

min (extension), and a final extension at 72°C for 10 min. S. mutans strain UA159 was used as a

positive control.

The PCR products were analyzed in a 1g/100mL agarose gel in Tris-acetate-EDTA buffer and

stained with ethidium bromide solution. The image of the gel was capture by a digital imaging

system (BioRad Gel Doc System). A 1-kb DNA ladder served as a molecular size marker in the

gel.

Analysis of early collected samples revealed difficulty in isolating S. mutans from the CF group,

whereas S. mutans isolates were detected in all S-ECC samples. S. mutans was isolated in only

six of the twenty-one CF samples. Therefore, for subsequent experiments, only the first ten S-

ECC samples collected and the 6 S. mutans-containing CF samples were used, in order to have

similar group sizes to analyze the genotypic and phenotypic properties of S. mutans isolates.

%MS = CFU/mL on MSB x100 = MS cells CFU/mL on BHI Total cells

23

4.7 Genotyping of S. mutans isolates by Arbitrarily-Primed

Polymerase Chain Reaction Arbitrarily Primed-Polymerase Chain Reaction (AP-PCR) assays were performed for each

isolate with the arbitrary primer OPA-02 (5ʹ- TGCCGAGCTG -3ʹ) (Tabchoury et al., 2008). The

reactions were processed in 20µL mixtures containing 1x Taq buffer, 0.2mM dNTP, 2µM OPA-

02 primer, 7mM MgCl2, 1ng/µL of genomic DNA, and 0.05 units/µL of Taq DNA polymerase.

Reactions were performed with one initial cycle of denaturation at 94°C for 2 min, followed by

45 cycles of 94°C for 30 s (denaturation), 36°C for 30 s (annealing) and 72°C for 1 min

(extension), and a final extension at 72°C for 5 min. The amplicons were evaluated in a

1.5g/100mL agarose gel in Tris-acetate-EDTA (TAE) buffer by a digital imaging system (Biorad

Gel Doc System). A 1-kb DNA ladder served as a molecular size marker in the gel. Isolates

were considered as having the same genotypic identity when presented identical AP-PCR

product-size profiles. Any repeatable difference regarding the strong bands was considered

discriminatory. The genotypes found were analyzed descriptively and their proportion, in

relation to the number of colonies isolated in each sample and condition, was calculated.

4.8 Acid Tolerance Response of S. mutans clinical isolates Acid Tolerance Response (ATR) analysis was performed in order to assess the S. mutans

isolates’ acidurance by measuring their survival at a “Killing” (pH 3.2) pH. Overnight cultures

of the S. mutans isolates were diluted (1:20) into TYE (10% tryptone, 5% yeast extract, 17.2 mM

K2HPO4) supplemented with 5 mM glucose at pH 7.5 and incubated at 37°C in a 5% CO2-

enhanced environment until mid-log phase (OD600 of ∼0.4). Cultures were then pelleted via

centrifugation for 10 min at 4000 revolutions per min. Media were discarded and cell pellets

were resuspended in TYE at the lethal pH value of 3.2. An aliquot was immediately removed

(T0) and serially diluted in 10 mM potassium phosphate buffer (pH 7.2), and the incubation of

the culture was continued for 1 h (T1) at 37°C in an atmosphere of 5% CO2. Twenty microliters

of each dilution was spotted in triplicate onto Todd Hewitt Yeast Extract (THYE) agar plates and

incubated at 37°C in a 5% CO2-supplemented atmosphere for 48 h. Adapted cells were first

resuspended in TYE at pH 5.5 for 2 h prior to being subjected to TYE at pH 3.2. Plates were

24

incubated at 37°C, 5% CO2 for 48 h and then colonies were counted. The percentage of

S. mutans surviving cells was determined by the following calculation:

4.9 Persister cell formation by S. mutans clinical isolates Overnight cultures of S. mutans clinical isolates were grown in 3mL of BHI, incubated at 37°C

in 5% CO2. A dilution (1:20) was plated onto BHI prior to the addition of the fluoroquinolone

antibiotic ofloxacin (20 µg/ml) at zero h (T0) and after 24 h (T24). In order to count colonies,

serial dilutions (by a factor of 10) were plated. These plates were incubated at 37°C in the

presence of 5% CO2. The total number of colonies were counted after 48 h of incubation. All

assays were performed in triplicate from three independent cultures. The percentage of persister

cells from each sample was determined by the following calculation:

4.10 Statistical analysis

Mean subject ages, %MS, ATR survival, and persister cell survival were compared and tested for

statistical significance using Student’s t-test. These tests were repeated with the Mann-Whitney

U Test in order to account for possible lack of data normality. Statistical significance was set at

p<0.05. Correlations between variables were measured with r2 values. Statistical analyses were

completed in Microsoft Excel Version 15.16.

% survival = CFU/mL at T1 x 100 CFU/mL at T0

% Persister Cells = CFU/mL at T24 x 100 CFU/mL at T0

25

5. Results 5.1 Subject data The S-ECC group consisted of 23 subjects (14 males, 13 females) and the CF group consisted of

21 subjects (9 males, 12 females) (Appendix A). The mean age of the subjects in the S-ECC

group was 50.8 ± 10.5 months and for the CF group, 43.4 ± 10.8 months (Table 1 and Appendix

A). Using Student’s t-test, this difference was statistically significant (p=0.027). The median

age of subjects in the S-ECC group was 55 months, whereas the median age of the CF group was

45 months (Table 1 and Appendix A). Using the Mann-Whitney U Test, this too represented a

statistically significant difference in age between the two groups (p=0.037).

Table 1. Subject age data. Subject #946 was removed from the mean age calculation due to an anomalously low %MS, which was unable to be replicated due to minimal quantity of plaque in their sample.

S-ECC CF p value

Age (months)

Mean ± Standard Deviation 50.8 ± 10.5 43.4 ± 10.8 0.027

Median 55 45 0.037

Of the S-ECC group, 40.9% were born via Caesarian section, whereas 19.0% of the CF group

were birthed in this manner (Appendix A). Using a Chi-Squared Test to compare proportions,

this difference was not statistically significant (p>0.05), however it represents a 2.2-fold increase

in the proportion of Caesarian section-delivered subjects in the S-ECC group as compared to the

CF group.

The mean dmft and dmfs scores of the S-ECC group were 10.0 ± 4.8 teeth and 25.7 ± 16.0

surfaces, respectively (Table 2 and Appendix B). The high standard deviations in the dmfs scores

reflected substantial variability in the S-ECC group with respect to the extent of caries

development among subjects. Three subjects (#194, #705 and #546) had greater than 50 carious

tooth surfaces (Appendix B), indicating that over half of the 100 possible tooth surfaces in their

26

mouths had developed dental caries. On the other end of the spectrum, subject #088 had a dmfs

score of 5 (Appendix B), demonstrating a comparatively smaller prevalence of carious teeth. To

help quantify the extent and severity of caries in each subject, the number of teeth with carious

pulpal involvement was also noted and found to be 2.8 ± 3.3 teeth (Table 2 and Appendix B). As

per the inclusion criteria, all subjects in the CF group had dmft and dmfs scores of zero.

Table 2. Subject caries data for S-ECC group.

Mean ± Standard Deviation Median dmft 10.0 ± 4.8 9

dmfs 25.7 ± 16.0 24

Number of Teeth with Carious Pulpal Involvement 2.8 ± 3.3 2

5.2 Mutans streptococci percentage determination in plaque samples There was a wide variability in the percentages of MS relative to the total bacterial cell count

from plaque samples from both groups (Table 3). Using Student’s t-test, there was no

statistically significant difference (p>0.05) in mean %MS from the total bacterial cell count

between the S-ECC (24.71 ± 17.79%) and CF (23.25% ± 16.10%) groups (Table 3, Figure 1, and

Appendix C). The median %MS in the S-ECC group was 23.91% as compared to 16.19% in the

CF group (Appendix C). While not statistically significant, this represented nearly a 1.5-fold

increase between the two groups. As age may be a confounding variable, since it is correlated

with caries experience, an analysis was performed with only the subjects aged 36-71 months

(i.e., 3 years of age and older). When subjects below age 36 months were excluded, no

statistically significant differences were observed in the mean %MS in the S-ECC (24.97 ±

18.88%) and CF groups (23.58 ± 16.52%) (Table 3, Appendix D). Among samples testing

positive for S. mutans, the mean %MS was 23.33 ± 20.65% in the S-ECC group and 41.84 ±

15.46% in the CF group (Table 3). Despite a nearly 2-fold difference in %MS, this difference

was not statistically significant (p>0.05).

27

Table 3. %MS in individual subjects (top) with mean %MS for all subjects, subjects greater than 3 years of age, and subjects with positive S. mutans identification (bottom). “*” denotes S-ECC subjects chosen for subsequent analyses. Subjects highlighted indicate those testing positive for S. mutans. Subject #946 was removed from the mean %MS calculation due to an anomalously low %MS, which was unable to be replicated due to minimal quantity of their plaque sample.

S-ECC CF Subject %MS Subject %MS

*063 14.62 ± 1.91 481 40.52 ± 4.78 *194 44.44 ± 1.63 776 15.01 ± 4.78 *485 6.70 ± 1.43 623 24.59 ± 2.43 *018 10.33 ± 1.08 443 15.63 ± 9.32 *438 10.17 ± 6.82 053 22.42 ± 5.35 *190 8.68 ± 0.67 041 45.58 ± 4.66 *200 7.63 ± 1.66 973 38.06 ± 1.21 *749 24.74 ± 2.81 780 9.81 ± 1.16 *705 37.50 ± 1.86 800 16.19 ± 1.29 *546 68.44 ± 14.16 946 0.7 832 32.22 ± 3.93 326 7.02 ± 0.22 566 31.57 ± 7.43 519 25.14 ± 0.90 417 9.45 ± 6.29 060 25.58 ± 18.77 171 45.69 ± 2.08 035 14.65 ± 6.37 900 6.27 ± 1.09 943 39.00 ± 1.84 652 23.91 ± 1.31 750 16.01 ± 3.62 088 0.17 ± 0.04 886 23.75 ± 1.63 201 12.99 ± 8.91 607 67.03 ± 0.51 198 43.73 ± 9.48 097 1.58 ± 1.05 172 12.69 ± 4.76 787 1.94 ± 0.62 881 29.67 ± 16.33 726 15.47 ± 4.90 736 40.00 ± 10.61 489 46.72 ± 10.80

Mean %MS

S-ECC CF p value (Student’s t-test)

All subjects 24.71 ± 17.79 23.25 ± 16.10 0.78

Subjects >3 years of age 24.97 ± 18.88 23.58 ± 16.52 0.82

Samples testing positive for S.mutans 23.33 ± 20.65 41.84 ± 15.46 0.10

28

Figure 1. Mean %MS with standard deviations for S-ECC and CF subjects.

5.3 Identification of S. mutans and genotyping Twenty-five MS isolates from each subject were screened for S. mutans using htrA gene-specific

primers (example of typical electrophoretic gel of PCR-amplified samples from #607, Figure 2).

S. mutans was identified in all of the first 10 S-ECC subjects tested, but only 6 of the 21 CF

subjects (Table 3, grey boxes indicate subjects where S. mutans was identified). As a result, only

samples from these 6 CF subjects were used for the remainder of the study. In order to have

similar group sizes, only the first 10 S-ECC subjects were analyzed for the remainder of the

study.

Among the CF subjects, S. mutans was more commonly isolated in subjects where the %MS was

greater than 20%, whereas in the S-ECC group, %MS did not appear to influence the ability to

isolate S. mutans.

0

5

10

15

20

25

30

35

40

45

Perc

enta

ge M

utan

s St

rept

ococ

ci (%

) S-ECC

CF

29

Figure 2. Gel showing the amplified PCR product of 479bp using the htrA S. mutans gene-specific primers of MS isolates from Subject #607 (CF group). Twenty-three of twenty-five S. mutans clones (asterisks in gel) were identified in this subject. “+” denotes the positive control (UA159)

In most subjects, only one S. mutans genotype was detected, however five subjects (#063, #194,

#749, #973, and #623) had more than one genotype isolated from their samples (Table 4). Ten

isolates containing S. mutans were considered representative of all the S. mutans strains in each

subject, in order to ensure that all strains were accounted for. In subjects where more than one

S. mutans genotype was identified, a specific genotype predominated in the majority of the

isolates and this genotype was regarded as the “dominant” genotype for the particular subject

(each distinct genotype denoted by a number following the patient’s identification). For example,

in the 8 S. mutans-containing isolates tested for subject #194, 7 of them were the same genotype

(#194-5-8 and 19-21, the dominant genotype), whereas one (#194-1) was a different genotype

(Figure 3). When applicable, both dominant and non-dominant genotypes were subsequently

used to test for the ATR and persister assays. Thirteen unique genotypes of S. mutans were

identified among the 10 S-ECC subjects (Figure 4), while 9 genotypes were identified among the

6 CF subjects (Figure 5). Two genotypes were identified from subjects #063, #194, #749, and

#973, while three genotypes were identified in the CF subject #623.

30

Table 4. Number of genotypes identified per subject.

Subject # of Genotypes 481 1 623 3

CF 973 2 946 1 943 1 607 1

Figure 3. AP-PCR gel demonstrating multiple S. mutans isolates from subject #194. All isolates were the dominant genotype, except #194-1 (Lane #1), which possessed a different “fingerprint” from the AP-PCR. The dominant genotype is circled in red, the non-dominant genotype is circled in green.

Isolate 1 2 3 4 5 6 7 8

Subject # of Genotypes 063 2 194 2 485 1 018 1

S-ECC 438 1 190 1 200 1 749 2 705 1 546 1

31

Figure 4. AP-PCR gel demonstrating the 13 different S. mutans genotypes identified from the 10 S-ECC subjects analyzed. Isolates circled in red represent different genotypes isolated from the same subject.

Figure 5. AP-PCR gel demonstrating the 9 different S. mutans genotypes identified from the 6 CF subjects analyzed. Isolates circled in red represent different genotypes isolated from the same subject.

5.4 Acid Tolerance Response of S. mutans clinical isolates Using Student’s t-test, the ATR assays for both groups showed no statistical significant

difference (p>0.05) in the mean survival percentage of S. mutans between S-ECC (21.60 ±

16.37%) and CF (17.95 ± 9.73%) groups (Table 5, Figure 6, and Appendix E). However, there

appeared to be a trend toward higher survival in the S-ECC group.

Interestingly, in some subjects where there was more than one genotype present, a statistically

significant difference in ATR was demonstrated. In the three S-ECC subjects where more than

one genotype was present, as in subjects #063, #194, and #749, the dominant genotype had

1 kb 018-3 063-4 063-22B 438-1 194-1 194-8B 485-16 190-2 705-5 749-3 749-4 546-4 200-11

The distinct genotypes, when present, as in subjects #973 and 623 were confirmed using a separate primer (not shown).

1 kb 973-1 973-2 481-11 946-12 623-1 623-4 623-9 943-A 607-1

32

significantly higher cell survival than the non-dominant genotype by a factor of 2.8x, 2.0x, and

3.5x, respectively (Figure 7, Appendix E). Higher ATR in the dominant genotype was not

observed in the two CF samples where more than one genotype was present (i.e. subjects #973

and #623). In the CF subject #973, there was no difference between the dominant (#973-2) and

non-dominant genotype (#973-1) in cell survival percentage (Figure 7, Appendix E). In the CF

subject #623, where three distinct genotypes were isolated, both of the non-dominant genotypes

(#623-4 and #623-9) had similar survival percentages to the dominant genotype (#063-1) (Figure

7 and Appendix E).

Table 5. Mean Acid Tolerance Response survival percentage from S-ECC and CF isolates.

S-ECC CF p value (Student’s t-test)

Mean Survival (%) 21.60 ± 16.37 17.95 ± 9.73 0.52

Figure 6. Mean Acid Tolerance Response with standard deviations between S-ECC versus CF

S. mutans isolates.

1

10

100

Perc

enta

ge C

ell S

urvi

val

(log

%)

S-ECC

CF

33

Figure 7. Mean Acid Tolerance Response of isolates with standard deviations (analysis was performed in triplicate) indicating survival percentage at non-adaptive pH 3.2. Isolate numbers with “D” in parenthesis indicates the dominant genotype.

5.5 Persister cell formation by S. mutans clinical isolates Using Student’s t-test, persister cell formation analysis demonstrated a statistically significant

difference (p=0.004) of a 15-fold increase in the survival percentage of the S-ECC group (1.51 ±

1.41%) as compared to the CF group (0.10 ± 0.07%) (Table 6, Figures 8 and 9, and Appendix F).

Table 6. Mean persister cell survival percentage from S-ECC and CF isolates.

S-ECC CF (%) p value (Student’s t-test)

Mean Survival (%) 1.51 ± 1.41 0.10 ± 0.07 0.004

1

10

100

018-

3…06

3-4

(D)

063-

2243

8-1

194-

8B (D

)19

4-1

485-

1619

0-2

705-

574

9-3

(D)

749-

454

6-4

200-

1197

3-2

(D)…

973-

148

1-11

946-

1262

3-1

(D)

623-

462

3-9

943-

A60

7-1

Cel

l Sur

viva

l (lo

g %

)

Isolate #

S-ECC

CF

34

Figure 8. Mean persister cell survival with standard deviation in S-ECC and CF isolates.

Figure 9. Mean persister cell survival percentages with standard deviations (analysis was performed in triplicate) of all isolates. Isolate numbers with “D” in parenthesis indicates the dominant genotype.

0.00

0.01

0.10

1.00

10.00

Perc

enta

ge S

urvi

val (

%)

S-ECC

CF

0.01

0.10

1.00

10.00

018-

3…06

3-4

(D)

063-

2243

8-1

194-

8B (D

)19

4-1

485-

1619

0-2

705-

574

9-3

(D)

749-

454

6-4

200-

1197

3-2 …

973-

148

1-11

946-

1262

3-1

(D)

623-

462

3-9

943-

A60

7-1

Pers

ister

Cel

l Sur

viva

l (lo

g %

)

Isolate #

S-ECCCF

35

5.6 Correlations between measures of caries severity, mutans

streptococci percentage, acid tolerance, and persister cell survival Weak correlations were found between any measures of caries severity (dmfs, dmft, or number

of teeth with carious pulpal involvement) with mutans streptococci percentage, acid tolerance or

persister cell survival (Appendix G). There was also a weak correlation between acid tolerance

and persister cell survival in S-ECC or CF samples (Appendix H).

5.7 Summary of Results The CF group was on average 7 months younger than the S-ECC group, however this difference

was not statistically significant. There was no difference in the %MS in the plaque samples

between the groups. However, S. mutans was isolated in all subjects screened in the S-ECC

group, regardless of %MS, whereas in the CF group, S. mutans was isolated more frequently

when %MS was relatively high (greater than 20%). In most subjects, only one S. mutans

genotype was isolated; however, two genotypes were isolated from the plaque of four subjects

and three genotypes were isolated from the plaque of one subject. Although there was no

difference overall in the ATR between the S-ECC and CF groups, the dominant genotype in S-

ECC subjects with more than one genotype showed greater cell survival after exposure to the

lethal pH. Increased acid tolerance was not observed in the dominant genotypes of CF subjects

where more than one genotype was present.

A statistically significant, 15-fold increase was found in the persister cell survival in the S-ECC

group as compared to the CF group. This novel finding demonstrates a markedly enhanced

persistence phenotype present in the S. mutans strains from the S-ECC group.

36

6. Discussion The present study was undertaken to investigate genotypic and phenotypic differences in

S. mutans samples collected from the dental plaque of young children with and without dental

caries. While previous studies have examined some of these areas, our research is novel in that it

was the first to investigate the differences in S. mutans persister cell formation and ATR from

plaque sampled directly from human subjects. This study helps to provide insight into the

relationship between MS, S. mutans, and dental caries in children. The results demonstrate the

genotypic and phenotypic variation among S. mutans strains both between and within

individuals. Some of these differences may account for the clinical presence or absence of caries

lesions and its clinical severity.

6.1 Sample population data Age has been shown to correlate with dental caries experience (Declerck et al., 2008; Parisotto et

al., 2010; Dye et al., 2015). This association has the potential to affect the %MS data because

although some of the younger CF subjects may not have manifested clinical caries yet, with more

time there is potential for caries lesions to develop in the future. The S-ECC group was on

average 7 months older than the CF group, which was statistically significant. Even when

subjects below 36 months of age were excluded in an attempt to control for subject age, there

was still no statistical difference in the mean %MS. Nevertheless, subject age may be considered

a confounding variable, as the longer a tooth is present in the oral cavity, the greater the

S. mutans colonization, which subsequently increases the risk of manifesting clinical caries

lesions as compared to a newly erupted tooth (Wan et al., 2003; Wen et al., 2012; Nelson et al.,

2014). As some of the subjects, particularly in the CF group, were under age 3 years old, some

of their teeth would have just recently erupted into the oral cavity, with less potential time to

develop dental caries lesions (Smith, 1991).

The variability in the severity of caries between subjects in the S-ECC group suggests that future

studies of this nature should take into consideration the fact that the current classification of

S-ECC might not be discriminative enough, or may need to be revised or sub-categorized, to

differentiate between relatively mild S-ECC and the most severe cases. While all members of the

37

S-ECC group in the current study fell into the AAPD’s classification of S-ECC, some individuals

had far more severe presentations than others with respect to both dmfs scores and the number of

teeth with carious pulpal involvement. For example, subject #063 had 15 carious teeth, of which

13 had pulpal involvement, indicating a very severe progression of caries, such that nearly all of

that subject’s teeth had carious lesions that extended to the dental pulp (Appendix B). However,

several other subjects had zero or only one tooth with carious pulpal involvement, despite having

numerous teeth with more minor carious lesions. The high standard deviation in the number of

teeth with carious pulpal involvement, which was higher than the mean number, demonstrated

the variability in severity of caries progression among the subjects in the study. Regardless, dmft,

dmfs, and number of teeth with carious pulpal involvement did not correlate with %MS, ATR, or

persister cell survival (Appendix G). This suggests that these measures of caries severity may

not be directly associated with presence of MS or the acidurance and persister phenotypes of

S. mutans. As dental caries is a multi-factorial disease process involving numerous other factors

such as diet, oral hygiene habits, and salivary buffering capacity, among others (Featherstone,

2006; Mitchell et al., 2009), the interplay between these variables is complex and not solely

linked to one alone.

In the present study there was a lack of statistical significant difference between the two groups

of participants born via Caesarian section with respect to presence of dental caries. There was,

however, a 2.6-fold increase in the percentage of Caesarian section-delivered subjects in our

sample that presented with S-ECC. Some studies have correlated an increased susceptibility to

dental caries and earlier acquisition of S. mutans to those born via Caesarian section (Li et al.,

2005; Pattanaporn et al., 2013). The rationale is that infants delivered by Caesarean section are

more aseptic and the atypical microbial environment increases the chances of S. mutans

colonization (Li et al., 2005). Other studies, however, have found that mode of delivery does not

correlate with the early colonization of S. mutans in the oral cavity of infants (Thakur et al.,

2012; Ubeja and Bhat, 2016; Brandquist et al., 2017). Lif Holgerson et al. (2011) found

differences in the oral microbiota in infants related to mode of delivery, with vaginally delivered

infants having a higher number of taxa detected by the Human Oral Microbe Identification

Microarray (HOMIM) microarray. Nelun Barford et al (2012) found that in 3-year-old Danish

pre-school children, caries prevalence was not related to the mode of delivery; however, there

38

was a tendency for more severe caries activity in the C-section group. Despite the lack of

statistical significance, our data showed a trend toward Caesarian section as a risk factor for

ECC.

6.2 Mutans streptococci percentage The %MS within the plaque samples from the S-ECC and CF subjects in the current study was

not statistically different, calling into question the value of using this measurement to predict

caries status, as has been done in the past (Hong et al., 2010; Seki et al., 2003; O’Sullivan and

Thibodeau, 1996). Multiple studies, however, have found an association between MS

concentrations with ECC (Fan et al., 2016; Seki et al., 2003; Vachirarojpisan et al., 2004;

Ghasempour et al., 2013; Ma et al., 2015), including a systematic review by Parisotto et al.

(2010), which found that the salivary count of MS was a strong risk indicator for ECC. Valdez

et al. (2017) found that counts of MS in biofilms of ECC and S-ECC children did not differ from

each other, but %MS were both higher than those found in CF children.

The absence of correlations between dmft, dmfs, and pulpally involved carious lesions to %MS

in the S-ECC group (R2 = 0.20, 0.20, and 0.08, respectively) agree with other studies that have

demonstrated a lack of association between %MS and manifest caries lesions. No difference was

found in %MS from total streptococci population in CF versus CA Venezuelan children

(Acevado et al., 2009) and high levels of MS were detected in a Sudanese population with very

low caries prevalence (Carlsson et al., 1987). A systematic review of the accuracy of different

caries risk assessment methods found that mutans streptococci sampled from plaque and saliva

demonstrate low sensitivity and high specificity (Senneby et al., 2015). This indicates that high

%MS from plaque or saliva may be useful to rule in disease (caries), but low %MS cannot be

used to rule out disease. They concluded that this finding indicates the importance of other non-

MS acidogenic species in caries pathogenesis. This appeared to be in accordance with our data,

where many of the S-ECC subjects had high %MS, but other S-ECC subjects, such as #088,

#190, #900, #485, #200, and #417, had %MS below 10% (Table 3), but all had dmfs scores

greater than or equal to 15 (Appendix B). Our data did, however, include multiple subjects in the

CF group where %MS was relatively high, such as subjects #481, #041, #607, #973, and #943,

39

who all had %MS greater than 35%. Overall, we feel this indicates a relatively poor capacity for

%MS to predict the clinical presence of dental caries lesions.

A possible explanation as to why %MS as a predictor of caries status could be flawed is that

some of the MS species are not acidogenic, and therefore are not cariogenic, e.g., a

spontaneously occurring lactate dehydrogenase deficient mutant of Streptococcus rattus has been

shown to be non-acidogenic and suggested as a probiotic (Hillman et al., 2009). Furthermore,

genotypic diversity exists among S. mutans and S. sobrinus strains with respect to adhesion to

tooth surfaces, acidogenicity, and acidurance, resulting in differences in cariogenicity (Banas,

2004). Therefore, two individuals with similar %MS may have differing caries experience

dependent on the specific strains of these bacteria that are present. Lembo et al. (2007) found

differences in susceptibility to acid challenge between S. mutans strains, with greater acidurance

statistically more frequent among isolates from CA children than among those from CF children.

Valdez et al. (2017) found that although genotypic diversity and acidogenicity of S. mutans were

similar among CF, ECC, and S-ECC children, strains isolated from the CA groups formed more

biofilms than those from the CF group. The strains they isolated from the S-ECC group were

also highly aciduric (Valdez et al., 2017).

6.3 Identification of S. mutans Despite statistically similar proportions of MS in the plaque samples collected from both groups,

S. mutans were identified far more readily from the S-ECC group as compared to the CF group

in the current study. In a similar study, plaque from S-ECC children presented the highest

percentage of S. mutans isolates as compared to plaque from CF and ECC children, which

showed no difference between them (Valdez et al., 2017). These results were also in agreement

with Kouidhi et al. (2014), who found that S. mutans detection in biofilms increased with

increasing severity of dental caries activity. Multiple studies have evaluated the differences in

plaque and salivary microbial profiles between CA and CF children (Ma et al., 2015; Parisotto et

al., 2010; Tanner et al., 2011; Jiang et al., 2016; Johansson et al., 2016). Using the HOMIM, Ma

et al. (2015) detected a significantly higher prevalence of S. mutans in both the plaque and saliva

of S-ECC children as compared to CF children. Tanner et al. (2011) isolated bacteria from

plaque samples from 42 children with S-ECC and 40 CF children and identified the presence of

40

various bacterial species by comparison of 16S rRNA taxon sequences with the Human Oral

Microbiome Database. They concluded that the major caries-associated species were S. mutans

and S. wiggsiae. In our study, S. mutans was most commonly found in the CF samples where

MS made up greater than 20% of the proportion of colonies, whereas %MS did not seem to

influence the ability to isolate S. mutans in the S-ECC group. Speculatively, this may have been

due to a qualitative observation of a generally larger bulk of plaque that was available for

collection in the S-ECC group. As a result, the S-ECC group’s plaque may have been more

mature, and thereby more likely to have S. mutans, a middle colonizer, whereas the CF group’s

plaque may have been made up of more early colonizers (Li et al., 2004).

6.4 Genotyping S. mutans Numerous studies have established the diversity of distinct genotypes of S. mutans colonizing the

oral cavity (Klein et al., 2004; Lembo et al., 2007; Mattos-Graner et al., 2004; Napimoga et al.,

2004; Palmer et al., 2013). Some authors have reported that CA individuals seem to harbour

more S. mutans genotypes than CF individuals, with the simultaneous action of multiple strains

possessing different virulence factors increasing the risk for caries development (Tabchoury et

al., 2008; Napimoga et al., 2004; Alaluusua et al., 1996). We were able to identify 13 distinct

genotypes among the 10 S-ECC subjects and 9 distinct genotypes among the CF subjects: 2

genotypes in 3 of the S-ECC subjects; 2 and 3 genotypes in 1 CF subject each. The relatively

small degree of genotypic diversity in our study is in accordance with previous studies of saliva

and plaque in children (Klein et al., 2004; Lembo et al., 2007; Mattos-Graner et al., 2004;

Napimoga et al., 2004; Tabchoury et al., 2008). As in our study, Valdez et al. (2017) used

AP-PCR with the OPA-02 primer to identify distinct S. mutans genotypes. Interestingly, they

found that most subjects in their study (53.6%) had two S. mutans genotypes, whereas the

majority of subjects in our study had only one genotype. Although subject age was similar in

their study to ours, perhaps ethnicity or variations in the environment may play a role in the

variety of genotypes present, as their study was performed in Brazil. The dominant strain’s

ability to out-compete the other S. mutans genotypes reflects an evolutionary advantage (Cheon

et al., 2013). This advantage may be characterized by an increased phenotypic capacity of the

dominant strain to survive the conditions present in the oral cavity, such as greater acidurance or

biofilm formation (Valdez et al., 2017; Lembo et al., 2007). It may also reflect a greater efficacy

41

or potential for mutacin production by these strains, capable of killing other MS bacteria,

including other S. mutans strains (Hillman et al., 1987). Hillman et al. (2000) suggested

exploiting this anti-S. mutans mutacin production by replacement therapy, wherein a non-

cariogenic (lactate-dehydrogenase deficient) strain of S. mutans with increased mutacin

production capacity would be introduced to eliminate the cariogenic strains of S. mutans.

6.5 Acid Tolerance Response of S. mutans isolates Acidurance, the ability to endure low pH conditions, is an important virulence factor for

S. mutans to survive in the oral cavity, and substantial variation in this phenotype exists amongst

different strains (Palmer et al., 2013). S. mutans’ acid-adaptive mechanisms are multi-factorial,

and involve modifications of the plasma membrane and extracellular matrix, management of

sugar import and output of fermentation end-products, extrusion of protons, influx of other

cations, generation of intracellular basic molecular species, and repair of acid-damaged protein

and DNA (Baker et al., 2017). Genotypic variation related to any of these factors can influence

the acidurance of any particular strain. Our results found substantial variation in the ATR among

the various S. mutans strains; however, there was no statistically significant difference in the

survival percentage of S-ECC versus CF isolates after being subjected to the non-adaptive

pH 3.2. There was, however, a trend that showed increased survival of the S-ECC isolates as

compared to the CF isolates (S-ECC = 21.60 ± 16.37%; CF = 17.95 ± 9.73%). The standard

deviations in these data were quite high, reflecting the large variability in the ATR (acidurance)

of the S. mutans isolates. Interestingly, the ATR data in our study showed considerable variation

within subjects where more than one S. mutans genotype was present. In the S-ECC samples

where two genotypes were present, the dominant genotype had significantly higher cell survival

than the non-dominant genotype, whereas in the CF samples with more than one genotype, the

dominant genotype had statistically similar cell survival than the non-dominant genotype(s). It

would appear that these dominant genotypes were better suited to survive the acidic conditions

present in the oral cavities of the S-ECC subjects, and their genotype was selected more

favourably (Marsh, 1994; Welin-Neilands and Svensäter, 2007).

In contrast to our study, other studies showed statistically significant differences in acid tolerance

between CA and CF samples. Valdez et al. (2017) found a statistically significant difference in

42

ATR, with their S-ECC S. mutans isolates demonstrating higher percentage of bacterial growth

at pH 2.8 as compared to the CF isolates. Perhaps the difference in findings could be accounted

for by the difference in pH used as the “Killing/Lethal pH” in their study (2.8) versus ours (3.2),

or by differences in the sample population. Lembo et al. (2007) also found that genotypes with

low susceptibility to acid challenge (i.e., high acidurance) were more common among CA than

CF isolates. Jiang et al. (2017) found that an S. mutans strain isolated clinically from a CA adult

exhibited greater acidurance compared to strains isolated from a CF adult and a reference strain,

indicating that the colonization of highly aciduric, and therefore cariogenic, clinical strains may

lead to high caries risk in individuals. A possible explanation for the contrasting data of our

study as compared to Jiang et al. (2017) is that they only analyzed S. mutans strains isolated from

one CA subject, one CF subject, and a reference strain, whereas our study evaluated multiple

strains of S. mutans isolated from multiple subjects in each group. The variability of ATR

demonstrated by strains within both group we studied and even between different genotypes

from the same subject, reflects the necessity to evaluate multiple strains of S. mutans to

determine the differences in ATR between CA and CF individuals. In agreement with our

findings, were the results of de Camargo et al. (2018), who found no difference in the acid

tolerance of S. mutans salivary samples from CA and CF children using a similar experimental

protocol. While our data did not show a statistically significant increase in ATR in the S-ECC as

compared to the CF group, this trend was observed, which is in accordance with the previous

studies of Valdez et al. (2017) and Jiang et al. (2017).

6.6 Persister cell formation of S. mutans isolates The most striking finding in our data was the 15-fold increase in the S. mutans persister cell

survival percentage in the S-ECC compared to the CF groups, a finding that supported our

hypothesis. This novel finding suggests that the environmental stressors in the oral cavities of

children with S-ECC selects for the propagation of S. mutans strains with greater expression of

the persistence phenotype. This study was also novel in that, to our knowledge, this is the first

study to demonstrate the formation of persister cells from bacteria harvested in vivo, as opposed

to stock laboratory strains.

43

The ability of S. mutans to utilize the persistence phenomenon can help explain the recurrent

nature of dental caries. Previous studies investigating persister cell formation in other bacterial

species, such as Escherichia coli and S. aureus, have reported the proportion of persister cells

among the total bacterial count to be in the range of 0.1% to 1% (Keren et al., 2004). Similar

proportions have also been found in persister cell formation in S. mutans in studies where

ofloxacin (Leung and Levesque, 2012; Leung et al., 2015) and dimethylaminohexadecyl

methacrylate or chlorhexidine (Jiang et al., 2017; Wang et al., 2017) were used as antibiotic

agents. Therefore, the higher persister cell survival in the S-ECC group is of interest, as these

particular strains have demonstrated an enhanced persistence phenotype beyond that which is

normally reported. Persistence is not limited to bacteria, having been demonstrated in

Candida albicans, a fungus commonly found in the oral cavity (Lafleur et al., 2006; Lafleur et al.

2010). Lafleur et al. (2010) investigated the levels of persister cells of C. albicans between

cancer patients with differing time periods of oral carriage of the fungus. Interestingly, they

found that persister cell levels were significantly higher in individuals with longer-term carriage

of C. albicans. This may help explain our findings wherein the S-ECC group, who were on

average older and likely had S. mutans carriage for longer periods of time than the CF group,

demonstrated significantly higher persister cell counts.

An important factor in the persistence phenomenon is that it appears to be increased in cells

found in biofilms (Lewis, 2005). The importance of the biofilm in persister cell formation is in

the intercellular communication via quorum sensing, as this allows the bacteria to determine

which cells are to become persisters (Leung et al., 2015; Leung and Levesque 2012; Reuter et al.

2016). However, planktonic bacterial cells, including S. mutans, have been shown to be capable

of forming persisters (Jiang et al., 2017; Leung and Lévesque, 2012; Lewis, 2012). Another key

feature of persister cell formation is the involvement of toxin-antitoxin systems, composed of a

toxin (protein) and an antitoxin (protein or non-coding RNA) (Hayes and Van Melderen, 2011;

Page and Peti, 2016). The toxin arrests cellular growth by interfering with a vital cellular

process, whereas the antitoxin neutralizes the toxin activity during normal growth conditions

(Hayes and Van Melderen, 2011). Under stressful conditions the antitoxins are selectively

degraded, leaving the toxins to exert their effects, leading to growth arrest and dormancy (i.e.,

persister formation) (Christensen et al., 2004; Schuster et al., 2013). Inhibition of quorum

44

sensing and toxin-antitoxin pathways in S. mutans may be a useful target to eliminate persister

cell formation, thereby reducing infection relapse (Leung and Levesque, 2012; Page and Peti,

2016).

Targeting the metabolic pathways of S. mutans appears to be a promising strategy to combat

persister cells, as their lack of metabolism, evidenced by reduced adenosine triphosphate (ATP)

levels, is what renders them tolerant to antibacterial agents (Wood, 2017; Kwan et al., 2013;

Conlon et al., 2016). Jiang et al. (2017) induced S. mutans colonies to form persister cells

exposing them to a variety of different antibiotics and to an antibacterial quaternary ammonium

compound (dimethylaminododecyl methacrylate). They then monitored the expression of

several glycolysis- and citrate cycle-related genes, including adhE, ldh, and pdhA/B/C/D, to

evaluate the carbon source metabolism of the S. mutans persisters. As they had expected, the

metabolically dormant persister cells downregulated the expression of these genes as compared

to the control group’s normal cells. They hypothesized that the activation of carbon source

metabolism may be a possible strategy for the eradication of the otherwise dormant S. mutans

persisters. Addition of extra glucose to the culture significantly reduced levels of persisters in

both planktonic and biofilm conditions, which they hypothesized was due to changing of

dormant persister cells into “active” cells. Waking of persister cells by the addition sugars or

glycolysis intermediates and then killing them by bactericidal has also been shown to be

effective with other bacterial species (Allison et al., 2011; Marques et al., 2014); however, more

research will be required to determine how to best deliver this type of therapy to patients.

45

7. Conclusions and Future Studies Genotypic and phenotypic diversity exists among S. mutans derived from the plaque of children

with and without S-ECC. Possible important phenotypic traits that may play a role in increasing

virulence of S. mutans include increased tolerance to acid (acidurance) and increased production

of persister cells capable of surviving environmental stressors such as antibiotics. Genotypic

diversity in terms of the presence of dominating genotypes that are more acid tolerant may help

explain the increased ability of these strains to cause dental caries. The ability of S. mutans to

utilize persistence in the face of high levels of antibiotics is one of the reasons for the chronicity

and recurrence of dental caries activity. The novel finding of the enhanced persister cell

formation found in the plaque of children with S-ECC further emphasizes the importance of

targeting microbial persistence in the fight against dental caries, particularly in high-risk

populations.

Future studies should aim at further understanding the underlying mechanisms involved in

establishing chronicity of infection via persister cell formation. This will allow for the

development of pharmacological agents that can target the mechanisms by which S. mutans is

able to cause infection relapse. Further research will be required to determine the optimal

vehicle for delivery of these agents, perhaps topically or via dental restorative materials.

46

8. Appendices

Appendix A. Subject data for S-ECC (n=23) and CF (n=21) subjects.

S-ECC

Subject Age Age (months) Gender Pre-Term C-Section

018 4y10m 58 Male No Yes

190 3y4m 40 Female Yes Yes

485 3y11m 47 Female No No

438 4y7m 55 Male Yes Yes

194 4y7m 55 Male No No

063 5y5m 65 Male No Yes

705 4y11m 59 Male No No

749 2y9m 33 Male Yes Yes

546 3y8m 44 Female No Yes

200 4y11m 59 Male No No

171 5y6m 66 Female No No

417 4y9m 57 Female No Unknown

881 3y6m 42 Female No No

652 4y2m 50 Female No Yes

900 3y9m 45 Male No Yes

201 3y4m 40 Male No Yes

489 4y11m 59 Female No No

736 4y8m 56 Female No No

088 5y1m 61 Female No No

172 2y9m 33 Male No No

566 2y7m 31 Female No No

832 4y5m 53 Male No No 198 5y0m 60 Male No No

Mean 50.8 SD 10.5

Median 55

47

CF

Subject Age Age (months) Gender Pre-Term C-Section

623 4y0m 48 Male No No

481 4y4m 52 Female No No

946 4y4m 52 Female No No

973 2y7m 31 Male No No

943 2y4m 28 Male No No

607 3y7m 43 Male No No

519 3y3m 39 Male No No

041 3y9m 45 Female No No

780 2y5m 29 Male No No

326 3y6m 42 Female No No

800 5y1m 61 Male No Yes

776 5y2m 62 Male No No

443 3y10m 46 Female No No

053 2y0m 24 Female No Yes

060 3y9m 45 Female Yes No

035 4y5m 53 Female No No

750 4y9m 57 Female No No

886 3y11m 47 Male No Yes

097 3y9m 45 Female No Yes

787 2y8m 32 Male No No

726 3y2m 38 Male No No

Mean 43.8 SD 10.7

Median 45

48

Appendix B: Caries data for S-ECC subjects (n=23).

Subject dmft dmfs Pulpal Involvement

018 7 19 0

190 7 15 2

485 8 24 2

438 5 16 2

194 18 52 6

063 15 42 13

705 17 61 7

749 8 11 0

546 16 55 6

200 10 27 2

171 8 16 5

417 11 31 2

881 17 31 3

652 3 8 0

900 10 22 1

201 12 35 2

489 8 17 0

736 18 41 2

088 5 5 0

172 4 8 1

566 4 4 0

832 11 26 8 198 9 24 1

Mean 10.0 25.7 2.8

SD 4.8 16.0 3.3

Median 9 24 2

49

Appendix C. Percentage mutans streptococci for S-ECC (n=23) and CF (n=20) subjects. Subject #946 was removed from the mean %MS calculation due to an anomalously low %MS (0.7%), which was unable to be replicated due to minimal quantity of their plaque sample.

S-ECC CF

Subject %MS Subject %MS

063 14.62 ± 1.91 481 40.52 ± 4.78

194 44.44 ± 1.63 776 15.01 ± 4.78

485 6.70 ± 1.43 623 24.59 ± 2.43

018 10.33 ± 1.08 443 15.63 ± 9.32

438 10.17 ± 6.82 053 22.42 ± 5.35

190 8.68 ± 0.67 041 45.58 ± 4.66

200 7.63 ± 1.66 973 38.06 ± 1.21

749 24.74 ± 2.81 780 9.81 ± 1.16

705 37.50 ± 1.86 800 16.19 ± 1.29

546 68.44 ± 14.16 326 7.02 ± 0.22

832 32.22 ± 3.93 519 25.14 ± 0.90

566 31.57 ± 7.43 060 25.58 ± 18.77

417 9.45 ± 6.29 035 14.65 ± 6.37

171 45.69 ± 2.08 943 39.00 ± 1.84

900 6.27 ± 1.09 750 16.01 ± 3.62

652 23.91 ± 1.31 886 23.75 ± 1.63

088 0.17 ± 0.04 607 67.03 ± 0.51

201 12.99 ± 8.91 097 1.58 ± 1.05

198 43.73 ± 9.48 787 1.94 ± 0.62

172 12.69 ± 4.76 726 15.47 ± 4.90

881 29.67 ± 16.33

736 40.00 ± 10.61

489 46.72 ± 10.80

Mean 24.71 Mean 22.18

SD 17.79

SD 16.44

Median 23.91 Median 16.19

50

Appendix D. Percentage MS excluding subjects below age 36 months.

S-ECC (n=20) CF (n=16)

Subject %MS Subject %MS

063 14.62 ± 1.91 481 40.52 ± 4.78

194 44.44 ± 1.63 776 15.01 ± 4.78

485 6.70 ± 1.43 623 24.59 ± 2.43

018 10.33 ± 1.08 443 15.63 ± 9.32

438 10.17 ± 6.82 041 45.58 ± 4.66

190 8.68 ± 0.67 800 16.19 ± 1.29

200 7.63 ± 1.66 326 7.02 ± 0.22

705 37.50 ± 1.86 519 25.14 ± 0.90

546 68.44 ± 14.16 060 25.58 ± 18.77

832 32.22 ± 3.93 035 14.65 ± 6.37

417 9.45 ± 6.29 750 16.01 ± 3.62

171 45.69 ± 2.08 886 23.75 ± 1.63

900 6.27 ± 1.09 607 67.03 ± 0.51

652 23.91 ± 1.31 097 1.58 ± 1.05

088 0.17 ± 0.04 726 15.47 ± 4.90

201 12.99 ± 8.91

198 43.73 ± 9.48

881 29.67 ± 16.33

736 40.00 ± 10.61

489 46.72 ± 10.80

Mean 24.97 Mean 22.23

SD 18.88 SD 16.85

Median 19.27 Median 16.19

51

Appendix E. Acid Tolerance Response survival data for S-ECC and CF isolates. Isolates with “D” in parenthesis refer to the dominant genotype when more than one genotype was present.

Status Isolate % Survival SD

S-ECC

018-3 30.99 4.46

063-4 (D) 70.22 9.37

063-22 24.88 0.49

438-1 13.32 0.89

194-8B (D) 18.94 3.5

194-1 9.31 0.53

485-16 20.48 2.49

190-2 12.41 2.87

705-5 7.36 0.77

749-3 (D) 21.99 2.05

749-4 6.34 0.52

546-4 23.66 2.98

200-11 20.85 0.91 Mean 21.60

SD 16.37

Median 20.48

CF

973-2 (D) 25.62 2.25

973-1 27.58 4.45

481-11 9.86 1.56

946-12 35.88 13.66

623-1 (D) 6.98 0.58

623-4 10.33 0.18

623-9 12.84 5.64

943-A 18.23 1.19

607-1 14.24 1.78 Mean 17.95

SD 9.73

Median 14.24

52

Appendix F. Persister cell survival percentage for S-ECC and CF isolates. Isolates with “D” in parenthesis refer to the dominant genotype when more than one genotype was present.

Status Isolate % Survival SD

S-ECC

018-3 0.51 0.01

063-4 (D) 0.07 0.02

063-22 0.19 0.02

438-1 2.31 0.52

194-8B (D) 1.37 0.04

194-1 5 1.02

485-16 1.56 0.16

190-2 3.13 1.64

705-5 2.41 1.12

749-3 (D) 0.2 0.13

749-4 0.97 0.28

546-4 0.6 0.71

200-11 1.35 0.3

Mean 1.51

SD 1.41

Median 1.35

CF

973-2 (D) 0.1 0.01

973-1 0.2 0.05

481-11 0.17 0.01

946-12 0.02 0.01

623-1 (D) 0.11 0.03

623-4 0.05 0.02

623-9 0.04 0.01

943-A 0.2 0.01

607-1 0.023 0.006

Mean 0.10

SD 0.07

Median 0.10

53

Appendix G. i. Correlations between measures of caries severity (dmft, dmfs, and number of teeth with carious pulpal involvement) with mutans streptococci percentage.

R² = 0.20273

0

10

20

30

40

50

60

70

80

0 5 10 15 20

%M

S

dmft

R² = 0.20065

0

10

20

30

40

50

60

70

80

0 20 40 60 80

%M

Sdmfs

R² = 0.07597

0

10

20

30

40

50

60

70

80

0 5 10 15

%M

S

# Teeth with Pulpal Involvement

54

ii. Correlations between measures of caries severity (dmft, dmfs, and number of teeth with carious pulpal involvement) with ATR survival percentage.

R² = 0.01743

0

10

20

30

40

50

60

70

80

0 5 10 15 20

ATR

Sur

viva

l (%

)

dmft

R² = 0.00921

0

10

20

30

40

50

60

70

80

0 20 40 60 80AT

R S

urvi

val

(%)

dmfs

R² = 0.26961

0

10

20

30

40

50

60

70

80

0 5 10 15

ATR

Sur

viva

l (%

)

# Teeth with Pulpal Involvement

55

iii. Correlations between measures of caries severity (dmft, dmfs, and number of teeth with carious pulpal involvement) with persister cell survival percentage.

R² = 0.01971

0.00

1.00

2.00

3.00

4.00

5.00

6.00

0 5 10 15 20

Pers

iste

r Sur

viva

l (%

)

dmft

R² = 0.03288

0.00

1.00

2.00

3.00

4.00

5.00

6.00

0 20 40 60 80Pe

rsis

ter S

urvi

val (

%)

dmfs

R² = 0.01893

0

1

2

3

4

5

6

0 5 10 15

Pers

iste

r Sur

viva

l (%

)

# Teeth with Pulpal Involvement

56

Appendix H. Correlation between acid tolerance and persister cell survival in both groups. S-ECC

CF

R² = 0.31052

0

1

2

3

4

5

6

0 20 40 60 80

Pers

iste

r Cel

l Sur

viva

l (%

)

Acid Tolerance Response Survival (%)

R² = 0.00065

0

0.05

0.1

0.15

0.2

0.25

0 10 20 30 40

Pers

iste

r Cel

l Sur

viva

l (%

)

Acid Tolerance Response Survival (%)

57

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