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Understanding the Regulation and Biological Function of the ComRS Competence Signaling System in Streptococcus mutans

Iwona Bernadetta Wenderska

A thesis submitted in conformity with the requirements for the degree of Doctorate of Philosophy Faculty of Dentistry University of Toronto

Understanding the Regulation and Biological Function of the ComRS Competence Signaling system in Streptococcus mutans

Iwona Wenderska

Doctorate of Philosophy

Faculty of Dentistry University of Toronto

2016 Abstract

Genetic competence provides bacteria with an opportunity to increase genetic diversity by allowing the acquisition of novel traits that can confer virulence or a survival advantage. The oral biofilm, containing a diverse microbiota at close proximity, is an ideal environment for this form of genetic exchange. Several oral streptococci have been demonstrated to engage in DNA uptake from their environment by the process of genetic transformation. In the cariogenic organism, Streptococcus mutans, competence for transformation is regulated by the ComRS peptide signaling system, comprised of the ComR transcriptional regulator and the ComS pre-peptide required to produce the mature ComX-inducting peptide, XIP. XIP in conjunction with ComR activates the ComX alternate sigma factor critical for DNA uptake and recombination. In this study, we examined the molecular pathways and biological functions of the ComRS regulon in S. mutans. We demonstrated a novel role for ComRS in competence-associated self-lysis.

Further analysis of the global transcriptome in response to XIP revealed a role for XIP in the regulation of bacteriocin genes, encoding antimicrobial peptides used by bacteria to lyse closely related species. The coordination of competence development and cell lysis may ensure availability of eDNA for DNA uptake by the competent subpopulation. Aside

ii from cell lysis, supplementation with XIP regulated a number of virulence-associated genes, and resulted in a global transcription profile that mimicked that of the starvation- induced stringent response in S. mutans. Others have shown that genetic competence and the stringent response were regulated by the rcrRPQ operon, encoding an RcrR transcriptional regulator and the RcrPQ ABC transporters. Our work further implicated

RcrRPQ in the regulation of comS expression and endogenous XIP levels. Together, our work suggests that the ComRS regulon in S. mutans exhibits a biological role beyond competence induction that includes virulence and stress adaptation.

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Acknowledgments

I would like to extend my gratitude to my supervisors, Dr. Dennis Cvitkovitch and Dr. Dilani Senadheera, for their mentorship and guidance throughout my studies. I appreciate all of the opportunities, challenges and invaluable advice. I would also like to thank my committee members, Dr. William Navarre and Dr. Tara Moriarty, for their insightful suggestions and critiques.

Thank you to all the members of the Cvitkovitch and Senadheera labs for their friendship and support. A special thank you to Gursonika and Andrew. I would also like to extend my deepest gratitude to Dr. Milos Legner for great scientific discussions, guidance and friendship.

I am forever grateful for the unwavering support and love of my parents and siblings. I am blessed to have you in my life.

Thank you.

Table of Contents

Acknowledgments ...... iv

Table of Contents ...... v

List of Figures and Tables ...... ix

List of Abbreviations ...... xi

Publications ...... xiii

Awards ...... xiv

Chapter 1: Literature Review...... 1

1.1 The oral environment ...... 2

1.1.1 The oral cavity ...... 2

1.1.2 Dental plaque and microbial succession ...... 3

1.1.3 Dental caries: microbial community in health and disease ...... 7

1.2 Streptococcus mutans ...... 8

1.2.1 Attachment to the tooth ...... 10

1.2.2 Sugar metabolism and acidogenicity ...... 11

1.2.3 Aciduricity ...... 14

1.2.4 Two-component signal transduction systems ...... 15

1.2.5 Bacteriocins ...... 17

1.2.5.1 Regulation of bacteriocin production ...... 18

1.2.6 Genetic transformation ...... 20

1.2.6.1 Regulation of genetic competence: the core competence regulators ...... 24

1.2.6.1.1 The core competence regulators and competence- associated cell lysis ...... 27

1.2.6.2 Regulation of genetic competence: TCSTSs ...... 27

1.2.6.3 Regulation of genetic competence: The RcrRPQ operon ...... 28

Chapter 2: Research rationale and hypothesis ...... 31

2.1 Research Rationale ...... 32

2.2 Hypothesis...... 32

Chapter 3: A novel function for the competence inducting peptide, XIP, as a cell death effector of Streptococcus mutans ...... 34

3.1 Abstract ...... 35

3.2 Introduction ...... 36

3.3 Materials and methods ...... 37

3.3.1 Bacterial strains and growth conditions ...... 37

3.3.2 Transformation frequency (TF) assays ...... 38

3.3.3 Cell viability assays ...... 38

3.3.4 Time-course killing analyses ...... 39

3.3.5 Biofilm formation assays ...... 39

3.3.6 Quantitative real-time PCR (qRT-PCR) analyses ...... 39

3.3.7 XIP detection and quantification ...... 40

3.4 Results ...... 41

3.4.1 Transformation frequencies (TF) with CSP and/or XIP in CDM and complex media ...... 41

3.4.2 Measurement of XIP in the culture supernatants of S. mutans UA159, comR/S, comE and comX knockout mutants ...... 43

3.4.3 Effect of XIP on cell viability of S. mutans UA159 grown in CDM ...... 45

3.4.4 Regulation of comR/S, and comX expression by XIP, CSP and the ComDE system ...... 47

3.5 Discussion ...... 51

3.6 Acknowledgements ...... 54

Chapter 4: The effects of the RcrRPQ operon on comS expression and XIP levels . 55 vi

4.1 Abstract ...... 56

4.2 Introduction ...... 57

4.3 Materials and methods ...... 60

4.3.1 Strains and growth conditions ...... 60

4.3.2 Transformation assays ...... 60

4.3.3 Quantification of ß-glucuronidase released ...... 61

4.3.4 Quantitative real-time PCR (qRT-PCR) analyses ...... 61

4.3.5 XIP quantification ...... 62

4.3.6 RcrR purification and electrophoretic mobility shift assays (EMSAs) ...... 62

4.4 Results ...... 63

4.4.1 RcrR regulates comS expression and extracellular XIP levels...... 63

4.4.2 Increased competence of ∆SMRcrP is the result of an increase in endogenous levels of XIP ...... 66

4.4.3 The lack of the RcrQ transporter affects the amount of secreted XIP ...... 69

4.4.4 High expression of the full-length ComS peptide inhibits exogenous XIP activity ...... 71

4.5 Discussion ...... 74

Chapter 5: Transcriptional profiling of the oral Streptococcus mutans in response to the competence signaling peptide XIP ...... 77

5.1 Abstract ...... 78

5.2 Introduction ...... 79

5.3 Materials and methods ...... 81

5.3.1 Strains and growth conditions ...... 81

5.3.2 RNA extraction and preparation for sequencing ...... 81

5.3.3 Preparation of mRNA libraries for Illumina deep-sequencing (RNAseq) .. 81

5.3.4 Data analysis ...... 82

5.3.5 qRT-PCR ...... 82

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5.3.6 Northern blot detection ...... 83

5.3.7 Rapid amplification of cDNA ends (RACE) ...... 84

5.3.8 Transformation assays ...... 84

5.4 Results ...... 85

5.4.1 Transcriptome changes in S. mutans UA159 in response to XIP ...... 85

5.4.2 Changes in gene expression in ∆SMcomS strain in response to 1 µM XIP ...... 88

5.4.3 Putative small RNAs expressed in the presence of XIP ...... 92

5.5 Discussion ...... 95

5.6 Conclusions ...... 97

5.7 Acknowledgements ...... 98

Chapter 6: Summary, future directions and significance ...... 99

6.1 Summary ...... 99

6.2 Future directions...... 102

6.3 Significance ...... 103

Supplemental Tables ...... 105

References ...... 135

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List of Figures and Tables

Figure 1.1 Dental plaque succession…………………………………………………………5

Figure 1.2 Models of DNA uptake……………………………………………………………23

Figure 1.3 Genetic competence regulation in anginosus and mitis groups of streptococci (A), pyogenic, bovis and salivarius groups of streptococci (B), and mutans streptococci (C)…………………………………………………………………....26

Figure 3.1 TF of Streptococcus mutans strains in THYE and CDM growth media…..…42

Figure 3.2 Secretion of XIP by S. mutans UA159 and comX-, comR-, and comS- mutant strains……………………………………………………………………..………..44

Figure 3.3 Effect of CSP and XIP on cell survival of S. mutans……………….…………46

Figure 3.4 Gene expression of comR, comS, and comX in response to CSP and XIP…………………………………………………………………………….……48

Figure 3.5 Differential regulation of comRS in THYE and CDM media grown in the presence or absence of CSP and XIP, respectively……………...... ………...50

Figure 3.6 Model for competence and cell killing pathways of S. mutans in THYE vs. CDM………………………………...... ……………………..…………..….53

Figure 4.1 The effect of ∆SMrcrR on XIP-associated phenotypes, comS expression and extracellular levels of XIP…………………………………………………...... …65

Figure 4.2 Electrophoretic mobility shift assays of RcrR protein binding to promoters of rcrRPQ, comS and fruA…………………………………………………………..66

Figure 4.3 The effect of ∆SMrcrP on XIP-associated phenotypes, comS expression and extracellular levels of XIP………………………………………………………...68

Figure 4.4 The effect of ∆SMrcrQ on XIP-associated phenotypes, comS expression and extracellular levels of XIP………………………………………………………...70

Figure 4.5 Expression of oppD, pep1 and pep2 in ∆SMrcrQ compared to WT UA159………………………………………………………………………………72 ix

Figure 4.6 Transformation frequency in UA159, ∆SMcomS, ∆SMrcrQ, and ∆SMcomS/rcrQ double deletion strain in the presence or absence of XIP………………………………………………………………………………….72

Figure 4.7 Transcription of pep1 and pep2 competence inhibitory peptides in ∆SMcomS, ∆SMrcrQ and ∆SMcomS/rcrQ as compared to UA159…………………….....73

Figure 5.1 Functional characterization of the changes (>2-fold) in the UA159 transcriptome in response to 1 µM XIP…………………………………………86

Figure 5.2 The effect of increasing amounts of XIP on genes involved in bacteriocin regulation (a) and sugar uptake (b)……………………………………………..87

Figure 5.3 Functional characterization of the changes (>2-fold) in the ∆SMcomS transcriptome in response to 1 µM XIP…………………………………………90

Figure 5.4 Expression of selected genes in ∆SMcomS in response to increasing concentrations of XIP……………………………………………………………..91

Figure 5.5 Expression of selected intergenic regions in response to increasing concentrations of XIP……………………………………………………………..94

Figure 5.6 The effect of ∆SMU.82-83 on transformation frequency……………………...94

Figure 6.1 Overview of the current model for ComRS signaling in S. mutans including specific contributions from chapters of this dissertation……………………..101

Table 5.1 Genes up- or down-regulated in S. mutans UA159 in response to 1 µM XIP……………………………………………………………………………….....93

Table S5.1 Genes up- or down-regulated in ∆SMcomS in response to 1 µM XIP…....105

Table S5.2 Intergenic regions up- or down-regulated in UA159 in response to 1 µM XIP………………………………………………………………………………...124

List of Abbreviations

ATP adenosine triphosphate CCR carbon catabolite repression CDM chemically defined medium cDNA complementary DNA CFUs colony forming units CSP competence stimulating peptide DMSO dimethyl sulfoxide DNA deoxyribonucleic acid dsDNA double-stranded DNA eDNA extracellular DNA EDTA ethylenediaminetetraacetic acid EMSA electrophoretic mobility shift assay EPS extracellular polysaccharides Ftf fructosyltransferase Gbp glucan binding protein Gtf glucosyltransferase GUS β-glucuronidase h hour HK histidine kinase IPS intracellular polysaccharides IPTG isopropyl β-D-1-thiogalactopyranoside LC liquid chromatography mRNA messenger RNA MS/MS tandem mass spectrometry MSM multiple sugar metabolism MU Miller units OD optical density ORF open reading frame PBS phosphate buffered saline

PCR polymerase chain reaction PEP-PTS phosphoenolpyruvate sugar: phosphotransferase system pH potential hydrogen (p)ppGpp pyrophosphorylated GDP or GTP qRT-PCR quantitative real time PCR RACE rapid amplification of cDNA ends RNA ribonucleic acid RNA-seq RNA sequencing RR response regulator rRNA ribosomal RNA sCSP synthetic CSP spec spectinomycin spp species ssDNA single-stranded DNA sXIP synthetic XIP TCSTS two-component signal transduction system TF transformation frequency THYE Todd Hewitt yeast extract UV ultraviolet v/v volume per volume wt/vol weight per volume XIP comX-inducing peptide

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Publications

Publications reproduced as dissertation chapters

Wenderska, IB. Lukenda, N. Cordova, M. Magarvey, N. Cvitkovitch, DG. Senadheera, DB. (2012) A novel function for the competence inducting peptide, XIP, as a cell death effector of Streptococcus mutans. FEMS Microbiol Lett 336(2):104-12.

Additional publications

Wenderska, IB. Chong, M. Wright, GD. and Burrows LL. Palmitoyl-DL-carnitine affects multiple pathways that influence P. aeruginosa biofilm development. ChemBioChem, 12(18):2759-66.

Nguyen, UT. Wenderska, IB. Chong, MA. Koteva, K. Wright, GD. and Burrows LL. Small molecule modulators of Listeria monocytogenes biofilm development. Appl Environ Microbiol 78(5): 1454-65.

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Awards

2012-2015 Ontario Graduate Scholarship

2012-2014 Wilson G. Harron Scholarship

2014 American Society for Microbiology Travel Award

2014 Third Prize for Research Day poster presentation

2013 Second Prize CADR-NCOHR Student Research Award

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Chapter 1: Literature Review

1.1 The oral environment

1.1.1 The oral cavity

As the gateway to the human gastrointestinal tract, the oral cavity is a complex and dynamic environment that houses one of the most diverse microbial communities associated with the body (Eren et al., 2014; The Human Microbiome Project Consortium, 2012; Wade, 2013). It is the only anatomical location that contains exposed hard non-shedding tissue surfaces which provide a unique location for the successive accumulation of a diverse microbial community referred to as dental plaque. The microbial composition of plaque is associated with oral health, and its accumulation and maturation is described in further detail in section 1.1.2. Aside from teeth and its surrounding mucosal surfaces, the oral cavity is also comprised of hard palate, tongue and the floor of the mouth, all of which provide distinct habitats for microbial colonization (Eren et al., 2014; The Human Microbiome Project Consortium, 2012; Wade, 2013).

Fluctuations in the oral environment, including variations in temperature, pH, oxygen levels and the frequency and content of dietary intake influence the composition of the microbial community. For example, oxygen levels vary greatly between the dorsal surface of the tongue (16%) and the pits and fissures of teeth (no detectable oxygen), and dictate which microbes can colonize each locale, depending on their sensitivity to oxygen and requirement for growth (Eskow and Loesche, 1971; Kenney and Ash Jr, 1969; Loesche, 1969).

Bacteria adapted to live in the oral cavity also have mechanisms to contend with drastic variations in carbohydrate levels. Following the ingestion of food, these microbes cope with increases in sugar levels up to 10,000-fold (Carlsson and Hamilton, 1994). Many oral bacteria are capable of rapid metabolism of these carbohydrates. In some instances, carbohydrate metabolism leads to the production of acid by-products, and the acidification of the local environment (Geddes, 1975; Miller et al., 1940; Stephan, 1940, 1944; Stephan and Hemmens, 1947). As a result, the pH of dental plaque has been shown to vary from above 8.0 on smooth surfaces to below 4.0 in carious lesions (Dirksen et al., 1963). In between meals, when concentrations of free carbohydrates are

3 low, oral bacteria are also able to utilize components of saliva as their source of nutrients (Bradshaw et al., 1994; De Jong and Van der Hoeven, 1987; De Jong et al., 1984; Kolenbrander, 2011; Palmer et al., 2001). Aside from providing a source of nutrients, saliva is able to buffer the organic acids produced by microbial metabolism (Kleinberg and Jenkins, 1964). Saliva contains a bicarbonate buffering system that works to neutralize temporary shifts in the oral pH (Bardow et al., 2000; Lilienthal, 1955). Other salivary components, including antimicrobial factors such as lysozyme, lactoferrin and immunoglobulin A (IgA), modulate the resident oral microflora, and can prevent colonization by foreign pathogens (Groenink et al., 1999; Kilian et al., 1981; Laible and Germaine, 1985; Reinholdt and Kilian, 1987; Williams and Gibbons, 1972). The sheer force of salivary flow further sweeps away bacteria unable to adhere to the surfaces of the oral cavity (Bloomfield, 1919, 1920). As a result, all bacteria commonly found residing in the oral flora have the ability to adhere to and form sessile microbial communities known as biofilms.

1.1.2 Dental plaque and microbial succession

Due to its accessibility and the ease of sampling, dental plaque is one of the best-described microbial biofilms associated with the human body (Eren et al., 2014; The Human Microbiome Project Consortium, 2012; Wade, 2013). It is comprised of approximately 300 species of bacteria that assemble on the tooth in a successive manner starting with the primary colonizers (Keijser et al., 2008) (Figure 1). The primary colonizers are microbes that directly interact with and colonize the salivary pellicle on the surface of the tooth. The salivary pellicle contains saliva-derived proteins, glycoproteins and lipoproteins that coat the tooth almost immediately after brushing (Hannig, 1999; Hay, 1967, 1969; Kraus et al., 1973; Lee et al., 2013; Yao et al., 2001). Some of the salivary components, named cryptitopes, only expose their microbial receptors after binding to the tooth surface (Gibbons, 1989). Streptococci constitute 60- 90% of the early colonizers, and include Streptococcus gordonii, Streptococcus oralis, Streptococcus intermedius, Streptococcus mitis and Streptococcus sanguinis (Nyvad and Kilian, 1987). These pioneer species have been shown to bind an array of proteins

4 in saliva, including mucins, agglutinins, α-amylase and proline-rich peptides (Brady et al., 1992; Gibbons et al., 1991; Murray et al., 1992; Scannapieco et al., 1989; Scannapieco et al., 1995). Multiple adhesion receptors may confer a major selective advantage for the primary colonizers over other oral bacteria with less flexibility in receptor recognition. Aside from binding to the salivary pellicle, streptococci exhibit extensive coaggregation not only with other members of its genus but also with other genera of oral bacteria. Interactions have been observed with other early colonizers including members of the Actinomyces spp., Capnocytophaga spp., Haemophilus spp., Eikenella spp., Prevotella spp., Propionibacterium spp., and Veillonella spp. (Ciardi et al., 1987; Gibbons and Nygaard, 1970; Hughes et al., 1992; Hughes et al., 1988; Kolenbrander and Andersen, 1984, 1986; Kolenbrander and Williams, 1981, 1983; Weiss et al., 1990).

In addition to the physical interactions, the structure of the microbial community is also dependent on the nature of the pioneer species and their metabolic needs. Close associations of microbial species within the community often reflect metabolic synergies and dependencies for growth. For example, the lactate end-product of streptococcal metabolism is used as a nutrient source by members of the Veillonella spp. (Mikx and Van der Hoeven, 1975). Colonization of certain species can also prevent or retard the colonization of another species. This has been observed in the antagonistic relationship between S. sanguinis and S. mutans, where early colonization of S. sanguinis delays the colonization of S. mutans resulting in lower rates of dental caries (Caufield et al., 2000). Hydrogen peroxide produced by S. sanguinis inhibits the growth and colonization of S. mutans (Kreth et al., 2008). If colonized first, S. mutans in turn produces antimicrobial peptides, termed bacteriocins or mutacins (specifically mutacin I and IV), to inhibit the growth S. sanguinis (Kreth et al., 2005). Colonization of the teeth by a commensal microorganism early on during plaque development may therefore be beneficial towards establishing a healthy microbial community.

Figure 1.1 Dental plaque succession. The early colonizers including Streptococcus and Actinomyces spp., adhere to the salivary pellicle coating of the tooth enamel, and provide attachment sites and a favourable environment for the colonization of Corynebacterium. The Corynebacterium taxa functions as a bridging organism providing attachment sites for the successive accumulation of other microbes including Fusobacterium, Leptotrichia, and Capnocytophaga. Image adapted from Welch et al., 2016.

The streptococci and Actinomyces species of the early microbial community interact with Corynebacterium, a highly abundant taxa in dental plaque (Welch et al., 2016). The Corynebacterium filaments grow outward from the tooth, and provide

6 structure to the plaque biofilm and attachment sites for succeeding organisms (Welch et al., 2016). Combinatorial Labeling and Spectral Imaging Fluorescence In Situ Hybridization (CLASI-FISH) revealed hedgehog structures within dental plaque that contain Corynebacterium filaments at the base, radiating outwards, and interacting with stratified zones of succeeding organisms (Welch et al., 2016) (Figure 1.1). The periphery of the structure contains taxa able to participate in aerobic respiration, including Streptococcus and Haemophilus/Aggregatibacter. Also present was the Porphyromonas species, and to a lesser extent Neisseriaceae. Within this outer layer, further organization was observed where the Haemophilus/Aggregatibacter group was observed only in the presence of streptococci. The lactate, produced by streptococcal metabolism, is a preferred growth substrate for Aggregatibacter (Brown et al., 2008), as a result the close association may reflect a metabolic dependency. Below the periphery lies a filamentous annulum, containing the anaerobes Fusobacterium, Leptotrichia and Capnocytophaga. The anaerobic Fusobacterium nucleatum has been shown to further facilitate the attachment of the “red complex” species consisting of Treponema denticola, Porphyromonas gingivalis and Tannerella forsythia (Kolenbrander and Andersen, 1986; Kolenbrander et al., 1989; Kolenbrander et al., 1995; Socransky et al., 1998). The presence of the red complex group of bacteria confers an increased risk for periodontal disease (Socransky et al., 1998).

The mature dental plaque contains hundreds of microbial species embedded in an exopolysaccharide matrix of host and microbial origin. If the plaque is removed regularly and good oral hygiene is maintained, then the successive accumulation of the microbial community repeats. Plaque that is inaccessible to removal can modulate polymicrobial infections of the oral cavity as a result of dysbiosis that fosters a community dominated by pathogenic organisms within the plaque biofilm. If systematically disseminated, plaque bacteria can cause infections of the cardiovascular, musculoskeletal and nervous systems (Bartzokas et al., 1994; Debelian et al., 1994; Herzberg and Meyer, 1996; Marks et al., 1988).

1.1.3 Dental caries: microbial community in health and disease

In 1940 Stephan showed a rapid decline in plaque pH following a sugar rinse that was linked to the production of organic acids by oral bacteria (Stephan, 1940, 1944). Sustained plaque pH below 5.5 favours demineralization of the enamel and dental caries (tooth decay) (Stephan, 1944). Dental caries is considered one of the most significant global oral health burdens, experienced by over 95% of the population by the time they reach middle age (Cooney, 2010). If left untreated tooth decay can progress from enamel, through the dentin to the pulp of the tooth, where it can cause infection or an abscess (Hahn et al., 1991; Hahn and Liewehr, 2007).

Dental caries is a multifactorial disease that requires a carbohydrate-rich diet, a susceptible site (the tooth surface), and the presence of caries-causing microbes. Lactobacilli and mutans streptococci have been identified as the major contributors to enamel demineralization (Krasse, 1988). Among the mutans group, S. mutans has been considered as the major etiological agent of tooth decay (Clarke, 1924; Fitzgerald and Keyes, 1960; Kristoffersson et al., 1985; Loesche, 1986; Zinner et al., 1965).

The mineral of our teeth is composed of hydroxyapatite crystals that contain many impurities, including the incorporation of carbonate ions that substitute phosphate in the crystal lattice (Kühl and Nebergall, 1963; Robinson et al., 2000; Zapanta- LeGeros, 1965). Approximately 1 out of 10 phosphate ions in the enamel are replaced by carbonate, creating defects and calcium-deficient regions that are more susceptible to acid solubility (Aoba and Moreno, 1990; LeGeros and Tung, 1983; Robinson et al., 2000). Demineralization of the tooth occurs when organic acids, produced from fermentable carbohydrates by the aforementioned cariogenic bacteria, diffuse into the water among the crystals and reach a susceptible site on the crystal surface, causing dissolution of calcium and phosphate ions. Initially the demineralization that occurs is reversible and can be repaired by the process of remineralization, facilitated by salivary components (Cate and Arends, 1977; Johansson, 1965; Koulourides et al., 1965). The saliva contains calcium and phosphate ions in a supersaturated state (Hay et al., 1982; Moreno et al., 1979). These high concentrations of ions can be used to rebuild the crystal lattice by building onto the existing crystal remnants. This process is further

8 facilitated by the presence of fluoride, often found in toothpaste or in some areas of the world, in the water supply (Jenkins, 1985; Petersson and Bratthall, 1996). During the remineralization process fluoride is incorporated into the enamel by substituting hydroxyl ions in the crystal structure to create fluorapatite, which is less soluble and therefore less susceptible to demineralization than natural enamel (Kay et al., 1964; Moreno et al., 1974; Robinson et al., 2000). The importance of saliva, and fluoride, in the process of remineralization and the maintenance of good oral health is well established (Humphrey and Williamson, 2001; Petersson and Bratthall, 1996). Patients with impaired salivary flow (hyposalivation, xerostomia), due to antibiotic treatments, radiation or genetic disorders (Sjögren’s syndrome), do not have sufficient calcium and phosphate ions available for the remineralization process, and as a result suffer from rampant caries (Christensen et al., 2001; Guggenheimer and Moore, 2003). In a healthy individual, dental caries occurs with increased carbohydrate intake and prolonged acidification of dental plaque that shifts the demineralization/remineralization equilibrium towards enamel dissolution and subsequent cavitation (Kleinberg and Jenkins, 1964; Stephan, 1940, 1944). This equilibrium imbalance correlates with a microbial shift in the dental plaque favouring the growth of acidogenic (acid-producing) and aciduric (acid- tolerant) bacteria (Bradshaw et al., 1989; Dennis et al., 1975; Staat et al., 1975).

1.2 Streptococcus mutans

In 1924, J. Clarke isolated what appeared to be an oval-shaped mutant of streptococci from human carious lesions, and thus named it Streptococcus mutans (Clarke, 1924). Decades later S. mutans was recognized as one of the major etiological agents of dental caries (Fitzgerald and Keyes, 1960; Kristoffersson et al., 1985; Zinner et al., 1965), and as a result gained much attention from the scientific community. Since then research has focused on the pathogenesis of the organism, identifying several main virulence factors associated with its cariogenic potential. Unlike some other pathogens, which secrete typical virulence factors including toxins that damage the host, the pathogenicity of S. mutans is primarily linked to its metabolism (Fitzgerald et al., 1989; Hillman et al., 1996; Johnson et al., 1980). S. mutans is highly effective at

9 producing large amounts of organic acids as a byproduct of the fermentation of a wide variety of dietary sugars (acidogenicity) (Carlsson and Griffith, 1974; Dashper and Reynolds, 1996; Stephan and Hemmens, 1947; Yamada and Carlsson, 1975). Resulting acid accumulation causes a decrease in the plaque pH and consequent demineralization of the tooth enamel (LeGeros and Tung, 1983). To withstand the low environmental pH, S. mutans has also developed mechanisms to tolerate highly acidic conditions (aciduricity) (Bender et al., 1986; Hamilton and Buckley, 1991; Harper and Loesche, 1983, 1984). This resiliency along with its ability to maintain glycolysis at pH levels well below that needed for enamel demineralization, are major contributors to the cariogenicity of S. mutans (Harper and Loesche, 1983, 1984).

S. mutans has also evolved a number of mechanisms to adapt to a life in the oral cavity, and outcompete the non-cariogenic commensals when the conditions are favourable. It has evolved mechanisms to attach to the tooth surface and form resilient biofilm communities. This sessile lifestyle is essential for survival in the oral cavity, as cells that cannot adhere to the enamel are eliminated by the host salivary flow (Bloomfield, 1919, 1920). Residing in the dental plaque on the surface of teeth, S. mutans has further developed mechanisms to sense and adapt to the ever-changing environment of the oral cavity (Smith and Spatafora, 2012). All of these adaptive mechanisms and stress responsive pathways contribute to the survival and persistence of S. mutans in the oral cavity and thus contribute to the virulence of the organism.

The main virulence attributes of S. mutans are discussed in greater detail in the sections below. These include: S. mutans adhesion properties and significant contributions to the formation of the extracellular matrix of dental plaque; its capacity to produce large amounts of acids and the ability to withstand these acidic conditions; its ability to produce antimicrobial peptides against closely related species to compete for common resources; the ability of S. mutans to acquire genetic diversity by genetic transformation; and its ability sense and respond to environmental changes by regulating the aforementioned virulence attributes through two-component signaling systems.

1.2.1 Attachment to the tooth

The initial attachment of S. mutans to the tooth surface is mediated by two main mechanisms: a sucrose-independent manner that is largely facilitated by the surface adhesion protein P1 (SpaP, antigen I/II, Pac), or a sucrose-dependent manner that is mediated by the aggressively sticky polymers of glucose (glucans) (Bowen et al., 1991; Gibbons and Nygaard, 1968; Jordan and Keyes, 1966). Following initial attachment, bacterial cells multiply and form microcolonies embedded in a matrix of polysaccharides, proteins, and nucleic acids. This exopolymeric matrix allows the bacterial community to develop a complex, three-dimensional structure (Koo et al., 2010; Liao et al., 2014; Lynch et al., 2007; Xiao et al., 2012).

In the absence of sucrose, the surface adhesin P1 mediates adhesion by directly interacting with the salivary agglutinin present in the salivary pellicle coating on the tooth (Bowen et al., 1991; Brady et al., 1993; Crowley et al., 1999; Crowley et al., 1993; Lee et al., 1989; Russell et al., 1980). Other surface associated proteins including WapA (Antigen III) and BrpA have also been implicated in sucrose-independent biofilm formation of S. mutans (Wen et al., 2006; Zhu et al., 2006). A deletion strain of either WapA or BrpA leads to perturbed biofilm formation and architecture (Wen et al., 2006; Zhu et al., 2006).

In the presence of dietary sucrose, glucan polymers facilitate attachment of cells to the tooth pellicle, and play an important role in the initiation of biofilm formation under these conditions (de Stoppelaar et al., 1971; Gibbons and Nygaard, 1968; Jordan and Keyes, 1966; Larrimore et al., 1983; Tanzer et al., 1985). These aggressively sticky polymers are generated from sucrose by the glucosyltransferase (Gtf) enzymes (Aoki et al., 1986; Hanada and Kuramitsu, 1988, 1989; Schilling and Bowen, 1988; Tamesada et al., 2004). S. mutans has been shown to produce three types of Gtfs (GtfB, GtfC and GtfD), which vary in the glucosidic linkages they assemble. GtfB and GtfC produce primarily water insoluble α-1,3-linked glucan polymers, whereas GtfD produces water soluble α-1,6-linked glucans (Aoki et al., 1986; Hanada and Kuramitsu, 1988, 1989; Mukasa et al., 1989). The cooperative action of all three enzymes is important for cellular adhesion (Ooshima et al., 2001; Tamesada et al., 2004). The

11 fructosyltransferase (Ftf) enzyme is also utilized by S. mutans to produce a third type of polymer composed of fructan monomers (Carlsson, 1970; Manly and Richardson, 1968). Due to its diffusion-limiting nature, insoluble glucan traps the organic acids produced by the acidogenic members of the biofilm (Xiao et al., 2012). This results in microenvironments of low pH that not only contribute to the dissolution of enamel (Robinson et al., 2000), but also promote conditions that reduce bacterial diversity in the plaque (Li et al., 2007; Preza et al., 2008). The fructan polymer and the water-soluble glucan produced by GtfD function as extracellular energy storage, and are believed to contribute to caries formation by allowing sugar metabolism and acid production beyond sucrose utilization (DaCosta and Gibbons, 1968; Manly and Richardson, 1968; Schroeder et al., 1989; Wood, 1967). Bacterial interactions with the glucan polymers are further reinforced by surface-associated glucan binding proteins (Gbps). S. mutans produces four different Gbps, namely GbpA, GbpB, GbpC and GbpD, all of which work together with the Gtfs to shape the architecture of the S. mutans biofilm (Banas and Vickerman, 2003; Lynch et al., 2007; Russell, 1979; Sato et al., 1997; Shah and

Russell, 2004; Smith et al., 1994).

Aside from sugar polymers, extracellular DNA (eDNA) also provides a structural component to the biofilm matrix (Liao et al., 2014; Perry et al., 2009). eDNA generated from the lysis of a subpopulation of the community has been shown to play a role in the maintenance of mature S. mutans biofilms (Perry et al., 2009). DNase treatment of S. mutans biofilms resulted in reduced biofilm biomass (Perry et al., 2009).

1.2.2 Sugar metabolism and acidogenicity

An increase in sugar consumption correlates with an increase in dental caries (Kleinberg and Jenkins, 1964; Stephan, 1940, 1944). In 1881, Miller attributed this correlation to the acid produced from fermentable carbohydrates by oral bacteria (Miller, 1890). S. mutans, as one of the main causative agents of caries, has the ability to ferment a variety of different dietary carbohydrates as part of its metabolism and energy production, including sucrose, glucose, fructose, lactose, mannose and galactose

(Stephan and Hemmens, 1947). For each sugar, S. mutans contains specialized machinery for import and metabolism into smaller end products that can enter the glycolytic pathway (Ajdic and Pham, 2007).

During periods of excess sugar intake, S. mutans accumulates intracellular and extracellular polysaccharides (IPS and EPS) as energy reserves for times of famine. The IPS are glycogen-like polymers accumulated by DaltA-D enzymes (Harris et al., 1992). The EPS, primarily used for storage, include the fructan and the water-soluble glucan polymers whose synthesis is catalyzed from dietary sucrose by the aforementioned Ftf and GtfD enzymes, respectively (DaCosta and Gibbons, 1968; Manly and Richardson, 1968; Wood, 1967). During famine, fructanases and dextranases are used to release carbohydrates from fructans and glucans, respectively, for import and use in glycolysis (Jacques et al., 1985; Pulkownik and Walker, 1977; Walker et al., 1981). Sugars such as melibiose, raffinose, and isomaltotriose derived from dextrans are further taken up by the multiple sugar metabolism (MSM) transport system (Russell et al., 1992).

At limiting sugar concentrations, the phosphoenolpyruvate (PEP) sugar: phosphotransferase system (PTS) is the major system for the import of sugars (Jacobson et al., 1989; Vadeboncoeur and Pelletier, 1997). The PEP-PTS system is a multicomponent phosphorelay system that simultaneously phosphorylates the incoming sugar during uptake. The PEP-PTS system is comprised of three major components: Enzyme I (EI), the histidine-protein (HPr), and the substrate-specific Enzymes II (EII) (Jacobson et al., 1989; Kundig et al., 1964). EI begins the phosphorylation cascade by autophosphorylating using PEP as the phosphoryl donor. The phosphoryl group is then transferred to the His-15 residue of HPr. From there it is transferred to the A domains of various substrate-specific transporters (EIIAs). Finally, the phosphate group is transferred to the B domain (EIIB), from where it is relocated to the incoming sugar during its import through the transmembrane C domain of EII (EIIC). The PEP-PTS system transports a variety of carbohydrates including glucose, fructose, sucrose and lactose (Ajdic and Pham, 2007; Brown and Wittenberger, 1973; Calmes, 1978; Cvitkovitch et al., 1995; Gauthier et al., 1984; Maryanski and Wittenberger, 1975; Neron and Vadeboncoeur, 1987; Schachtele and Mayo, 1973; Slee and Tanzer, 1979;

Vadeboncoeur, 1984). The imported sugars enter the glycolytic pathway to produce ATP and organic acid end-products.

Aside from facilitating sugar uptake, components of the PTS system also influence other cellular processes, including biofilm development, acid tolerance and the global regulation of gene expression via carbon catabolite repression (CCR) (Abranches et al., 2006; Abranches et al., 2003; Chen et al., 1998; Loo et al., 2003; Moye et al., 2014; Zeng and Burne, 2008). CCR involves regulators that activate or silence genes in response to carbohydrate availability. Specifically, the presence of a preferred carbon source, often glucose, prevents the uptake and metabolism of other, secondary, carbon sources (Dills and Seno, 1983; Liberman and Bleiweis, 1984). In low-GC Gram-positive bacteria CCR is controlled by HPr and the carbon control protein A (CcpA) transcriptional regulator (Fujita et al., 1995; Lokman et al., 1997; Simpson and Russell, 1998). During CCR, glycolytic intermediates, such as fructose-1,6-bisphosphate (F-1,6- bP) or glucose-6-phosphate, phosphorylate HPr at serine-46 at the expense of ATP (Lodge and Jacobson, 1988; Mimura et al., 1987). The HPr-(Ser-46-P) complexes with CcpA and stimulates its binding to conserved catabolite responsive elements (CRE) in the promoters of a variety of genes (Abranches et al., 2008; Zeng et al., 2013). In addition to repressing processes involved in the utilization of secondary carbon sources, CcpA has also been shown to regulate the expression of several virulence attributes (Abranches et al., 2008; Zeng et al., 2013).

Depending on growth conditions, S. mutans produces lactate, formate, acetate and ethanol as by-products of fermentation (Carlsson and Griffith, 1974; Yamada and Carlsson, 1975). During growth with excess carbohydrates, lactic acid is the predominant glycolytic end product (Carlsson and Griffith, 1974; Yamada and Carlsson, 1975). Formate, acetate and ethanol are mainly produced under glucose limiting conditions (Carlsson and Griffith, 1974; Yamada and Carlsson, 1975). Due to its low pKa value, lactic acid is able to demineralize the enamel more effectively than the other glycolytic by-products (Margolis et al., 1985).

1.2.3 Aciduricity

S. mutans is capable of maintaining glycolytic activity at pH values as low as pH 4.4, which provides it with a competitive advantage over other oral streptococci that cannot endure such highly acidic conditions (Harper and Loesche, 1983, 1984). This advantage allows S. mutans to become a dominant member of the dental plaque under conditions favouring caries formation (Minah and Loesche, 1977a, 1977b).

S. mutans employs several mechanisms to cope with low pH conditions. While some of these acid tolerance mechanisms are constitutive, others are inducible in response to a sub-lethal pH of approximately 5.5, and prepare the bacteria for enhanced survival at pH as low as 3.0 (Matsui and Cvitkovitch, 2010). The main mechanism for coping with highly acidic conditions is through the extrusion of protons by the constitutively expressed F1F0 ATPase proton pump (Bender et al., 1986). This allows S. mutans to maintain an alkaline intracellular pH relative to its environment. As the pH falls, the activity of the F1F0 ATPase increases, maintaining the intracellular ΔpH at 0.5-1 pH units relative to the external milieu (Hamilton and Buckley, 1991; Sturr and

Marquis, 1992). Studies have shown that the F1F0 ATPase can also function as an ATP synthase in staved cells, providing them with energy for growth under acidic conditions. (Dashper and Reynolds, 1992; Sheng and Marquis, 2006).

The inducible acid tolerance response includes changes in the membrane profile, increased production of alkaline products and the upregulation of mechanisms involved in DNA repair and protein biogenesis (Boyd et al., 2000; Hamilton and Buckley, 1991). The fatty acid profile of the membrane alters to incorporate longer mono-unsaturated fatty acids (C18:1 and C20:1), resulting in decreased permeability of the membrane to protons (Fozo and Quivey, 2004b; Quivey et al., 2000). A deletion of fabM, responsible for the generation of mono-unsaturated fatty acids, results in increased sensitivity to acidic conditions and the inability to maintain a ΔpH across the membrane (Fozo and Quivey, 2004a). S. mutans has also been shown to utilize the agmatine deiminase system (AgDS), to convert agmatine, a derivative of arginine, to produce ammonia and ATP (Griswold et al., 2004; Griswold et al., 2006). The ammonia produced helps increase the cytoplasmic pH, while the ATP can be used to power the F1F0 ATPase for

15 proton efflux (Griswold et al., 2006). All of the aforementioned acid tolerance processes work together to limit the amount of protons entering the cell and maintain a relatively alkaline intracellular pH.

Other inducible acid tolerance mechanisms are used to preserve the macromolecules of the cell from acid damage. The glycosidic bonds of deoxyribonucleotides are unstable under acidic conditions. A buildup of acid inside the cell can therefore cause loss of purines and pyrimidines from DNA due to the protonation of the nitrogenous base and subsequent cleavage of the glycosyl bond (Lindahl and Nyberg, 1972). The resulting abasic or AP sites are recognized by the DNA repair AP endonucleases, that are upregulated in response to low pH conditions, to repair the phosphodiester bond (Hahn et al., 1999; Sancar, 1996). Molecular chaperones are also upregulated in response to acidic conditions to prevent accumulation and aggregation of improperly folded proteins that may result in cell toxicity. In S. mutans, the GroEL and DnaK chaperones are induced in response to acid shock, to facilitate the folding process of new or denatured proteins and the transport, assembly and degradation of other proteins (Craig et al., 1993; Jayaraman et al., 1997).

1.2.4 Two-component signal transduction systems

In order to persist in the oral cavity, S. mutans employs 14 different two- component signal transduction systems (TCSTSs), and one orphan response regulator, to translate changes in the extracellular environment into changes in gene expression that ultimately result in appropriate changes in cellular physiology and stress adaptation (Ajdic et al., 2002; Biswas et al., 2008; Levesque et al., 2007). TCSTSs consist of a membrane-bound histidine kinase (HK) and its cognate response regulator (RR) (Stock et al., 2000). In response to environmental stimuli the HK autophosphorylates on a histidine residue, and subsequently transfers the phosphate to an aspartate of the RR. Phosphorylation results in a conformational change and activation of the RR. Activated RR directly binds to the promoters and alters the expression of specific genes under its regulation. Many of the 14 TCSTSs converge to regulate the main virulence attributes

16 of S. mutans (Ahn et al., 2006; Biswas et al., 2008; Gong et al., 2009; Levesque et al., 2007; Liu and Burne, 2009; Tremblay et al., 2009). The most studied TCSTSs in relation to S. mutans physiology are the ComCDE and VicRKX systems. The ComCDE system is comprised of the ComD HK that in response to a peptide pheromone, encoded by the comC gene, activates the ComE RR to regulate bacteriocin production, genetic competence and biofilm formation (Hung et al., 2011; Li, Lau, et al., 2001; Li et al., 2002; van der Ploeg, 2005). The ComC prepeptide contains a peptide leader region with a double glycine (GG) motif that targets the pre-peptide to a membrane bound ABC transporter (NlmTE) (Hossain and Biswas, 2012; Li, Lau, et al., 2001). During export the leader sequence is cleaved at the GG processing site. Outside the cell, ComC is further processed to its active form by the membrane bound SepM protease (Hossain and Biswas, 2012). The active 18-mer peptide, called CSP or competence stimulating peptide (named for its role in genetic transformation) interacts with the ComD HK to activate the signaling cascade. Individual deletions of the comCDE genes resulted in aberrant biofilm formation and reduced biofilm biomass (Li, Lau, et al., 2001). The regulatory role of the ComCDE system in bacteriocin production and genetic competence are further described in more detail in sections 1.2.5.1 and 1.2.6.1, respectively.

The VicRKX system has been demonstrated to regulate sucrose-dependent adhesion and biofilm formation, acid tolerance, oxidative and cell envelope stress response, genetic competence and bacteriocin production (Deng et al., 2007; Senadheera et al., 2009; Senadheera et al., 2005; Senadheera et al., 2007). It is comprised of the VicK HK, VicR RR and the VicX protein that shares significant amino acid identity with metal-dependent hydrolases. The VicR RR is the only RR in S. mutans that is essential, and has been shown to be involved in cell division and cell wall integrity (Senadheera et al., 2005). VicR also directly binds to the promoters of the gtfBCD genes to regulate their expression, and sucrose-dependent adhesion (Senadheera et al., 2005). The role of VicRKX in bacteriocin production and genetic transformation is indirect via the ComCDE system and it is further described in sections 1.2.5.1 and 1.2.6.2, respectively.

Other S. mutans TCSTSs have been characterized, including CiaHR, LiaSR, ScnKR, and have overlapping functions to control virulence attributes of S. mutans (Ahn et al., 2006; Biswas et al., 2008; Gong et al., 2009; Levesque et al., 2007). The LevQRST signal transduction system, is involved in nutrient acquisition, specifically fructose, and is unique in that it contains four components. The LevQ and LevT proteins are membrane sensors of extracellular fructose levels that in response to low fructose stimulate the LevRS TCSTS, comprised of LevS HK and LevR RR. Activation of LevR results in the upregulation of the fruA gene, encoding a fructan hydrolase that liberates fructose form inulin, and sucrose and raffinose from fructans (Burne et al., 1987). The LevQRST system also controls the expression of the levDEFG operon, encoding the EII of a fructose/mannose PEP:PTS system, allowing for fructose uptake by the cell (Zeng et al., 2006). The LevQRST system also plays a significant role in CcpA-independent CCR, where allosteric regulation of LevR by HPr-(Ser-46-P) inhibits the expression of fruA and levDEFG genes (the CcpA-dependent CCR was briefly described in section 1.2.2) (Zeng and Burne, 2010).

1.2.5 Bacteriocins

S. mutans produces bacteriocins, also referred to as mutacins, to compete with other oral streptococci for tooth colonization (Hamada and Ooshima, 1975a, 1975b; Kelstrup and Gibbons, 1969; Yamamoto et al., 1975). Bacteriocins are ribosomally synthesized antimicrobial peptides produced by bacteria to compete against closely related species for common resources (Cotter et al., 2005; Kelstrup and Gibbons, 1969). The types produced by S. mutans vary between strains, but nearly all strains seem to produce at least one (Hamada and Ooshima, 1975b; Kelstrup and Gibbons, 1969; Yamamoto et al., 1975). Bacteriocins are classified into two main groups: lantibiotics (class I) and non-lantibiotics (class II) (Cotter et al., 2005). The lantibiotics contain post-translational modifications and are characterized by the presence of lanthionine and methyllanthionine residues (Gross and Morell, 1967, 1971; Jung, 1991; van De Ven and Jung, 1996). They typically interact with lipid II and inhibit cell wall synthesis (Breukink et al., 2003; Brotz et al., 1997; Brotz et al., 1998; Hasper et al.,

2006; Hsu et al., 2003; van Heusden et al., 2002). The non-lantibiotics do not contain any post-translational modifications with the exception of disulfide bridges, and they induce cell lysis by disrupting the membrane potential, leading to cell leakage (Abee et al., 1994; Galvez et al., 1991; Hechard and Sahl, 2002; Moll et al., 1996). S. mutans produces both types, with mutacins I, II and III, and mutacins IV, V, VI and N, being in the lantibiotic and non-lantibiotic group, respectively (Hale et al., 2005; Hillman et al., 1998; Novak et al., 1994; Perry et al., 2009; Qi et al., 1999, 2000). The S. mutans UA159 strain, of interest to our studies, has thus far been demonstrated to produce the non-lantibiotic group of mutacins, including IV, V and VI (Hale et al., 2005; Hossain and Biswas, 2011; Perry et al., 2009).

Bacteriocins are often co-expressed with immunity proteins to prevent autolysis (self-killing). Although there is no single common mechanism for conferring immunity, most bacteriocin immunity proteins are specific to their cognate bacteriocin (Nishie et al., 2012). For the non-lantibiotic group, transporter proteins are involved in bacteriocin immunity by extruding the antimicrobial peptides out of the cell (Ra et al., 1999; Stein et al., 2003). Others have been shown to bind to the bacteriocin receptor and to block pore formation (Oppegard et al., 2010).

1.2.5.1 Regulation of bacteriocin production

The expression of bacteriocins and their immunity proteins are tightly controlled to prevent unregulated and unnecessary production. In S. mutans, the ComCDE TCSTS directly regulates a variety of mutacins, mostly of the non-lantibiotic group. The ComE TR has been shown to bind a conserved direct repeat element (TCNTAAANGGT-10-TCNTAAANGGT) within the promoters of mutacin IV, V and VI, which results in marked increase in bacteriocin production (Hung et al., 2011; Kreth et al., 2005; Kreth et al., 2006; van der Ploeg, 2005; Yonezawa and Kuramitsu, 2005). Recently, the VicRKX system has been shown to inhibit mutacin production likely by inhibiting the ComCDE system (Senadheera et al., 2012). VicR has been shown to bind the VicR-box located within the open reading frame (ORF) of comC to inhibit its

19 expression (Senadheera et al., 2012). A deletion in the vicR gene results in a marked increase in bacteriocin expression, whereas overexpression of the system inhibits bacteriocin production, as demonstrated by deferred antagonism assays against the Lactococcis lactis, S. bovis and S. thermophilus indicator strains (Senadheera et al., 2012).

Other membrane-bound systems involved in the direct regulation of bacteriocin production include the LytTR family of regulatory systems, comprised of a membrane bound inhibitor protein (M) that antagonizes the activity of its associated transcriptional regulator (R) (Merritt and Qi, 2012). In S. mutans UA159 two systems of the LytTR family have been identified to regulate mutacin production: HdrRM and BrsRM. A mutation in the membrane-associated inhibitor protein results in constitutive activation of the cognate response regulator. The activated HdrR and BrsR proteins then bind the ComE binding site within mutacin promoters to increase their expression and production (Okinaga, Niu, et al., 2010; Xie et al., 2010). Therefore, unlike the VicRKX system, the HdrRM and BrsRM systems function in parallel to the ComCDE system. Furthermore, the HrdRM and BrsRM systems have been shown to coregulate each other to determine whether bacteriocin production will lead to inhibition of competitor strains or autolysis of the producer (Xie et al., 2010). Specifically, it was shown that overexpression of HdrR has a moderate effect on bacteriocin expression, that leads to increased bacteriocin production and lysis of competitor strains, while BrsR overexpression was lethal to the producer. This lethality was attributed to strong bacteriocin production (Xie et al., 2010).

Many of the lantibiotic mutacins cluster with regulatory elements that control their expression (Merritt and Qi, 2012). The loci containing mutacins I, II and III also encode a transcriptional regulator (MutR) of the Rgg-family. A deletion in the mutR gene exhibits undetectable mutacin activity, as demonstrated by the lack of antagonism against the S. sobrinus OMZ176 indicator strain (Qi et al., 1999).

1.2.6 Genetic transformation

The ability to take up and incorporate extracellular DNA (eDNA) allows bacteria to modify their genomes, leading to an increase in genetic variability and population survival under changing environmental conditions (Attaiech et al., 2011; Avery et al., 1944; Baltrus et al., 2008; Croucher et al., 2011; Engelmoer and Rozen, 2011; Griffith, 1928; Hiller et al., 2010; Michod et al., 1988). Aside from its role in genetic diversity, the process of transformation by genetically competent cells has been suggested to play a role in the utilization of eDNA as a nutrient or as a template for DNA repair of chromosomal damage (Finkel and Kolter, 2001; Michod et al., 1988; Prudhomme et al., 2006). Due to the aforementioned benefits of DNA transformation on genome plasticity and repair, this process is often activated in response to a multitude of stressors. As a result, genetic competence has been suggested to function as a general stress response mechanism in many gram-positive bacteria (Claverys et al., 2006). In Streptococcus pneunomiae, genetic competence is upregulated as part of a stress response after treatment with DNA damaging agents (Prudhomme et al., 2006; Slager et al., 2014). In S. mutans, competence deficient mutants were also defective in acid tolerance, biofilm formation, and oxidative stress response (Li, Hanna, et al., 2001; Li, Lau, et al., 2001; Li et al., 2002).

The import of eDNA by gram-positive bacteria is a complex process that involves proteins related to components of the type IV pili (T4P) and type II secretion system (T2SS) of gram-negative bacteria (Muschiol et al., 2015). The T4P, used for adhesion (Giltner et al., 2006; Rudel et al., 1995; Saiman et al., 1990), biofilm formation (Chiang and Burrows, 2003; O'Toole and Kolter, 1998), motility (Bradley, 1980; Merz et al., 2000; Skerker and Berg, 2001) and DNA uptake (Fussenegger et al., 1997; Kang et al., 2002; Stone and Kwaik, 1999; Wolfgang et al., 1998), are comprised of repeating pillin subunits that are polymerized into a long fiber using energy provided by a cytoplasmic extension ATPase (Freitag et al., 1995; Nunn et al., 1990; Turner et al., 1993). The pillins assemble at the base pushing the growing pilus structure outwards from the cytoplasmic membrane to trasverse the cell wall and outer membrane. Pilus retraction occurs by the disassembly of the pilin subunits mediated by a retraction ATPase (Maier

21 et al., 2002; Merz et al., 2000). It is believed that this ATP-driven process of extension and retraction facilitates the uptake of eDNA by gram-negative bacteria (Muschiol et al., 2015; Okamoto and Ohmori, 2002; Wolfgang et al., 1998). Although no interaction has been observed between the major pilin subunit and eDNA, recent work in N. meningitidis has shown that a minor pilin ComP is involved in DNA binding (Cehovin et al., 2013). Once the DNA is captured and transported into the periplasm, it interacts with the DNA-binding protein ComE, that delivers the DNA to a cytoplasmic channel ComA (Chen and Gotschlich, 2001; Facius et al., 1996). A single strand is impororted into the cell in parallel with the degradation of its complementary strand (Barany et al., 1983).

Similar to the T4P, the T2SS, used by gram-negative bacteria for protein secretion across the outer membrane, is composed of pilin-like subunits that assemble by the action of an ATPase (Campos et al., 2013). Unlike the long T4P fibers, the T2SS is a short periplasmic fiber (the pseudopilus), whose retraction is not required for protein secretion. In fact, the T2SS does not contain a retraction ATPase, and has been proposed to disassemble passively by collapsing or degradation (Durand et al., 2005). The current model for T2SS activity includes ATPase mediated pistin like movement that pushes periplasmic proteins across the outer membrane (Campos et al., 2013).

The macromolecular structures involved in DNA uptake in gram-positive bacteria also contain many of the components involved in the assembly of a pilin-like structure important for DNA binding and uptake (Muschiol et al., 2015). Much of the current model for DNA uptake and translocation across the membrane of gram-positive bacteria has been based on research in Bacillus subtilis and S. pneumoniae. A cell competent for DNA uptake expresses a pseudopilus comprised of the ComGC major pilin-like subunit, and the ComGD, ComGE and ComGG minor pilin-like proteins (Chung et al., 1998; Chung and Dubnau, 1995, 1998; Pestova and Morrison, 1998). In S. mutans these proteins are encoded by the nine-gene comY operon (Merritt et al., 2005). It has been proposed that the pseudopilus functions as a DNA receptor, and, like the T4P of gram-negative bacteria, its disassembly and retraction allows the bound DNA to traverse the cell wall peptidoglycan (Figure 1.2a) (Laurenceau et al., 2013). However, similar to the T2SS, no competence-associated retraction ATPase has yet been identified in gram-positive bacteria. Recent observations that pilus shedding directly

22 correlates with transformability has led to the proposal of a different model, where the pilus is assembled and released to create a temporary opening in the peptidoglycan that allows the entry and access of the DNA to the membrane-associated DNA binding protein ComEA (Figure 1.2b) (Balaban et al., 2014). In both S. pneumonae and B. subtilis, dsDNA is bound preferentially over single stranded DNA (ssDNA) (Davidoff- Abelson and Dubnau, 1973; Morrison and Guild, 1973; Piechowska and Fox, 1971). Single stranded breaks are then introduced into the dsDNA by unknown nucleases. Degradation of the nicked strand by endonuclease EndA occurs in parallel with transport of the second strand through the transmembrane channel ComEC (Dubnau, 1999). The imported ssDNA is protected from degradation by the single stranded DNA binding protein, SsbB, and the DNA processing protein A (DprA) (Berge et al., 2003; Morrison and Mannarelli, 1979). DprA further loads the RecA recombinase onto the ssDNA to facilitate RecA-mediated recombination (Mortier-Barriere et al., 2007). Recombination of the newly acquired DNA allows for integration into the host cell genome.

Figure 1.2 Models of DNA uptake. The two models are described in detail in the text of section 1.2.6. Figure adapted from Laurenceau et al., 2015.

1.2.6.1 Regulation of genetic competence: the core competence regulators

In many bacteria, including S. mutans, expression of the DNA uptake and recombination machinery is tightly regulated. In streptococci the state of competence is often regulated by small signaling peptides (reviewed in (Fontaine et al., 2015)). Many species utilize one of two main types: CSP (briefly described in conjunction with its sensory TCSTS ComDE in section 1.2.4) or XIP (ComX inducting peptide) (Figure 1.3). The anginosus and mitis groups of streptococci, including S. pneumoniae, utilize CSP for competence induction (Figure 1.3a) (Havarstein, Coomaraswamy, et al., 1995; Havarstein, Diep, et al., 1995; Havarstein et al., 1996; Hui and Morrison, 1991; Martin et al., 2006). In these strains the CSP-activated ComDE TCSTS directly upregulates the expression of the alternate sigma factor, comX. ComX controls the expression of the DNA uptake and recombination machinery. The pyogenic, bovis and salivarius groups of streptococci utilize the XIP signaling peptide of the ComRS system to induce transformation (Figure 1.3b) (Fontaine et al., 2010; Mashburn-Warren et al., 2010). ComS is a prepeptide that is secreted and processed by an unknown mechanism to produce XIP. The mature peptide is reimported into the cell, by the Opp/Ami permease, where it interacts and activates the ComR transcriptional regulator. ComR-XIP complex directly binds to the comS promoter in a positive feedback loop, and to the promoter of comX to induce its expression (Fontaine et al., 2010; Fontaine et al., 2013; Mashburn- Warren et al., 2010). The mutans streptococci have been shown to produce both CSP and XIP peptides, and although CSP signaling was the first to be discovered for its role in transformation (Li, Lau, et al., 2001), research over the last six years has established ComRS as the main competence inducing system in S. mutans (Figure 1.3c) (Fontaine et al., 2015; Reck et al., 2015).

The S. mutans ComCDE system is paralogous to the ComCDE competence system of S. pneumoniae, and homologous to the pneumococcal bacteriocin regulatory system BlpCRH (Martin et al., 2006). As a result, while the pneumococcal ComE directly regulates comX expression, in S. mutans the ComCDE system directly controls bacteriocin production (as described in section 1.2.5.1) and exerts indirect control of

25 genetic competence. The model is further complicated by the differences in growth conditions that are optimal for CSP and XIP signaling: CSP signaling for competence is optimal under rich nutrient conditions; conditions under which the activity of exogenously added XIP is inhibited, likely by nonspecific polypeptides in the medium competing for import by the oligopeptide transporter (Desai et al., 2012; Wenderska et al., 2012). Although the XIP peptide is only active under chemically defined conditions devoid of any polypeptides, the comRS genes are still essential for CSP-induced competence in rich media (Wenderska et al., 2012). While the exact mechanism of competence regulation by the ComCDE system remains unclear, the current model for comRS regulation by CSP signaling includes an increase in cell permeability to XIP by bacteriocins (Reck et al., 2015) (Figure 1.3c). Once inside the cell, XIP interacts with ComR for direct regulation of comX expression and transformability (Fontaine et al., 2013). The ComR-XIP complex also binds to the promoter of the comS peptide, creating a positive feedback loop (Fontaine et al., 2013; Mashburn-Warren et al., 2010), to ensure the entire population expresses comX (Reck et al., 2015; Son et al., 2012). Our recent observation that a comX deletion strain contains significantly less XIP in the supernatant suggests a second positive feedback loop that allows the ComX sigma factor to regulate XIP production and/or secretion (Wenderska et al., 2012).

Figure 1.3 Genetic competence regulation in anginosus and mitis groups of streptococci (A), pyogenic, bovis and salivarius groups of streptococci (B), and mutans streptococci (C).

1.2.6.1.1 The core competence regulators and competence-associated cell lysis

In addition to regulating genes involved in DNA uptake, XIP signaling and ComX induction has been demonstrated to result in cell lysis of a subpopulation of cells (Wenderska et al., 2012). A ComX-regulated lysin, LytF, has been identified as a contributor to the competence-associated lysis (Dufour and Levesque, 2013). Lysis of a subpopulation of cells could provide DNA and nutrients to the surviving population under conditions of stress and has been shown to enhance long-term survival of S. mutans (Dufour and Levesque, 2013). Furthermore, XIP also regulates comDE, and as a result transcription of bacteriocin genes (Reck et al., 2015). This tight association of competence development and bacteriocin production may provide S. mutans with a mechanism to lyse its competitors and simultaneously utilize their genetic contents to increase genome plasticity (Reck et al., 2015). Furthermore, a strong induction in bacteriocin expression has also previously been demonstrated to lead to autolysis of the producer and thus may also contribute to the lytic phenotype associated with XIP signaling (Xie et al., 2010).

1.2.6.2 Regulation of genetic competence: TCSTSs

Like in many other streptococcal species, CSP and XIP production in S. mutans is often initiated in response to specific environmental stressors including acidic, oxidative and temperature stresses, the presence of antibiotics or mutagens, and in response to nutritional signals (Biswas et al., 2008; Lemos and Burne, 2008). These environmental changes are often transduced by several two component signal transduction systems (TCSTSs) and other signaling systems, which include the ScnCHR, VicRKX, HdrMR, BsrRM, and the serine/threonine kinase PknB. Aside from their specific genetic targets, these regulatory systems often co-regulate a number of common processes including genetic transformation, biofilm formation, bacteriocin production, and stress adaptation (Fontaine et al., 2015; Smith and Spatafora, 2012). During competence regulation, several of these appear to converge upstream of ComX,

28 either at the level of ComRS or ComCDE, to control competence development. The ScnCHR (SMU.1145c-47c) signal transduction system is composed of the ScnC peptide, and the ScnHR TCSTS (Kim et al., 2013). A deletion of the scnC gene resulted in a 10-fold reduction in comX expression and competence development that could not be complemented by exogenous XIP. The ScnC peptide therefore likely controls ComRS production and/or activity. Since the phenotypes of ∆scnC could be alleviated by a deletion of the scnRH two component system, Kim et al., suggested that the ScnC peptide may induce competence development by inhibiting the ScnRH signaling system. The HdrRM signaling system, comprised of a transcriptional activator, HdrR, and its negative regulator HdrM, regulates bacteriocin production and competence development. A deletion in hdrM or overexpression of hdrR results in significant increase in comX expression, and transformation. Since the HdrR activator has been suggested to bind to ComE-binding sites, it also likely regulates competence in an indirect manner via NlmC expression (Okinaga, Niu, et al., 2010; Okinaga, Xie, et al., 2010). The BsrRM signaling system also regulates genetic competence indirectly via HdrR stimulation and bacteriocin production (Xie et al., 2010). The VicRK signaling system is a negative regulator of competence development. Activated VicR (VicR~P) inhibits CSP expression by binding to a vicR-box located within the corresponding ORF (Senadheera et al., 2012). Whether the VicRKX system affects ComRS directly has not yet been investigated. The PknB serine/threonine kinase positively regulates comCDE expression, possibly by affecting the phosphorylation state of the VicR repressor (Banu et al., 2010; Fontaine et al., 2015).

1.2.6.3 Regulation of genetic competence: The RcrRPQ operon

The rcrRPQ operon has been demonstrated to play a critical role in genetic transformation and oxidative stress tolerance in S. mutans (Seaton et al., 2011). Furthermore, this system provides the first evidence of a link between competence development and the (p)ppGpp alarmone in S. mutans (Seaton et al., 2011). Collectively referred to as (p)ppGpp, these group of signaling molecules include the guanosine 3’-diphosphate, guanosine 5’-triphosphate, and guanosine 3’,5’-

29 bispyrophosphate, and are responsible for activation of the stringent response. This highly conserved stress response, usually to nutrient starvation, involves global transcriptional changes that favour adaptation to a semidormant state, reducing growth of the population and facilitating survival during stressful conditions (Potrykus and Cashel, 2008). Coordination of genome replication with natural transformation may provide a mechanism that ensures correct chromosome rearrangements following homologous recombination of transformed DNA. This link exists in several Bacillus subtilis strains, where reduced DNA and RNA replication occurs in the competent subpopulation, and (p)ppGpp signaling has been shown to regulate genetic competence (Erickson and Copeland, 1972; Inaoka and Ochi, 2002).

RcrR is a transcriptional regulator of the MarR-family of regulators and is part of an operon containing two predicted ABC exporters: rcrP and rcrQ. It has been demonstrated that a deletion of rcrR results in a downregulation of relP, encoding a (p)ppGpp synthetase that contributes to the bulk of (p)ppGpp present during exponential growth (Seaton et al., 2011). Aside from (p)ppGpp regulation, loss of particular genes in the rcrRPQ operon results in drastic changes to the competence phenotype (Seaton et al., 2015; Seaton et al., 2011). Specifically, a non-polar (NP) deletion of rcrR results in a competence negative phenotype, while its polar (P) counterpart results in increased transformation (Seaton et al., 2011). The drastic differences between the two strains were attributed to the differences in rcrP and rcrQ expression levels. RcrR functions as a negative regulator of the rcrRPQ operon, and as a result a non-polar deletion of rcrR increases rcrPQ expression 1000-fold. In the ∆RcrR-P mutant, on the other hand, the two transporters were expressed at wild-type levels. Seaton et al., further demonstrated that a deletion of the rcrPQ resulted in an increase in natural transformation frequency, and a decreased transformation in response to exogenously added CSP. Aside from the two putative ABC exporters, Ahn et al, identified two peptides as part of the rcrRPQ operon, encoded at the end of rcrQ. Deletion of both peptides (pep1 and pep2) resulted in increase in genetic transformation in the competence negative ∆rcrR-NP strain, suggesting a function for the peptides as negative regulators of competence (Ahn et al., 2014). Since overexpression of pep1 and pep2 resulted in decreased levels of the ComX protein, the authors suggested that

30 these peptides either interfere with the ComR-XIP activity to induce comX or increase ComX degradation by ClpPCMecA complex, previously demonstrated to regulate ComX stability (Tian et al., 2013). Further it was suggested that the activity of the peptides appeared independent of the phenotypes associated with the rcrPQ transporters. Specifically, Ahn et al., demonstrated that the competence phenotypes associated with the rcrPQ genes remained unchanged regardless of the presence or absence of the peptides encoded downstream, and therefore concluded that pep1 and pep2 are not substrates for export by RcrPQ (Ahn et al., 2014). Although the association of RcrRPQ with competence is clear, the mechanism of regulation and the substrates for the two ABC transporters remains unknown.

Chapter 2: Research rationale and hypothesis

2.1 Research Rationale

Prior to the discovery of ComRS, the ComCDE TCSTS was recognized for its role in genetic transformation in S. mutans (Cvitkovitch, 2001; Li, Lau, et al., 2001). However since no binding site for ComE was observed in the promoter of comX, this regulation was considered indirect. Direct regulation by the ComCDE system was observed for bacteriocin-encoding genes, whose expression resulted in the lysis of a subpopulation of cells (Hung et al., 2011; Perry et al., 2009). The eDNA released by the competence-associated cell lysis contributes to the DNA pool available for competent bacteria. In 2010, Mashburn-Warren and colleagues identified the ComRS system as the proximal regulator of comX (Mashburn-Warren et al., 2010). The direct regulation of comX by ComR-XIP was later demonstrated through electrophoretic mobility shift assays (EMSAs) (Fontaine et al., 2013). Unlike the characterization of ComCDE, which was performed under nutrient rich conditions, the ComRS system was characterized under chemically defined conditions devoid of polypeptides that could potentially inhibit XIP activity. By synthesizing different lengths of the ComS peptide, Mashburn-Warren determined the minimal sequence for activity and termed it the mature peptide, XIP. Although the role of ComRS in direct regulation of comX was established, the identity of the naturally produced XIP peptide and its role in CSP signaling and cell lysis was not clear. Furthermore, although many stress-response regulators have been shown to regulate genetic competence and the CSP pathway, including the RcrRPQ regulon expressing drastic transformation phenotypes, due to the novelty of the ComRS system in S. mutans their role in the regulation of XIP signaling has not been determined.

2.2 Hypothesis

We hypothesize that due to the close association of genetic competence and lysis the ComRS signaling system is also a key regulator of cell lysis. We further hypothesize that RcrRPQ has a direct role in the regulation of the ComRS signaling system.

Specific Aim 1: Identify the XIP peptide naturally produced in S. mutans UA159, and examine its role in competence-associated cell lysis, and CSP-induced competence under rich nutrient conditions. This research aim was carried out by:

-Characterizing and quantifying XIP in S. mutans UA159 strain

-Quantifying of XIP in comS, comR, comE and comX deletion strains

-Examining the lytic activity of XIP

-Examining the activity of XIP and CSP in the two growth media

Specific Aim 2: Examine the role for the RcrR regulator and the two ABC transporters, RcrP and RcrQ in comRS regulation. This research aim was carried out by:

-Examining the role of RcrRPQ on transformation and competence-associated lysis

-Examining the role of RcrRPQ on comS/R and comX gene expression

-Examining the role of RcrRPQ on levels of secreted XIP

Specific Aim 3: Examine the global effects of XIP signaling on the S. mutans transcriptome. This research aim was carried out by:

-Examining the transcriptiome of UA159 and the comS deletion strain in the presence or absence of exogenously added XIP

Chapter 3: A novel function for the competence inducting peptide, XIP, as a cell death effector of Streptococcus mutans

Iwona B. Wenderska, Nikola Lukenda, Martha Cordova, Nathan Magarvey, Dennis G. Cvitkovitch, Dilani B. Senadheera

FEMS Microbiol.

Author contributions: IBW performed the experiments and analyzed the data. IBW and DBS wrote the manuscript. DBS, NM and DGC provided scientific input. MC contributed to the qRT-PCR assays and assisted with the transformation and lytic assays. NL performed LCMS/MS quantification of XIP in cell extracts.

3.1 Abstract

In Streptococcus mutans, ComX, an alternative sigma factor, drives the transcription of the “late-competence genes” required for genetic transformation. ComX activity is modulated by inputs from two signaling pathways, ComDE and ComRS, that respond to the competence stimulating peptide (CSP) and the SigX-inducing peptide (XIP), respectively. In particular, the comRS, encoding the ComR regulatory protein and the ComS precursor to XIP, functions as the proximal regulatory system for ComX activation. Here, we investigated the individual and combinatorial effects of CSP and XIP on genetic transformation and cell killing of S. mutans. Our transformation results confirm the recent reports by Mashburn-Warren et al. and Desai et al. that XIP functions optimally in a chemically defined medium (CDM), whereas its activity is inhibited when cells are grown in complex medium. Using tandem mass spectrometry (MS/MS) fragmentation, a drastic reduction in XIP levels in ComX-deficient cultures were observed, suggesting a ComX-mediated positive feedback mechanism for XIP synthesis. Our evaluation of cell viability in the presence of 10µM XIP resulted in the killing nearly 82% of the population. The killing activity was shown to be dependent on the presence of comR/S and comX. These results suggest a novel role for XIP as a compelling effector of cell death. This is the first report that demonstrates a role for XIP in cell killing.

3.2 Introduction

The acquisition of novel, heritable DNA by genetically competent bacteria not only propagates antibiotic resistance and virulence determinants, but also shapes bacterial genomes contributing to rapid evolutionary changes (Cody et al., 2003; Didelot and Maiden, 2010; Feil et al., 1999; Seifert et al., 1988).

In Streptococcus mutans, an oral resident associated with dental caries, competence development relies on multiple input systems that relay environmental signals to ultimately modulate the transcription of comX, encoding an alternate sigma factor, ComX (SigX) (Ahn et al., 2006; Mashburn-Warren et al., 2010; Okinaga, Niu, et al., 2010; Okinaga, Xie, et al., 2010; Senadheera et al., 2012). In S. mutans, competence does not develop in the absence of ComX, as it is critical for the expression of genes involved in DNA uptake and recombination (Aspiras et al., 2004). Expression of comX was first shown to be regulated by the ComDE two component signaling system comprising of a sensor kinase and a response regulator, respectively, which responds to accumulation of the competence stimulating peptide (CSP) (Aspiras et al., 2004; Cvitkovitch, 2001). Recently, Mashburn-Warren et al. (2010) identified the ComR regulatory protein of the ComRS signaling pathway as the proximal regulator necessary for comX expression. ComR, in conjunction with its cognate signal peptide, XIP (SigX inducing peptide), modulates comX transcription in S. mutans (Mashburn- Warren et al., 2010). The XIP precursor encoded by comS is consequently exported, processed to its mature form, and then internalized via the Opp/Ami transporter to interact with ComR for comX regulation (Mashburn-Warren et al., 2010). The loss of ComR abolishes comX expression and competence development, which cannot be restored by the addition of CSP. Furthermore, XIP does not require a functional comE gene to induce the expression of comX (Mashburn-Warren et al., 2010). These observations highlight the central role of ComRS in the regulation of comX.

Previously, it has been demonstrated that S. mutans cultures exposed to high CSP concentrations (2-4 µM) cause growth arrest and eventually undergo cell death by lysis (Perry et al., 2009; Qi et al., 2005). In this work, we asked whether synthetic XIP can elicit a similar response to cause cell death of S. mutans. Our viability assays

37 revealed that supplementing 10µM XIP killed approximately 82% of the population. We further report that in addition to the comR/S, the presence of comX is vital for optimal killing. Moreover, we also report the effects of XIP on genetic transformation, which support findings by Mashburn Warren et al. (2011) and Desai et al., (2012). Further, using tandem mass spectrometry, we successfully detected the 7 amino acid XIP peptide (GLDWWSL) in the wild-type UA159 supernatant, but not in that of the ComS- deficient mutant. While these results concur with those recently reported by Khan et al. (2012), we further show that supernatant XIP levels are drastically reduced in ComX- deficient cultures, suggesting a positive role for ComX in ComS/XIP production, export or processing. Taken together, in addition to its widely discussed role in competence, our work reveals a novel role for XIP as a potent effector of cell death in S. mutans, which may be potentially used for the development of therapeutic strategies to prevent dental caries.

3.3 Materials and methods

3.3.1 Bacterial strains and growth conditions

S. mutans UA159 (Ajdic et al., 2002) and its mutant strains in comC (∆SMcomC), comD (∆SMcomD), comE (∆SMcomE) (Li, Lau, et al., 2001) and comX (∆SMcomX) (Li et al., 2002) were used in this study. The comS (∆SMcomS) and the comR (∆SMcomR) mutants were constructed using a non-polar ligation PCR mutagenesis method described previously (Ajdic et al., 2002). S. mutans strains were grown at 37oC with 5%

CO2 in either Todd-Hewitt broth (Becton Dickinson, MD) containing 0.3% yeast extract (Difco Laboratories) (THYE), or chemically defined medium (CDM) described previously (Mashburn-Warren et al., 2010). Erythromycin and spectinomycin were used as needed at concentrations of 10µg/mL and 1mg/mL, respectively. Synthetic XIP (sXIP) and synthetic CSP (sCSP) peptides were synthesized using F-MOC chemistry (Advanced Protein Technology Centre, Hospital for Sick Kids, Toronto, Canada). Stock concentrations of 1µM of sXIP and 0.4mM sCSP were prepared in DMSO and water,

38 respectively. Growth kinetics were monitored using an automated growth reader (Bioscreen C; Labsystems, Finland) as previously described.

3.3.2 Transformation frequency (TF) assays

Overnight cells grown in THYE were pelleted, washed and resuspended in phosphate buffered saline (1x PBS). The resuspended culture was diluted 1:50 using prewarmed THYE or CDM and grown to an OD600 ~0.1. Next, 1µg/mL of the donor plasmid DNA (pDL277; specR) (LeBlanc et al., 1992) was added to 1mL aliquots of the culture in the presence or absence of CSP (0.4µM) or XIP (10µM) and samples were incubated for 90 min. For XIP, control cultures containing 1% DMSO were utilized. After incubation, cultures were serially diluted and plated on THYE plates with and without antibiotics. TF was calculated as transformant colony forming units (CFUs) divided by the total number of viable CFUs, times one hundred.

3.3.3 Cell viability assays

Overnight cultures in THYE were pelleted, washed, resuspended in sterile 1X PBS, and diluted 1:50 using warm THYE or CDM. Each suspension was supplemented with either 2µM CSP or 10µM XIP. Cultures without peptides or containing 1% DMSO were used as controls. All cultures were grown to an OD600~0.8, at which point, the cells were gently sonicated on ice and used for viability assays. Cells were serially diluted, plated on THYE agar and CFUs counted. Results standardized using cellular dry weight. These standardized values were then used to calculate the percentage survival by dividing the standardized number of viable cells after treatment by the standardized total number of cells without peptide, times 100.

3.3.4 Time-course killing analyses

Overnight cultures of UA159 in THYE were pelleted, washed, resuspended in sterile 1X PBS, and diluted 1:20 using warm CDM. The subcultures were allowed to grow to an OD600 of 0.4, after which they were split into two where one was exposed to 1% DMSO and the other to 10µM XIP. Cultures were further incubated and samples were taken at varying time points (0h, 1h, 2h, 3h, 4h and 5h) after exposure to XIP, gently sonicated, serially diluted and plated on THYE plates for CFU determination. Results were standardized using cellular dry weight. Percentage viability was calculated as the number of viable cells after treatment divided by the total number of cells without peptide, times 100.

3.3.5 Biofilm formation assays

Overnight cultures in THYE were pelleted, washed, resuspended in sterile 1X PBS, and diluted 1:100 using warm CDM. Each suspension was supplemented with either 1% DMSO or 10µM XIP and used to inoculate polystyrene plates. After 24h incubation, the biofilms were dried and strained with 0.1% Safranin Red.

3.3.6 Quantitative real-time PCR (qRT-PCR) analyses

Overnight cultures of UA159 and its derivatives were diluted 20X in fresh THYE or CDM and grown to an OD600 of 0.4-0.5 in the presence or absence of 0.4µM CSP or 10µM XIP, respectively. For growth in CDM, overnight cells were washed, resuspended in 1xPBS prior to inoculation and harvesting. Controls included, THYE without added peptide, as well as CDM with 1%DMSO. RNA isolation, DNAse treatment, cDNA synthesis, qRT-PCR and expression analyses were carried out as previously described (Senadheera et al., 2005). Primers used for qRT-PCR: comR (For: CGTTTAG GAGTGACGCTTGG, Rev: TGTTGGTCGCCATAGGTTG), comS (For: TTTTGATGG GTCTTGACTGG, Rev: TTTATTACTGTGCCGTGTTAGC) and comX (For: ACTGTTTG

TCAAGTCGCGG Rev: TGCTC TCCTGCTACCAAGCG). Expression was normalized to that of 16SrRNA, and statistical analyses were performed on four independent experiments using Student’s t-test (P <0.05).

3.3.7 XIP detection and quantification

Overnight cultures in CDM were diluted 100-fold and grown for 48h at 37°C in

5% CO2 air mixture. Cell-free supernatants were obtained by centrifugation and filter sterilized using a 0.45µm syringe filter. Samples were lyophilized and, once dry, reconstituted in 2 mL of 5% MeOH/H2O (v/v) prior to analysis by HPLC-ESI-MS/MS (Dionex UltiMate 3000 HPLC system with variable UV detection in line to a Bruker amaZon X ion-trap mass spectrometer operating in positive ionization mode with auto MS/MS enabled). Analytical scale analysis was performed on a 250x 4.60 mm Phenomenex Luna 5µ C18(2) 100Å column (Serial no. 516161-20) with a flow rate of 1 ml min-1 and the following program consisting of solvents A (water + 0.1% formic acid) and B (acetonitrile + 0.1% formic acid): 0-2 min, equilibration at 5% B; 2-18 min, linear gradient to 100% B; 18-20 min, constant 100% B, 20-20.5 min, linear decrease to 5% B; 20.5-23 min re-equilibration at 5% B. The identity of XIP in culture supernatants was confirmed by comparison to the retention time and MS/MS fragmentation of sXIP. To quantify XIP levels a directed LC-MS/MS experiment was performed using selected- reaction monitoring (SRM) tandem mass spectrometry. The SRM m/z transition 876.4 → 658.4 was monitored, corresponding to a –SL loss from the GLDWWSL parent ion, generating a GLDWW daughter ion. Resulting peak areas were integrated and final concentrations calculated from a linear calibration curve created using CDM spiked with sXIP and processed in an identical way to CFSs. Peak areas from each sample were standardized to optical densities of 48-h culture samples prior to centrifugation for HPLC-MS analysis. Results were obtained for four independent experiments and statistics were conducted using the Student’s T-test.

3.4 Results

3.4.1 Transformation frequencies (TF) with CSP and/or XIP in CDM and complex media

The initial finding that XIP induces genetic transformation via ComX was reported by Mashburn-Warren et al. (2010) using cells grown in CDM. Recent work by Desai et al., (2012) reported that the induction of comX by XIP was largely inhibited when grown in rich nutrient THB, a medium commonly used to study CSP-induced competence. In accordance with these reports, our TF assays show that XIP is optimally functional in CDM in eliciting transformation and its activity is inhibited when cells are grown in complex medium (i.e. THYE) (Figure 3.1). In contrast, we observed that CSP was largely ineffective at inducing competence in CDM, and that it was optimally functional in complex medium (Figure 3.1). Since CSP and XIP were shown not to function optimally in the same growth medium, we did not obtain significant combinatorial effects in either THYE or CDM (data not shown).

Figure 3.1 Transformation frequency (TF) of S. mutans strains in THYE and CDM growth media. S. mutans UA159 and mutant strains were subcultured to an OD600 of 0.1 in either THYE (a) or CDM (b). Plasmid DNA pDL277 (specr) was added alone or with 0.4µM sCSP or 10µM sXIP for transformation experiments. Results show the mean TF of three independent experiments ± standard error.

3.4.2 Measurement of XIP in the culture supernatants of S. mutans UA159, comR/S, comE and comX knockout mutants

To elucidate the role of known S. mutans competence genes in the regulation of XIP production, its processing and/ or secretion, we used HPLC-ESI-MS/MS to monitor extracellular XIP levels in comR/S, comE and comX-deficient mutants. We were able to successfully identify the presence of XIP in the wild-type supernatant by comparison of the retention time and of the fragmentation patterns to the sXIP standard (Figure 3.2a and b). We were able to detect XIP at concentrations ranging from 95 ng/mL to 750 ng/mL (or 109nM to 857nM), and consistent with the loss of transformability ∆SMcomS, XIP was absent in their cell free supernatants (Figure 3.2c). These results are in accordance with that of Khan et al. (2012) who also reported their inability to detect mature XIP in culture supernatants of the ComS mutant. As expected of a positive regulator of comS expression, ∆SMcomR also displayed highly reduced levels of XIP. Our further quantification of XIP in the ComX and ComE mutants suggested a significant decrease (p<0.05) of this peptide in the ∆SMcomX supernatant, whereas it was significantly increased in the ∆SMcomE supernatant (Figure 3.2c). These results suggested that while ComX positively influenced the production, processing and/or secretion of XIP, the ComDE two component system negatively affected one or more of these processes in S. mutans.

Figure 3.2 Secretion of XIP by S. mutans UA159 and comX-, comR- and comS-mutant strains. (a) Chromatograms resulting from selected-reaction monitoring (SRM) LC-MS experiments used for extraction and quantification of XIP. A transition corresponding to the loss of S and L residues from the XIP peptide parent ion at 876.4m/z, generating a daughter ion at 658.4 m/z was monitored. (b) A comparison of MS2 fragmentation pattern of authentic standard XIP peptide (top panel) and wild-type S. mutans sample (bottom panel). Specific amino acid and water losses are annotated. (c). XIP levels in the supernatant of each strain in stationary phase were quantified using HPLC-ESI-MS/MS. Results shown represent those from at least 3

45 separate experiments, error bars indicate standard deviation. Statistical analyses were performed using a Student’s T-test: *p<0.05.

3.4.3 Effect of XIP on cell viability of S. mutans UA159 grown in CDM

While investigating the effects of sXIP on genetic transformation, we noted that growth of UA159 was drastically impaired by the addition of 10µM XIP in CDM (Figure 3.3a). Since this indicated a likely effect on cell death, we performed cell viability assays to determine whether XIP could act as a death effector of S. mutans. In the presence of 10µM XIP in CDM, we observed only an 18% survival rate relative to the no-peptide control, suggesting that XIP can function as a potent killing peptide under these conditions (Figure 3.3b). XIP was unable to induce killing in ∆SMcomR and was also largely ineffective in ∆SMcomX, demonstrating the importance of both comR and comX in the killing activity of XIP. Furthermore, in ∆SMcomS and ∆SMcomC grown in CDM exposed to XIP, we noted 80% and 89% killing, respectively (Figure 3.3b). In contrast to CDM, XIP was not able to induce killing when S. mutans strains were grown in THYE. To confirm the effect of XIP on cell viability, time course killing analyses were performed, which demonstrated a negative effect of XIP on the CFU counts of healthy cultures at varying time points (Figure 3.3c). Furthermore, S. mutans was not able to form biofilms in the presence of XIP (Figure 3.3d). This drastic effect on biofilm development may be attributed to XIP’s drastic effect on the viability of cells. These results suggest an important role for XIP as a novel killing peptide that can be targeted to kill S. mutans.

Similar to lysis by XIP, CSP-induced cell death was also largely diminished in the absence of comR/S or comX (Figure 3.3), suggesting that the CSP-induced killing pathway previously described requires the presence of comR/S and comX for optimal killing.

Figure 3.3 Effect of CSP and XIP on cell survival of S. mutans. (a). The effect of XIP at varying concentrations (10µM, 1µM, 0.5µM, 0.25µM, 1% DMSO) on the growth of S. mutans

UA159 was examined by monitoring OD600 using the Bioscreen C automated growth reader (b). S. mutans UA159, as well as comR, comS, comX and comC knockout mutant strains were grown in the presence or absence of either XIP (10µM) or CSP (2µM) to OD600~0.8 in THYE (top panel) or CDM (bottom panel). Viable CFUs were counted and results were standardized to the dry weight and presented as the percentage of the CFUs obtained in the absence of the peptide. Results shown here are the average of four independent experiments ± standard error. Statistical analyses were performed using Student’s T-test: *p<0.05, **p<0.01, ***p<0.001. (c). Actively growing cultures of UA159 were exposed to 10µM XIP for 0, 1, 2, 3, 4 and 5h. Following sonication, viable CFUs were counted and results were standardized to the dry weight and presented as the percentage of the CFUs obtained in the absence of the peptide. Results shown here are the average of four independent experiments ± standard error. Statistical analyses were performed using Student’s T-test: *p<0.05, **p<0.01, ***p<0.001. (d). S. mutans biofilms were grown on polystyrene plates overnight in the presence or absence of 10µM XIP,

47 and stained with 0.1% Safranin Red. The image presented here is a representative of four independent experiments.

3.4.4 Regulation of comR/S, and comX expression by XIP, CSP and the ComDE system

Our transformation and viability results as well as that obtained by Mashburn- Warren et al., (2010) and Desai et al., (2012), strongly suggest that the ComCDE system may regulate comX transcription through ComRS, although this was not directly tested. Hence, we examined comR/S and comX transcription in UA159, ∆SMcomD and ∆SMcomE strains grown with and without CSP or XIP. Due to the poor activity of CSP in CDM and no activity of XIP in THYE, experiments with CSP were performed in THYE, whereas those with XIP were conducted from cells grown in CDM. Supporting a hierarchal position of the ComCDE system upstream of ComRS, we observed that addition of CSP increased comS and comX expression by 73.9-fold and 2.3-fold, respectively (Figure 3.4a). In THYE without added CSP, comR/S, and comX expression was not significantly affected by loss of comD/E relative to wild-type (Figure 3.5a). However, with CSP, expression of comS was significantly decreased over 100-fold in both mutants (p<0.001), relative to wild-type (Figure 3.5b). Addition of CSP also decreased comX expression by nearly 30-fold in ∆SMcomD and ∆SMcomE strains, respectively, compared with the parent (Figure 3.5b). These results suggested that in complex medium, comS expression can be modulated by adding CSP, and that comS induction by the CSP is ComDE-dependent.

Figure 3.4 Gene expression of comR, comS and comX in response to CSP and XIP. Real- time analysis of gene expression of comR, comS and comX were preformed using mid- logarithmic UA159 cells grown in THYE (a) and CDM (b) in the presence or absence of either 0.4µM CSP (a) or 10µM XIP (b). Gene expression was normalized to UA159 with no peptide added. Results shown here are the average of at least three independent experiments ± standard error. Statistical analyses were performed using Student’s T-test: ***p<0.001

In wild type, addition of sXIP increased expression of comX and comS by 83-fold and 141-fold, respectively (Figure 3.5b), thus confirming the autoregulatory loop described by Mashburn-Warren et al., 2010. In ∆SMcomD and ∆SMcomE grown in CDM, comS and comX genes were upregulated almost 3-fold without added peptide, likely suggesting that ComDE may repress their expression in CDM medium (Figure 3.5c). This finding was also supported by the high levels of XIP detected in the ∆SMcomE culture supernatant. Further, upon addition of sXIP to ∆SMcomD and ∆SMcomE mutants no change in comR and comS expression was observed (Figure 3.5d), suggesting that the ComDE system does not affect XIP signaling, once the ComRS system is activated.

Figure 3.5 Differential regulation of comRS in THYE and CDM media grown in the presence or absence of CSP and XIP, respectively. Real-time analysis of comR-, comS- and comX-specific expression in comD- and comE-knockout mutants grown in THYE (a) and CDM (c) media, as well as in the presence of either 0.4µM CSP (b) or 10µM XIP (d). Gene expression was normalized with 16SrRNA and fold-expression was calculated relative to that of UA159 held at a user-defined value of 1.0. Results shown are the mean expression values of at least three independent experiments ± standard error. Statistical analyses were performed using Student’s T-test: *p<0.05, **p<0.01, ***p<0.001.

3.5 Discussion

Competence has been observed in a number of bacteria to occur in conjunction with lysis of a subpopulation of cells (Claverys et al., 2007; Lemme et al., 2011; Perry et al., 2009; Steinmoen et al., 2002). The lysed subpopulation is thought to contribute to the genetic pool used for DNA uptake by the competent cells. Herein, we have demonstrated a role for the XIP competence peptide as potent modulator of cell death in S. mutans. Our viability assays show XIP can kill nearly 82% of the population when supplied at a concentration of 10µM. To our knowledge, this is the first report that demonstrates a function for XIP as an effector of cell death.

We further report that XIP-mediated killing works via the ComR/S system and ComX, which positions the ComR/S and ComX in a more centralized position in the killing pathway of S. mutans. Although previous reports have attributed CSP-induced lysis to an imbalance between the ComE-regulated mutacin V and its immunity protein ImmB (Dufour et al., 2011; Lemme et al., 2011; Perry et al., 2009), here we argue that competence-associated cell death in S. mutans, is instead, largely due to activity downstream of ComX. This is also supported by the fact that nlmC (synonyms: cipB, bsmA) encoding mutacin V also modulates comX activity, which in turn, may contribute to its killing activity (Dufour et al., 2011). We are currently examining genes downstream of ComX stimulated by XIP that may function as killing effectors using global transcriptome analysis.

Although the killing activity of CSP harbors specificity towards its parent strain (Qi et al., 2005), the spectrum of activity of XIP has yet to be determined. XIP contains a double-tryptophan (WW) motif conserved among short hydrophobic peptides of the pyogenic and bovis groups of Streptococci, located within a conserved genomic context (Mashburn-Warren et al., 2010). Similar peptides specific for Streptococcus agalactiae, Streptococcus porcinus, and Streptococcus parauberis have been shown to bear no effect on competence or growth of S. mutans, suggesting that these peptides may be specific to their parental strain (Desai et al., 2012). XIP therefore may be exploited for targeted killing of S. mutans.

Our transformation and cell viability results with CSP and/or XIP in both THYE and CDM media showed that these peptides do not function optimally under the same conditions. Our transformation results are in agreement with Desai et al., (2012) who reported that titration of THB into UA159 cultures in CDM inhibited XIP-induced transformability. While they demonstrated some level of activity of XIP in 100% THB, our results showed complete inhibition of XIP in THYE. It is likely that the yeast extract in THYE is largely responsible for the inhibition observed.

Despite the observation that XIP is inactive in THYE-grown cells, the comS gene is still required for transformation despite growth in a complex medium. Hence, it is possible that the ComS peptide may also function intracellularly without its export and subsequent import into the cell. We have also taken into consideration that conditions tested in complex medium may not be optimal for the expression of the XIP exporter, which can likely result in the accumulation of ComS inside the cell, making it vulnerable to intracellular cleavage.

Our expression analysis combined with LC-MS/MS in CDM demonstrates a negative-regulatory role for the ComDE system in XIP production. Kreth et al. (2007) reported that ComDE repressed comC expression prior to CSP stimulation. It is possible that ComDE may prevent premature expression of comS, thereby delaying competence induction in CDM to the latter stages of growth. As observed by Desai et al., (2012), competence in CDM is first observed in mid-logarithmic cells of S. mutans and continues well into the stationary phase.

We further observe that the amount of XIP was significantly reduced in ∆SMcomX, suggesting a ComX-mediated positive feedback mechanism for XIP synthesis. Putative ComX binding sites were located within the comR gene, upstream of comS, suggesting that ComX may directly regulate comS expression (Figure 3.6a). This positive autoregulation of XIP production may contribute to the persistence of the competent state in CDM. Based on previous works and our findings presented here, we propose a growth condition-dependent model for genetic competence in S. mutans (Figure 3.6b).

(a)

(b)

Figure 3.6 Model for competence and cell killing pathways of S. mutans in THYE versus CDM. (a). Two putative ComX binding sites within the comR gene, located 102bp apart are underlined and shown in bold font (b). In complex medium (THYE), CSP is sensed by the ComDE system, which stimulates the expression of comS, leading to an intracellular accumulation of the ComS peptide or its cleavage products, which in turn, can activate ComR to induce comX expression. In CDM, the ComDE system represses comS expression. This repression is released by an unknown mechanism to upregulate comS, whose product is then secreted, processed and imported by the Opp/Ami transporter to activate comX transcription, as first described by Mashburn-Warren et al (2010). ComX positively regulates XIP production, which may contribute to the persistence of the competent state in CDM, as described by Desai et al (2012).

3.6 Acknowledgements

We thank Kirsten Krastel for technical assistance. We are thankful to Dr. Donald Morrison for his review of our manuscript and helpful suggestions provided along with Dr. Lauren Mashburn-Warren and Dr. Mike Federle. D.G.C. is a recipient of the NIH grant R01DE013230-03 and CIHR-MT15431.

Chapter 4: The effects of the RcrRPQ operon on comS expression and XIP levels

Iwona B. Wenderska, Nikola Lukenda, Dennis Liu, Nathan Magarvey, Dilani B. Senadheera, Dennis G. Cvitkovitch

Author contributions: IBW, DBS and DGC conceived and designed the study. IBW wrote the manuscript. IBW performed all experiments with the exception of LCMS/MS quantification of XIP in cell extracts, which was performed by NL and DL.

4.1 Abstract

In the cariogenic pathogen Streptococcus mutans, competence development is regulated by the ComRS signaling system, comprised of the ComR transcriptional regulator and the ComS pre-peptide to the mature ComX inducing peptide, XIP. ComR in conjunction with XIP regulate the expression of an alternate sigma factor ComX, which in turn regulates the expression of genes involved in DNA uptake and recombination. Genetic competence of S. mutans is also co-regulated with the (p)ppGpp stress alarmones, expressed in response to amino-acid starvation. Both DNA transformation and (p)ppGpp synthesis are regulated by the RcrRPQ operon consisting of a transcriptional regulator and two ABC transporters, respectively. Although both ComRS and RcrRPQ are crucial for DNA transformation, as deletions in any of these genes result in drastic competence phenotypes, their interaction has not yet been investigated. Herein we examined the role for RcrRPQ in the regulation of the main competence signaling system ComRS. We demonstrated that the phenotypes associated with various gene deletions in the rcrRPQ operon can be correlated with comS expression and concentrations of endogenous XIP. We further provide initial evidence to support the role of the RcrQ ABC transporter in the export of the ComS pre-peptide.

4.2 Introduction

Genetic competence is a physiological state that enables bacteria to bind and take up extracellular DNA (eDNA) by the process known as transformation. Many competent species rely on the incoming DNA to increase genetic diversity, which in turn allows for increased population survival under stressful conditions (Claverys et al., 2006; Didelot and Maiden, 2010). Streptococcus mutans, one of the main causative agents of dental caries, is naturally competent, and regulates the state of competence by two main signaling peptides: ComX inducing peptide (XIP) and Competence stimulating peptide (CSP) (Fontaine et al., 2015). Both peptides control the expression of the alternate sigma factor ComX, albeit under different growth conditions (Desai et al., 2012; Wenderska et al., 2012). ComX in turn regulates genes involved in DNA uptake and recombination, and as a result is essential for genetic transformation. ComX induction also results in lysis of a fraction of the population (Wenderska et al., 2012), contributing to the pool of eDNA available for competent cells. All signaling systems and transcriptional regulators affecting genetic competence converge to directly or indirectly regulate ComX (Fontaine et al., 2015).

The XIP peptide is part of the core competence regulatory system ComRS, comprised of the ComR transcriptional regulator and the ComS prepeptide to XIP. ComS is secreted out of the cell and processed by an unknown mechanism. Mature XIP is subsequently imported back into the cell by the Opp/Ami oligopeptide transport system to interact with ComR (Fontaine et al., 2013; Mashburn-Warren et al., 2010). Being dependent on the opp oligopeptide transport system for import, XIP is susceptible to competition by nonspecific polypeptides, and as a result has been shown to be active only under chemically defined conditions devoid of polypeptides (Desai et al., 2012; Wenderska et al., 2012). XIP-activated ComR binds directly to the comX promoter to increase comX transcription, and to the comS promoter creating a positive feedback loop (Fontaine et al., 2013; Mashburn-Warren et al., 2010). ComRS, like ComX, are essential for competence development, and for competence-associated lysis in a

58 subpopulation of cells (Mashburn-Warren et al., 2010; Wenderska et al., 2012).

The CSP peptide was initially identified for its role in genetic transformation, however it is now recognized mainly as a regulator of bacteriocins, antimicrobial peptides used by bacteria to gain a competitive advantage over closely related species (Li, Lau, et al., 2001; Perry et al., 2009; Reck et al., 2015; van der Ploeg, 2005). CSP signaling is transduced by the ComDE two-component signaling system, comprised of a histidine kinase and a response regulator, respectively (Li, Lau, et al., 2001). Activated ComE directly regulates a number of bacteriocin genes including nlmC, and has an indirect effect on the expression of comX (Hung et al., 2011; Perry et al., 2009; Reck et al., 2015). CSP signaling has been characterized under nutrient rich conditions, under which the activity of the exogenous XIP is inhibited (likely by nonspecific polypeptides in the medium competing for import by opp/ami) (Desai et al., 2012; Wenderska et al., 2012). The comRS genes, however, are still essential for CSP- induced competence in rich media (Wenderska et al., 2012). It has recently been proposed that the CSP-induced NlmC bacteriocin increases comX expression indirectly by increasing the permeability of the cell to the main competence stimulating peptide XIP (Reck et al., 2015).

Genetic competence in S. mutans has been shown to be co-regulated with (p)ppGpp synthesis. Collectively referred to as (p)ppGpp, this group of signaling molecules are responsible for global transcriptional changes that reduce chromosomal replication and the growth of the population (Potrykus and Cashel, 2008). This coordination between genome replication and natural transformation may ensure correct chromosome rearrangements following homologous recombination of transformed DNA. In S. mutans, DNA transformation and the expression of the (p)ppGpp synthases are regulated by the RcrR transcriptional regulator of the MarR-family of regulators (Seaton et al., 2011). RcrR is part of an operon containing two predicted ABC exporters: rcrP and rcrQ. The loss of individual genes in the rcrRPQ operon results in dramatic changes to the competence phenotype under nutrient rich conditions (Seaton et al., 2015;

Seaton et al., 2011). A non-polar (NP) deletion of rcrR results in a competence negative phenotype, while its polar (P) counterpart results in increased transformation (Seaton et al., 2011). Since RcrR functions as a negative regulator of the rcrRPQ operon, the drastic differences between the two strains were attributed to the differences in rcrP and rcrQ expression. In the ∆RcrR-NP mutant, expression of rcrPQ increased 1000-fold, while in the ∆RcrR-P mutant, the two transporters were expressed at wild-type levels. Seaton et al., further demonstrated that a deletion of rcrPQ resulted in increased natural transformation, and decreased transformation in response to exogenously added CSP.

Recently, two peptides (pep1 and pep2) involved in competence inhibition have been identified at the 3’ end of rcrQ, as part of the rcrRPQ operon (Ahn et al., 2014). Overexpression of pep1 and pep2 resulted in decreased levels of the ComX protein. As a result, it was proposed that these peptides inhibit transformation by either interfering with ComR-XIP ability to induce comX expression, or by increasing ComX degradation by the ClpPCMecA complex, which has been previously demonstrated to regulate ComX stability (Tian et al., 2013). The activity of the peptides also appears to be independent of the phenotypes associated with the rcrPQ transporters. Specifically, it was demonstrated that the phenotypes associated with the rcrPQ genes remained unchanged regardless of the presence or absence of the downstream peptides, and therefore concluded that pep1 and pep2 are not substrates for export by RcrPQ (Ahn et al., 2014).

The role of RcrRPQ in genetic competence has thus far been investigated under rich nutrient conditions not optimal for XIP signaling. Due to the strong competence phenotypes associated with the rcrRPQ operon, we investigated the direct effects of this operon on the core competence signaling system ComRS. Herein, we examine the role for the RcrRPQ operon in comS expression, secretion and the XIP-associated phenotypes. We argue that the phenotypes associated with various gene deletions in the rcrRPQ operon can be explained by

60 expression levels of the comS transcript and concentrations of endogenous XIP. We further investigated each of the ABC transporters separately for their role in genetic competence and suggest that due to the opposite phenotypes observed, RcrP and RcrQ likely secrete different substrates. We further propose that the RcrQ ABC transporter may play a role in the export of the ComS pre-peptide.

4.3 Materials and methods

4.3.1 Strains and growth conditions

S. mutans UA159 and its deletion strains rcrR (ΔSMrcrR), rcrP (ΔSMrcrP), rcrQ (ΔSMrcrQ), oppD (ΔSMoppD), rcrP/oppD (ΔSMrcrP/oppD), rcrQ/oppD (ΔSMrcrQ/oppD) were used in this study. S. mutans strains were grown at 37oC with 5% CO2 in either Todd-Hewitt Broth (Becton Dickinson, MD) containing 0.3% yeast extract (THYE) or chemically defined medium (CDM). Erythromycin and kanamycin were used as needed at concentrations of 10 µg/mL and 1 mg/mL, respectively. Synthetic XIP peptide was synthesized using F-MOC chemistry (Advanced Protein Technology Centre, Hospital for Sick Kids, Toronto, ON, Canada). Stock concentrations of 1 mM XIP were prepared in DMSO.

4.3.2 Transformation assays

Overnight cells grown in THYE were pelleted, washed, and resuspended in fresh CDM. The resuspended culture was diluted 1:20 using pre-warmed CDM and grown to an OD600nm ~ 0.6. Next 1 µg/mL of the donor plasmid DNA (pDL277; specR) was added to 1 mL aliquots of the culture in the presence of 10 µM XIP or 1% DMSO vehicle control, and samples were incubated for 90 min. After incubation, cultures were sonicated to disrupt any streptococcal chains, serially diluted, and plated on THYE plates with or without antibiotic.

Transformation frequency (TF) was calculated as transformant colony forming units (CFUs) divided by the total number of viable CFUs, times 100.

4.3.3 Quantification of ß-glucuronidase released

Cell lysis of S. mutans UA159 was determined by harvesting the supernatants of cultures expressing the gusA reporter gene cloned into pIB187 under a constitutive promoter (Syed et al., 2011). Briefly, supernatants were mixed in equal parts with GUS buffer, and incubated at 37oC for 120 min. 3.5mM

Na2CO3 was added to stop the reaction and absorbance was read at 420nm.

GUS activity was expressed as (1000 x A420)/(time [min] x OD600) in Miller Units.

4.3.4 Quantitative real-time PCR (qRT-PCR) analyses

Overnight cultures of wild-type UA159 and deletion strains in THYE were washed and resuspended in CDM. The cultures were then further diluted 20x in fresh CDM and grown to an OD600nm of 0.4-0.5 or 0.8-0.9. RNA isolation, DNAse treatment, cDNA synthesis, qRT-PCR, and expression analyses were carried out as previously described (Wenderska et al., 2012). Primers used for qRT-PCR were as follows: comR (For: CGTTTAGGAGTGACGCTTGG, Rev: TGTTGGTCGCCATAGGTTG), comS (For: TTTTGATGGGTCTTGACTGG, Rev: TTTATTACTGTGCCGTGTTAGC) and comX (For: ACTGTTTGTCAAGTCGCGG Rev: TGCTCTCCTGCTACCAAGCG), rcrQ (For: CGATTGTTGCCTTTATGACT TACG Rev: GGTTTTCGTCTTCTATTTCTTCCTC). Expression was normalized to that of the 16S rRNA gene, and statistical analyses were performed on four independent experiments using Student’s t-test (P < 0.05).

4.3.5 XIP quantification

Overnight cultures of wild-type UA159 and deletion strains were diluted o 100-fold in CDM and grown for 48 h at 37 C in 5% CO2 air mixture. Cell-free supernatants were obtained by centrifugation and filter sterilized using a 0.45 µm-syringe filter. Samples were lyophilized, and once dry, reconstituted in 2 mL of 5% MeOH/H2O (v/v) prior to analysis by HPLC-MS/MS as described previously (Wenderska et al., 2012).

4.3.6 RcrR purification and electrophoretic mobility shift assays (EMSAs)

A C-terminally tagged RcrR protein was obtained by cloning the rcrR gene into the pET32 vector. Protein expression was induced in the E. coli BL21(D3) Rosetta strain by addition of 1 mM isopropyl-β-D-thiogalactopyranoside (IPTG) and incubation for 2 hrs at 30oC. The protein was purified using the HisTALON cobalt-affinity column as per manufacturer instructions (Clontech). The purified protein was tested in electromobility shift assays with 20 ng of the pComS, and the pfruA as a negative control. Various concentrations of the purified RcrR were incubated with 20 ng DNA in binding buffer (10 mm HEPES (pH 7.9), 50 mm KCl,

5 mm MgCl2, 1 mm EDTA, 5 mm dithiothreitol and 10% glycerol) for 40 min. 2 µL of 10x Fermentas loading dye was added to each 20 µL reactions immediately prior to loading on a 4% native polyacrylamide gel. The DNA complexes were separated for 2.5 h at 70V on ice in 1xTAE buffer. The gel was stained with

SYBR Green for 20 min at room temperature, washed twice with ddH2O, and visualized using ultraviolet light.

4.4 Results

4.4.1 RcrR regulates comS expression and extracellular XIP levels.

To determine the relationship between RcrR and ComRS we examined the effect of the rcrR deletion on comS gene expression, extracellular XIP amounts and genetic transformation in response to exogenously added XIP. Transformation assays were performed on cells in late-log phase, the time of natural transformation of S. mutans UA159 cultured in CDM (Desai et al., 2012). This was done to clearly demonstrate the effect of various gene deletions on natural transformation as compared to wild type. Addition of increasing amounts of XIP partially restored the previously characterized competence deficient phenotype of the rcrR deletion strain (Figure 4.1a). To reinforce our observations, addition of XIP also induced competence-associated lysis of ∆SMrcrR. By measuring the release of a constitutively expressed cytoplasmic enzyme we demonstrated that XIP increased cell lysis in ∆SMrcrR in a concentration dependent manner (Figure 4.1b). These results suggest that exogenously added XIP can in part complement competence-associated phenotypes of ∆SMrcrR, and suggests that limiting levels of endogenous comS may in part contribute to the competence negative phenotype of this strain. To determine whether comS levels were altered as a result of the rcrR deletion, we examined the expression of comS, and its regulatory target comX, in ∆SMrcrR by qRT-PCR. Gene expression was monitored prior to (OD600 ~ 0.4) and during competence (OD600 ~ 0.8) of S. mutans UA159 cultured in CDM. The rcrQ gene, previously shown to be negatively regulated by RcrR was used as a control (Seaton et al., 2011) (Figure 4.1c). qRT-PCR analyses revealed a drastic decrease in comS transcript levels in ∆SMrcrR during competence (Figure 4.1c).

We observed that during competence (OD600 0.8) comS expression was reduced 200-fold, and subsequently resulted in a 1600-fold reduction in comX expression (Figure 4.1c). Furthermore, XIP quantification revealed a significant decrease in XIP amounts in the supernatant of this strain (Figure 4.1d). These results

64 suggested that the phenotypes associated with the rcrR deletion may in part be explained by its role in positive regulation of comS expression.

To determine whether the regulation of comS expression by RcrR was direct, purified RcrR protein was incubated with the pComS promoter, and the DNA complex analyzed using EMSAs. The prcrRPQ promoter was used as a positive control since RcrR has been shown to autoregulate its own operon (Seaton et al., 2015; Seaton et al., 2011). The pfruA promoter was used as a negative control since its activity is not under the direct control of RcrR (Seaton et al., 2015). Specific binding of RcrR to the promoter of pComS was not observed under the conditions tested (Figure 4.2). It is therefore likely that RcrR regulates comS expression in an indirect manner.

a c

0.1

0.01 1000 *** OD 0.4 100 *** OD 0.8 0.001 10

0.0001 1 0.1 *** % Transformation frequency Transformation % 0.00001 UA159 to relative 0.01 *** 0.001 0.000001 expression gene Normalized rcrQ rcrQ comScomRcomX comScomRcomX M XIP M XIP M XIP M XIP µ µ µ µ

UA159 + 1 SMrcrR + 1 UA159 + 1% DMSO UA159 + 10SMrcrR + 1%Δ DMSO SMrcrR + 10 Δ Δ

b d

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100 40

Miller Units (MU) *** 20 50 % XIP production XIP % relative to UA159 to relative

0 0

M XIP M XIP µ µ UA159 SMrcrR Δ

UA159 + 1 SMrcrR + 1 UA159 + 1% DMSOUA159 + 10 µMSMrcrR XIP + Δ1% DMSOSMrcrR + 10 µM XIP Δ Δ

Figure 4.1 The effect of ∆SMrcrR on XIP-associated phenotypes, comS expression and extracellular levels of XIP. (a). UA159 and mutant strains were subcultured to an r OD600 of 0.6 in CDM. Plasmid DNA pDL277 (spec ) was added alone or with XIP at varying concentrations. Results showed the mean %TF of three independent experiments ± standard error. (b). XIP-induced cell lysis, demonstrated by the release of the cytoplasmic β-glucuronidase (GUS) enzyme into the extracellular medium of UA159 pIB187 or ΔSMrcrR pIB187 grown in the presence of varying concentrations of XIP. (c). qRT-PCR analysis of comR/S, comX and rcrQ expression was performed in UA159 and ΔSMrcrR strains grown in CDM to OD600 0.4 or OD600 0.8. ***p<0.001 (d). XIP levels in the supernatant of UA159 and ∆SMrcrR were quantified using HPLC-ESI-MS/MS. Results shown represent those from at least three separate experiments, and error bars

66 indicate standard deviation. Statistical analyses were performed using Student’s t-test: ***p< 0.001.

prcrRPQ pcomS pfruA RcrR (pM) 0 167 333 500 0 167 333 500 0 167 333 500

Figure 4.2 Electrophoretic mobility shift assays of RcrR protein binding to promoters of rcrRPQ, comS and fruA. EMSAs were conducted using purified RcrR. As indicated, the purified protein was added to 379 bp pRcrRPQ, 216 bp pComS or 231 bp pfruA DNA fragments in increasing protein concentrations. DNA/protein complexes were separated by 4% native polyacrylamide gel and stained with SYBR green.

4.4.2 Increased competence of ∆SMRcrP is the result of an increase in endogenous levels of XIP

To determine whether the increased transformability of ∆SMrcrP can also be explained by increased comS expression and XIP levels, we examined gene

67 expression and peptide amounts in this strain as compared to wild type. The ∆SMrcrP strain is highly transformable and cannot be further induced by XIP (Figure 4.3a). The increased competence phenotype correlates with early comS expression. Specifically, qRT-PCR revealed a 60-fold increase in comS expression prior to competence development in ∆SMrcrP, and wild-type levels during competence, when comS is highly expressed in both strains (Figure 4.3b). The early onset of comS induction in ∆SMrcrP results in increased XIP levels in the supernatant of this strain as determined by LCMS/MS (Figure 4.3c), and resulting 14-fold increase in comX expression.

To further demonstrate the importance of comS for the highly competent phenotype of ∆SMrcrP we deleted the comS gene in this strain. A deletion of the comS gene in the highly transformable ∆SMrcrP strain abolishes transformation, suggesting that the high levels of comS are solely responsible for the increased competence in this strain (Figure 4.3a). Furthermore, the competence negative phenotype of the ∆SMrcrP/∆SMcomS double deletion strain can in part be complemented by exogenously added XIP. These results suggest that perhaps ∆SMrcrP produces XIP at saturating levels and therefore cannot be further induced by XIP, however when endogenous comS levels are reduced, as a result of a comS deletion, exogenous XIP is able to induce competence despite the ∆SMrcrP background.

a b

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1 / P + 50 Mrcr 0 S UA159 + 1 SMrcrP + 9 SMcomS + 1 SMcomS 15 / A U SMrcrP

SMrcrP

Figure 4.3 The effect of ∆SMrcrP on XIP-associated phenotypes, comS expression and extracellular levels of XIP. (a). UA159 and mutant strains were subcultured to an r OD600 of 0.6 in CDM. Plasmid DNA pDL277 (spec ) was added alone or with 10 µM XIP for transformation experiments. Results show the mean %TF of three independent experiments ± standard error. (b). qRT-PCR analysis of comR/S, comX and rcrQ expression was performed in UA159 and ΔSMrcrP strains grown in CDM to OD600 0.4 or

OD600 0.8. ***p<0.001 (c). XIP levels in the supernatant of wild-type and deletion strain were quantified using HPLC-ESI-MS/MS. Results shown represent those from at least three separate experiments, and error bars indicate standard deviation. Statistical analyses were performed using Student’s t-test: ***p< 0.001.

4.4.3 The lack of the RcrQ transporter affects the amount of secreted XIP

Aside from increased comS levels, the rcrQ exporter was also highly upregulated in the highly transformable ∆SMrcrP strain (Figure 4.3a) and lead us to hypothesize that rcrQ may be involved in ComS transport. A deletion of the rcrQ gene resulted in abolished transformation that could not be complemented with exogenously added XIP (Figure 4.4a). To determine whether this loss in transformability was a result of low levels of the ComS peptide we examined its expression in this strain using qRT-PCR. Despite the low transformation phenotype, comS levels were increased 39-fold at mid-logarithmic (OD600 0.4-

0.5) and 5-fold at late-logarithmic (OD600 08-0.9) growth phases (Figure 4.4b). Despite the increase in comS transcription, ∆SMrcrQ supernatant contained five times less XIP than wild-type (Figure 4.4c), and subsequently resulted in decreased comX levels (Figure 4.4b). The low levels of XIP in the supernatant of ∆SMrcrQ, despite the high levels of comS transcript in this strain, suggested a potential role for RcrQ in ComS export.

a b OD 0.4 OD 0.8 on i ss

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UA15 o

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0 0 t p

e v P i I t 50 X a l

e *** % UA159 + 1 r SMrcrQ + 1

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UA15 SMrcr

Figure 4.4 The effect of ∆SMrcrQ on XIP-associated phenotypes, comS expression and extracellular levels of XIP. (a). UA159 and ∆SMrcrQ were subcultured to an OD600 of 0.6 in CDM. Plasmid DNA pDL277 (specr) was added alone or with 10 µM XIP for transformation experiments. Results show the mean %TF of three independent experiments ± standard error. (b). qRT-PCR analysis of comR/S and comX expression was performed in UA159 and ΔSMrcrQ strains grown in CDM to OD600 ~ 0.4 or OD600 ~ 0.8. ***p<0.001 (c). XIP levels in the supernatants of UA159 and ∆SMrcrQ were quantified using HPLC-ESI-MS/MS. Results shown represent those from at least three separate experiments, and error bars indicate standard deviation. Statistical analyses were performed using Student’s t-test: ***p< 0.001.

4.4.4 High expression of the full-length ComS peptide inhibits exogenous XIP activity

As described above and depicted in Figure 4.4a, the ∆SMrcrQ mutation results in competence deficiency that cannot be complemented by exogenous XIP. Assuming that ComS export is required for processing and XIP activity, a negative competence phenotype like the one observed in ∆SMrcrQ is expected of the XIP exporter. However a mutation in the exporter of XIP should not affect the import of exogenously supplied XIP and therefore its activity. The observation that exogenously added XIP cannot complement the competence negative phenotype of ∆SMrcrQ was therefore puzzling to us. The OppD/Ami oligopeptide transporter is expressed at WT levels in this strain (Figure 4.5), suggesting that XIP import is not likely affected. Interestingly, a deletion of the comS gene in ∆SMrcrQ restores sensitivity of the competence negative strain to exogenous XIP (Figure 4.6). This suggests that the high level of comS expression in the ∆SMrcrQ strain may inhibit XIP activity, perhaps by competitively inhibiting the XIP-ComR interaction.

Aside from increased comS transcription in the ∆SMrcrQ strain, an increase was also observed in the previously-characterized competence- inhibitory peptides pep1 and pep2 (Figure 4.7). Since these peptides were suggested to inhibit competence by interfering with ComR-XIP activity, it is possible that the lack of sensitivity of the ∆SMrcrQ strain to exogenously added XIP is due to increased levels in pep1 and pep2. To determine whether pep1 and pep2 are responsible for the lack of sensitivity of the ∆SMrcrQ mutant to XIP, we examined the expression of pep1 and pep2 in the ∆SMrcrQ mutant, insensitive to exogenous XIP, and the ∆SMrcrQ/comS mutant, with restored XIP sensitivity. Expression of the two peptides was unaltered in the two strains suggesting that the pep1 and pep2 peptides are not responsible for the lack of sensitivity of ∆SMrcrQ to exogenous XIP.

Figure 4.5 Expression of oppD, pep1 and pep2 in ∆SMrcrQ compared to WT UA159. qRT-PCR analysis of oppD, pep1 and pep2 expression was performed in

UA159 and ΔSMrcrQ strains grown in CDM to OD600 0.4 or OD600 0.8. ***p<0.001

Figure 4.6 Transformation frequency in UA159, ∆SMcomS, ∆SMrcrQ, and ∆SMcomS/rcrQ double deletion strain in the presence or absence of XIP. UA159,

∆SMcomS, ∆SMrcrQ and ∆SMcomS/rcrQ were subcultured to an OD600 of 0.6 in CDM. Plasmid DNA pDL277 (specr) was added alone or with 10 µM XIP for transformation experiments. Results show the mean %TF of three independent experiments ± standard error.

100 OD 0.4 OD 0.8

0.1 pep1 gene expression relative to UA159

SMrcrQ SMcomS Δ Δ SMcomS/rcrQ Δ

100 OD 0.4 OD 0.8

0.1 pep2 gene expression relative to UA159

SMrcrQ SMcomS Δ Δ SMcomS/rcrQ Δ

Figure 4.7 Transcription of pep1 and pep2 competence inhibitory peptides in ∆SMcomS, ∆SMrcrQ and ∆SMcomS/rcrQ as compared to UA159. RT-PCR analysis of pep1 (top panel) and pep2 (bottom panel) expression was performed in UA159,

ΔSMcomS, ΔSMrcrQ and ΔSMcomS/rcrQ strains grown in CDM to OD600 0.4 or OD600 0.8. ***p<0.001

4.5 Discussion

Horizontal gene transfer via DNA transformation is an important evolutionary mechanism among many bacteria to increase genetic diversity and population survival under stressful conditions (Claverys et al., 2006; Didelot and Maiden, 2010). The state of competence for transformation occurs at a physiological cost due to the expression of protein machineries involved with DNA uptake and recombination, and as a result is tightly regulated. In genetically competent bacteria, DNA transformation is often controlled by a core competence signaling network of genes, including the ComRS and ComCDE systems present in many streptococcal species (Fontaine et al., 2015). In some bacteria competence is also co-regulated with mechanisms that control genome replication and bacterial growth to ensure proper chromosome rearrangements after homologous recombination of transformed DNA. In S. mutans, DNA transformation and the (p)ppGpp signaling alarmones that control replication and cell growth are regulated by the same operon: rcrRPQ, containing a transcriptional regulator and two ABC transporters, respectively (Seaton et al., 2011). RcrRPQ has only been investigated under rich nutrient conditions in relation to the ComCDE system, which regulates the transformability of S. mutans in an indirect manner. Herein we demonstrate the role for RcrRPQ in the regulation of the main competence signaling system ComRS.

Our data suggested that the RcrR transcriptional regulator contributed to genetic competence in S. mutans in part by positively regulating comS expression. The ΔSMrcrR strain expresses low levels of the comS transcript (Figure 4.1c) which in turn results in low amounts of extracellular XIP (Figure 4.1d) and low expression of the downstream sigma factor comX (Figure 4.1c) that ultimately leads to a 1000-fold reduction in DNA uptake (Figure 4.1a). Exogenous XIP could not fully complement competence in the ΔSMrcrR strain, likely because the previously identified inhibitory peptides pep1 and pep2, expressed as part of the rcrRPQ operon and under the control of RcrR, also play a role in the phenotype of ΔSMrcrR (Ahn et al., 2014). Since we did not observe

75 binding of the RcrR protein to the comS promoter (Figure 4.2), RcrR likely regulates comS expression in an indirect manner. RcrR has previously been demonstrated to bind the promoter of comX, albeit with lower affinity than the promoter of its own operon (Seaton et al., 2015). Since we have previously observed that a ∆SMcomX deletion strain produced lower amounts of extracellular XIP possibly through an uncharacterized positive feedback mechanism (Wenderska et al., 2012), it is possible that RcrR may exert its effects on comS in an indirect manner via its regulation of comX.

Although previous reports addressing the role of the RcrP/Q transporters in genetic competence have examined them as a unit (Ahn et al., 2014; Seaton et al., 2011), our data indicated that they each have their own, distinct, roles in competence. By deleting each putative exporter separately we demonstrate that they have opposite effects on DNA transformation, and therefore likely export different substrates. The ΔSMrcrP strain is 100-fold more competent than wild- type (Figure 4.3a). A deletion of the comS gene in the highly competent ΔSMrcrP strain resulted in abolished transformation. The increased transformation frequency of the ΔSMrcrP strain is therefore dependent on the presence of comS. In agreement with our transformation results, expression analysis by qRT-PCR revealed that comS is significantly induced in ΔSMrcrP prior to competence development (OD600 ~ 0.4) (Figure 4.3b), allowing for the accumulation of high levels of XIP in the supernatant of this strain (Figure 4.3c) and the corresponding increase in DNA uptake. RcrP may therefore play a role in negative regulation of competence, perhaps by excreting a competence factor that positively regulates ComRS signaling.

Our results suggested that the second putative exporter RcrQ is essential for DNA transformation in S. mutans (Figure 4.4a). The competence negative ΔSMrcrQ strain displayed significantly reduced levels of extracellular XIP as measured by LCMS/MS (Figure 4.4c), which resulted in reduced expression of the downstream comX competence factor (Figure 4.4b). The expression of comS however was significantly increased, both prior to (OD600~0.4) and during

transformation (OD600~0.8) (Figure 4.4b). The increased levels of comS transcript and low amounts of XIP in the supernatant of ΔSMrcrQ suggested that the ComS pre-peptide may be expressed but not secreted. The RcrQ exporter may therefore play a role in the secretion of the ComS pre-peptide. Since we did not measure intracellular ComS peptide levels, we cannot exclude the possibility that the ΔSMrcrQ strain may be impaired in comS translation or the stability of the ComS pre-peptide. However, since the ΔSMrcrQ can only be complemented with exogenously added XIP in the absence of the comS gene (Figure 4.6), the high levels of comS transcripts may not only be expressed, but may also play a role to inhibit exogenous XIP activity.

Due to the essential role of RcrQ in genetic competence we would expect the rcrPQ double deletion strain to reflect the phenotype of ΔSMrcrQ alone. However, previous reports on the rcrPQ double deletion strain yielded phenotypes similar to our highly competent ΔSMrcrP strain (Seaton et al., 2011). This research was carried out under rich nutrient conditions optimal for CSP- induced competence. Since CSP signaling induces competence indirectly likely by increasing the permeability of the cell to XIP (Reck et al., 2015), the increased permeability of the cell might also allow for nonspecific passage of the ComS peptide. These conditions may therefore render both the ComS and XIP transport systems dispensable for transformation, and the rcrPQ double deletion strain under these circumstances may in fact reflect the phenotype of the ΔSMrcrP strain. The competence phenotype of the rcrPQ double deletion strain under chemically defined conditions (optimal for XIP signaling) was not investigated in our work and remains to be characterized in future studies.

Overall our work demonstrated that competence phenotypes associated with the deletion of each of the genes in the rcrRPQ operon strongly correlate with levels of extracellular XIP produced by each strain. Therefore, the transcriptional regulator RcrR and the two putative ABC exporters RcrPQ regulate competence development in part by regulating comS expression and/or XIP secretion.

Chapter 5: Transcriptional profiling of the oral Streptococcus mutans in response to the competence signaling peptide XIP

Iwona B. Wenderska, Andrew Latos, Benjamin Pruitt, Sara Palmer, Grace

Spatafora, Dilani B. Senadheera, Dennis G. Cvitkovitch

Author contributions: IBW, DBS and DGC conceived and designed the study. IBW wrote the manuscript. DGC, DBS, SP and GS provided scientific inputs and edits to the manuscript. RNA libraries were prepared for sequencing by BP and GS. Bioinformatics analysis was performed by SP. IBW performed the qRT-PCR experiments. Northern analyses and the transformation assay were performed by AL.

5.1 Abstract

In the cariogenic Streptococcus mutans, competence development is regulated by the ComRS signaling system comprised of the ComR regulator and the ComS prepeptide to the competence signaling peptide XIP. Aside from competence development, XIP signaling has been demonstrated to regulate cell lysis, and recently, the expression of bacteriocins, small antimicrobial peptides used by bacteria to inhibit closely related species. Our study further explores the effect of XIP signaling on the S. mutans transcriptome. RNA sequencing revealed that XIP induction resulted in a global change in gene expression that was consistent with a stress response. An increase in several membrane bound regulators, including HdrRM and BrsRM, involved in bacteriocin production, and the VicRKX system involved in acid tolerance and biofilm formation, was observed. Furthermore, global changes in gene expression corresponded to changes observed during the stringent response to amino acid starvation. Effects were also observed on genes involved in sugar transport and carbon catabolite repression, and included the levQRST and levDEFG operons. Finally, our work identified a novel heat-shock responsive intergenic region, encoding a small RNA, with a potential role in competence shut-off.

5.2 Introduction

The ability to take up and incorporate extracellular DNA (eDNA) allows bacteria to modify their genomes, leading to an increase in genetic variability and population survival under changing environmental conditions (Claverys et al., 2000). Aside from its role in genetic diversity, the process of transformation by genetically competent cells has been suggested to play a role in utilization of eDNA as a nutrient or as a template for DNA repair of chromosomal damage (Finkel and Kolter, 2001; Prudhomme et al., 2006). In fact, genetic competence has been suggested to function as part of a general stress response mechanism in many gram-positive bacteria (Claverys et al., 2006).

In streptococci the process of transformation is often regulated by peptide signaling mechanisms that drive the expression of an alternate sigma factor comX, which, in turn, tightly regulates the expression of the DNA uptake and recombination machinery (Fontaine et al., 2015). In Streptococcus mutans, a caries-causing oral pathogen, genetic competence is regulated by the ComRS signaling system, comprised of the ComS pre-peptide to the ComX inducing peptide, XIP, and its cognate response regulator ComR (Mashburn-Warren et al., 2010). The ComS pre-peptide is secreted out of the cell and processed by an unknown mechanism. Upon import by the OppD/Ami permease, the mature XIP interacts with and activates ComR to directly regulate the expression of comX (Fontaine et al., 2013 (Fontaine et al., 2013; Mashburn-Warren et al., 2010). The ComR-XIP complex also binds to the promoter of the comS peptide, creating a positive feedback loop (Fontaine et al., 2013; Mashburn-Warren et al., 2010). Our recent observation that a comX deletion strain contains significantly less XIP in the supernatant suggests a second positive feedback loop that allows the ComX sigma factor to regulate XIP production and/or secretion (Wenderska et al., 2012). Aside from its essential role in competence regulation, ComRS and ComX also regulate lysis in a subpopulation of cells (Wenderska et al., 2012). A ComX-regulated lysin, LytF, has recently been identified as a contributor to the competence-associated cell lysis (Dufour and Levesque, 2013). This mechanism ensures the availability of DNA for genetic exchange, nutrition or chromosome repair during the stress-activated state of competence.

Recent work demonstrated that XIP signaling also regulates the expression of the ComDE two component system (TCS) that regulates the bacteriocin genes (Reck et al., 2015), which encode small antimicrobial peptides that inhibit the growth of closely related microbial species. The ComDE TCS is in an operon with the ComC prepeptide, which is processed to the competence stimulating peptide, CSP. The ComDE system directly regulates several bacteriocin genes in response to CSP and plays an indirect role in the activation of competence via ComRS (Hung et al., 2011). The tight association of competence development and bacteriocin production may provide S. mutans with a mechanism to lyse its competitors and simultaneously utilize their genetic contents to increase genome plasticity (Reck et al., 2015).

Aside from its role in competence and cell lysis, little is known regarding the role of XIP signaling in S. mutans physiology. Herein we employed strand-specific RNA sequencing (RNA-seq) to determine the effect of XIP treatment on the whole transcriptome of S. mutans UA159. As expected, we observed a large increase in competence-associated genes, the comX-regulated lysin LytF and a large number of the bacteriocin genes. In addition to the known modulation of ComDE, we identified a novel role for XIP-signaling to control HdrRM, BrsRM, and VicRKX signaling systems. These regulators function in parallel with the ComDE system to control competence and/or bacteriocin production (Banu et al., 2010; Okinaga, Niu, et al., 2010; Okinaga, Xie, et al., 2010; Senadheera et al., 2012; Senadheera et al., 2005; Senadheera et al., 2007; Smith and Spatafora, 2012; Xie et al., 2010). We also observed an increase in the expression of genes involved in nutrient stress, including the relQ ppGpp synthase, involved in the stringent response, and the LevQRST four-component system important to sugar metabolism and carbon catabolite repression. We further report the discovery of 11 intergenic regions that expressed putative short RNAs (sRNAs), and provided the first evidence for a heat shock-regulated intergenic region that negatively regulates genetic competence.

5.3 Materials and methods

5.3.1 Strains and growth conditions

S. mutans UA159 (Ajdic et al., 2002) and its comS deletion strain (ΔSMcomS) (Wenderska et al., 2012) were used in this study. S. mutans strains were grown at 37oC with 5% CO2 in either Todd-Hewitt Broth (Becton Dickinson, MD) containing 0.3% yeast extract (THYE) or chemically defined medium (CDM). Erythromycin was used as needed at a concentration of 10 µg/ml. Synthetic XIP peptide was synthesized using F- MOC chemistry (Advanced Protein Technology Centre, Hospital for Sick Kids, Toronto, ON, Canada). Stock concentrations of 1 mM of XIP were prepared in DMSO.

5.3.2 RNA extraction and preparation for sequencing

Bacterial cultures of UA159 and ΔSMcomS used for RNA extraction were grown overnight in THYE, washed and resuspended in fresh CDM. The cultures were then diluted 1:20 and grown to OD600 of 0.4-0.5 in the presence of 1 µM XIP or 1% DMSO vehicle control. Cells were harvested by centrifugation and resuspended in Trizol reagent (Invitrogen) prior to RNA isolation using the FastPrep system (Bio 101 Savant) as specified by the manufacturer. Total RNA was purified using the RNeasy purification kit and enriched for messenger RNA (mRNA) with the MicrobExpress kit to remove 23s and 16s ribosomal RNA (rRNA).

5.3.3 Preparation of mRNA libraries for Illumina deep-sequencing (RNAseq)

mRNA was fragmented into 50-200 nucleotide fragments by incubation in RNA Fragmentation Reagent (Ambion) for 15 min. Strand-specific cDNA libraries were prepared as described by Parkhomchuk et al. (Parkhomchuk et al., 2009). Briefly, the fragmented mRNA was then converted into double-stranded cDNA using the

SuperScript Double-Stranded cDNA Synthesis Kit (Invitrogen), as per manufacturer instructions with the following changes: After the first strand synthesis, the products were purified using Illustra Microspin G-50 columns to remove all the dNTPs, and a dUTP mixture containing dUTP, in lieu of dTTP, was used for the second strand synthesis. Samples were sequenced by the University of Vermont Cancer Center Advanced Genome Technologies Core. Each sample was sequenced at a minimum of 1000X coverage using 100 bp single end-reads on an Illumina HiSeq1000. Staphylococcus aureus was included as a control for strandedness.

5.3.4 Data analysis

RNAseq data was analyzed using the Galaxy server hosted by the research computing center at the University of Florida as previously described by Zeng and colleagues (Zeng and Burne, 2016). Briefly, reads were mapped to the S. mutans UA159 genome using Map with Bowtie for Illumina (version 1.1.2), with the default setting for single-end reads. Mapped reads corresponding to each gene were counted using htseq-count for both sense and anti-sense transcripts. The data presented in Table 1 are based on standardized mapped reads for the respective genes from single biological samples. Count data was standardized by reads per kilobase per million total reads per sample (RPKM). Specific results of interest were verified by qRT-PCR.

5.3.5 qRT-PCR

Overnight cultures of wild-type UA159 and ΔSMcomS strain in THYE were washed and resuspended in CDM. The cultures were then further diluted 20x in fresh

CDM and grown to an OD600nm of 0.4-0.5 in the presence of varying concentrations of XIP. RNA isolation, DNAse treatment, cDNA synthesis, qRT-PCR, and expression analyses were carried out as previously described (Wenderska et al., 2012). Primers used for qRT-PCR were as follows: comC (For: GACTGATGAATTAGAGATTATCATT GG, Rev: TTTCCCAAAGCTTGTGTAAAACT), comD (For: CGATGCTGTCAAGGGTAT CTTGTC, Rev: CAAGCAACTCCATCTCAGGCAG), comE (For: CCTGAAAAGGGCAA TCACCAG Rev: GGGGCATAAACTCAGAATGTGTCG), relQ (For: CGGTAAAGTTCA

TCTGTGTCATCAC Rev: AGACGAGGAAATGCGGCAG), vicR (For: CGCAGTGGCTG AGGAAAATG Rev: ACCTGTGTGTGTCGCTAAGTGATG), vicK (For: CGCAGTGGCTG AGGAAAATG Rev: ACCTGTGTGTGTCGCTAAGTGATG), vicX (For: TGCTCAACC ACAGTTTTACCG Rev: GGACTCAATCAGATAACCATCAGC), levD (For: GGAAGCCC TTTGACAACAGC Rev: CTGCCATTGGTAAGTTCATCCC), levR (For: ACATCTGGAT TAATCATGGC Rev: AAAGCTCTTCAATATGGTGC), manL (For: TGGCTATCGGA ATCGTTATCGC Rev: ATCATCAGGTCCTTCACTTGGC), hdrR (For: AAGCCATTTGC TTCTGCG Rev: TGGGGGTAGAGGAGAAAGAC), hdrM (For: GGTGAAACCAATCTGC GTATTC Rev: CCATTTGTGCTAGGAAAACCTG), brsR (For: CACGGAAAACAAACAG GTC Rev: TTCACCTTGGGAGATACG), brsM (For: GGCGTTTTACAAGGATTTGC Rev: GCTAAGAGAAGTGGTAGGACAATG). Expression was normalized to that of the 16S rRNA gene, and statistical analyses were performed on four independent experiments using Student’s t-test (P < 0.05).

5.3.6 Northern blot detection

Total RNA was isolated from UA159 cultures using Direct-zolTM RNA MiniPrep (Zymo Research). Five micrograms of total RNA was loaded into each lane and resolved on a 8% (wt/vol) polyacrylamide denaturing gel with 7.0 M urea. The RNA was transferred to a nylon membrane (Fermentas) and cross-linked using UV light for 5 minutes. The membranes were pre-hybridized in digoxygenin (DIG) Easy Hyb buffer (Roche) for 30 minutes at 42°C which was followed by a hybridization step using DIG High Prime DNA probes (25 ng/ ml) in DIG Easy Hyb buffer at 42°C overnight. Genomic DNA was amplified using region specific primers to generate probes that were then labeled using the DIG High Prime Labeling Kit (Roche) following the supplier’s instructions. The membrane was then incubated with CSPD (Roche), which resulted in a chemiluminescent signal that was visualized using a chemiluminescent detector (Bio- Rad). The transcript expression levels were quantified by densitometry using the ImageJ64 program (National Institutes of Health, Bethesda). The 5S rRNA was used as a loading control.

5.3.7 Rapid amplification of cDNA ends (RACE)

The transcription start and stop sites were mapped for several intergenic regions. The start sites were determined by using 5′ rapid amplification of cDNA ends (RACE) techniques using the RLM-RACE kit (Thermo Fisher, MA). Briefly, 10 µg of total RNA was treated using calf intestinal phosphatase. The RNA was then treated using tobacco acid pyrophosphatase and ligated to an adapter sequence, which was reverse transcribed using an outer primer. PCR was then performed using an inner nested primer with a region-specific primer to amplify the region along the transcript, this DNA was then pooled and column purified and sequenced by ACGT (Toronto, ON). The sequenced result was aligned to the NCBI sequence of the same region in order to locate the 5’ end.

A search for 3’ stop sites was also performed. The procedure used the RLM- RACE kit (Thermo Fisher, MA) according to the manufacturer’s instructions. Briefly, 1 µg of total RNA was reverse transcribed in the presence of the 3’RACE adapter. The reaction was then PCR amplified using the 3’RACE adaptor primer and a region specific primer. To determine the location of the 3’ end, the resulting PCR product was column purified, sequenced and aligned to the NCBI sequence of the same region.

5.3.8 Transformation assays

To perform genetic transformation assays, an overnight culture of S. mutans grown in THYE at 37°C was centrifuged at 4,500 RPM for 4 min at room temperature and resuspended in 1 volume of THYE. The washed culture was diluted 1:20 in THYE. Aliquots (200 µl) of the culture were transferred into a sterile microplate (Costar, Corning, Corning, NY). In test samples, 1 µg of plasmid (pDL277) was added. The sample was incubated at 37°C with 5% CO2 under static conditions. After 5 hours, test samples (20 µL using serial dilutions in PBS) with or without plasmid were plated on THYE plates supplemented with spectinomycin (10 µg/mL). After 48 hours of incubation at 37°C, the transformation frequency was calculated as percent survivability by counting the antibiotic resistant colony forming units (CFUs) divided by the total number

85 of viable CFUs x 100. The assay was performed with a minimum of three biological replicates and three technical replicates.

5.4 Results

5.4.1 Transcriptome changes in S. mutans UA159 in response to XIP

To determine the global effects of XIP signaling, we examined the effects of the peptide on the transcriptome of S. mutans UA159 using RNA deep sequencing

(RNAseq). The cells were grown in CDM to OD600 0.4-0.5 in the presence of either 1 µM XIP or the 1% DMSO vehicle control. Overall, exposure to XIP resulted in the upregulation of 105 genes by greater than twofold in the wild-type strain (Figure 5.1, Table S5.1). Of these, approximately 14 genes were involved in genetic competence and transformation, 8 genes in DNA metabolism, recombination and repair, and 10 genes in signal transduction and transcriptional regulation. As expected, this list includes the central competence regulator comX (4.4-fold), directly regulated by ComR- XIP, as well as many of the ComX-regulated late competence genes involved in DNA uptake and recombination. The strongest upregulation was observed in the 23 genes involved in cell killing and bacteriocin production, including the ComCDE signaling system (2.9-, 2.8- and 3.6-fold, respectively) that directly regulates bacteriocin expression, and the SMU.166-168 operon (2.8-, 3.1-, and 4.4-fold), proposed to encode a putative 3-component toxin-antitoxin system, comprised of the toxin-antitoxin complex and their transcriptional regulator, respectively (Mankovskaia, 2013). Upregulation was also observed in 6 genes involved in energy metabolism, 9 genes in transport and binding, and 23 genes with unknown functions. The XIP peptide caused a repression of 580 genes in UA159, 22 of which were involved in amino acid biosynthesis, 23 encoding purines, pyrimidines, nucleosides and nucleotides, 85 genes involved in protein synthesis, 53 encoding tRNA and 199 genes with unknown functions (Table S5.1). Quantitative real time PCR (qRT-PCR) was used to confirm the role of XIP in the regulation of the bacteriocin regulatory system comDE, as previously reported by Reck et al, but also in the regulation of the comC gene (Figure 5.2a).

Functional Category: 10- Biosynthesis of cofactors 1- amino acid biosynthesis 11- Cell envelope 2- competence and transformation 12- Adaptations to atypical conditions 3- DNA metabolism, recombination and repair 13- Cell division 4- Energy metabolism 14- mobile and extrachromosomal element functions 5- Central intermediary metabolism 15- pathogenesis and toxin production 6- purines, pyrimidines and nucleosides and nucleotides 16- fatty acid and phospholipid metabolism 7- transport and binding 17- Protein synthesis and protein fate 8- signal transduction and transcriptional regulation 18- tRNA 9- cell killing and bacteriocin production 19- other 100 M XIP µ

0.1 Fold-change in expression in UA159 in response to 1 1 to response in UA159 in expression in Fold-change

0.01 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

Functional Category

Figure 5.1 Functional characterization of the changes (>2-fold) in the UA159 transcriptome in response to 1 µM XIP.

a 1 µM XIP 5 µM XIP 10 µM XIP

** * 10 ** * * *

5 * *

0 Gene expression relative to DMSO to relative expression control Gene comC comD comE

8 * 1 µM XIP 5 µM XIP 10 µM XIP 6

4 * * * * 2

* *

Gene expression relative to DMSO to relative expression control Gene 0

levR levD manL

Figure 5.2 The effect of increasing amounts of XIP on genes involved in bacteriocin regulation (a) and sugar uptake (b). RT-PCR analysis of comC/D/E (a) or levR/D and manL

(b) gene expression was performed in UA159 grown in CDM to OD600 0.4 in the presence of 1 µM XIP. *p<0.05, **p<0.005

Notably, XIP exposure increased the expression of the levQRST four-component signaling system (2.3-, 2.4-fold, 2.7- and 2.7-fold, respectively), comprised of the levRS two-component system and two sugar-binding proteins LevQ and T, responsible for sensing fructose levels in the extracellular milieu (Smith and Spatafora, 2012; Zeng et al., 2006). The levDEFG operon encoding components of enzyme II of the fructose/mannose phosphotransferase (PTS) system was also upregulated 2.2-, 2.5-, 2.3-, and 2.4-fold, respectively. The LevDEFG is primarily involved in fructose uptake and is upregulated by the LevQRST signaling system in response to low fructose levels (Smith and Spatafora, 2012; Zeng et al., 2006). Three transcripts corresponding to Enzyme II components of the mannose PTS system required for transporting glucose, mannose, sucrose or cellibiose were downregulated (-4.2-fold, -3.8-fold and -3.9-fold for manN/L/M, respectively). These components, including manL, are important for carbon catabolite repression of the levDEFG operon (Zeng and Burne, 2010). The low expression of manL could, therefore in part, explain the increase in levDEFG expression. The effects of XIP on representative genes of the lev operons and the manL gene were confirmed by qRT-PCR (Figure 5.2b).

5.4.2 Changes in gene expression in ∆SMcomS strain in response to 1 µM XIP

Although genetic transformation in S. mutans UA159 does not occur until later stages of growth in CDM, we wanted to ensure that any endogenous XIP production in WT mid-logarithmic cells did not mask any effects on the S. mutans transcriptome. As a result, we examined the transcriptome of the comS deletion strain that does not produce any endogenous XIP. Treatment of ∆SMcomS with 1 µM XIP resulted in increased expression in almost 300 genes (Figure 5.3, Table S5.2). Of these, the highest upregulation was observed in genes involved in competence and transformation. The central competence regulator comX was upregulated by 61.4-fold, whereas the genes encoding components of the DNA uptake machinery were upregulated as much as 1000-fold.

Aside from competence related genes, several genes involved in virulence were upregulated in ∆SMcomS in response to 1 µM XIP. These included ABC transporters

89 with predicted involvement in multidrug tolerance, and genes involved in adaptation to atypical conditions and detoxification, including the relQ synthase (2.8-fold) of ()ppGpp; a mediator of the stringent response that reduces cell growth in response to nutrient starvation (Potrykus and Cashel, 2008). Consistent with the stringent response, we also observed a downregulation in genes involved in the synthesis of purines, pyrimidines, nucleosides and nucleotides. A purine-operon repressor was upregulated 4.0-fold and likely contributed to the low expression of its target genes. Genes involved in pathogenesis, toxin production and resistance were also upregulated and included several genes involved in mutanobactin production (SMU.1335c-1340), a CSP- regulated secondary metabolite implicated in interkingdom competition with Candida albicans (Joyner et al., 2010). In addition, at least a 2-fold increase was observed in 10 genes encoding transposases and other mobile elements.

A number of signal transduction and transcriptional regulators were also increased in response to XIP treatment, and included the vicRKX signaling system (2.0- , 2.0- and 2.4-fold, respectively), strongly associated with virulence of S. mutans (Senadheera et al., 2012; Senadheera et al., 2005; Senadheera et al., 2007), and the hdrRM (1.5- and 2.1-fold, respectively) and brsRM (5.0- and 4.2-fold, respectively) signaling systems, both of which have previously been demonstrated to function upstream of ComX to regulate competence development and bacteriocin production (Okinaga, Niu, et al., 2010; Okinaga, Xie, et al., 2010; Xie et al., 2010). The levQRST system was upregulated 3.5-, 3.9-, 4.1-, 4.3-fold, respectively, along with its associated fructose/mannose phosphotransferase system, levDEFG, which was increased by 3.4-, 3.0-, 2.8-, and 3.3-fold, respectively. The effects of XIP on these membrane-bound regulators of bacteriocin production, sugar metabolism and virulence expression were confirmed by examining one or more genes representative of each operon by qRT-PCR (Figure 5.4).

1000

100 SMcomS in response to 1 response in SMcomS ∆

1 Fold-change in expression in in expression in Fold-change 0.1 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

Functional Category

Figure 5.3 Functional characterization of the changes (>2-fold) in the ∆SMcomS transcriptome in response to 1 µM XIP.

4 1 µM XIP 3 1 µM XIP 5 µM XIP 5 µM XIP 10 µM XIP * 10 µM XIP 3 * * * 2

1 1

0 0 Gene expression relative to DMSO to relative expression control Gene Gene expression relative to DMSO to relative expression control Gene relQ vicR vicX

10 1 µM XIP 5 1 µM XIP 5 µM XIP 5 µM XIP 10 µM XIP 10 µM XIP 4

* 3 * * 5 ** * * ** * 2

0 0 Gene expression relative to DMSO to relative expression control Gene Gene expression relative to DMSO to relative expression control Gene brsR brsM hdrR hdrM

15 10 1 µM XIP * 1 µM XIP 5 µM XIP 5 µM XIP 10 µM XIP * * 10 µM XIP *

10 * * 5 * ** ** * 5 * * *

0 0 Gene expression relative to DMSO to relative expression control Gene Gene expression relative to DMSO to relative expression control Gene levR levD comE comC comD

Figure 5.4 Expression of selected genes in ∆SMcomS in response to increasing concentrations of XIP. RT-PCR analysis of gene expression was performed in ∆SMcomS grown in CDM to OD600 0.4 in the presence of increasing amounts of XIP. *p<0.05, **p<0.005

5.4.3 Putative small RNAs expressed in the presence of XIP

Intergenic regions serve several important regulatory functions and often contain small RNAs (sRNAs). These small transcripts can bind to proteins to regulate their function, or base-pair with RNAs, modulating their stability and translation. Identification and characterization of sRNAs is therefore important for understanding the regulation of bacterial processes. As a result, we examined large intergenic regions of greater than 500 bp for unidentified transcripts. Of these, products were identified in 11 regions, 5 of which were responsive to XIP treatment (Table 5.1). The transcriptional 5’ start sites were mapped for several of the transcripts and are listed in Table 5.3. Of note is the intergenic region between SMU.153 and SMU.154, which is located downstream of bacteriocin genes and is induced 5-, 10- and 20-fold in response to 1 µM, 5 µM and 10 µM XIP, respectively (Figure 5.5). Furthermore, an induction of the intergenic region between SMU.82 (dnaK) and SMU.83 (dnaJ) was observed at higher XIP concentrations. Specifically, this region increased by 2- and 4-fold in response to 5 µM and 10 µM XIP, respectively. The SMU.82-83 intergenic region is located within the heat shock operon and in addition to increased transcription with increasing concentrations of XIP, this region has been shown to be upregulated in response to heat stress when mid-logarithmic cells were exposed to high temperatures of 50 oC for 20 min (Latos, 2016). A deletion of the intergenic region (∆SMU.82-83) exhibited increased natural competence as compared to UA159 (Figure 5.6) suggesting a role in negative regulation of competence.

Table 5.1. Intergenic regions up- or down-regulated in UA159 in response to 1 µM

XIP.

Upstream US gene Downstream DS gene sRNA 5' RACE XIP sensitivity gene direction gene direction direction Northern start site smu.60 > smu.61 > -> Yes nd nd smu.82 > smu.83 > -> Yes 87,809 upregulated smu.97 > smu.99 > -> Yes 99,434 downregulated smu.97 > smu.99 > <- Yes 99,836 smu.97 > smu.99 > <- Yes 99,989 smu.97 > smu.99 > -> Yes 99,930 smu153 > smu.154 > Yes nd upregulated smu.217c < smu.218 > Yes nd nd smu.259 > smu.260 > Yes nd nd smu.305 > smu.307 > Yes nd nd smu.770c < smu.771c < <- No 719,052 upregulated smu.788 > smu.789 > Yes nd downregulated smu.1063 < smu.1064c < <- No 1,008,320 nd smu.1405c < smu.1406c < Yes nd nd nd = not determined

1 µM XIP 25 * 5 µM XIP 10 µM XIP

15 * ***

10 * * *** * 5

Gene expression relative to DMSO to relative expression control Gene * *

SMU.82-83 SMU.97-99 SMU.153-154 SMU.788-789 SMU.770c-771c

Figure 5.5 Expression of selected intergenic regions in response to increasing concentrations of XIP. qRT-PCR analysis of gene expression was performed with UA159 grown in CDM to OD600 0.4 in the presence of increasing amounts of XIP. *p<0.05, ***p<0.001

101

100

10-1

10-2

10-3 Transformation frequency (%) frequency Transformation

10-4 UA 159 ΔSMU.82-83

Figure 5.6 The effect of ∆SMU.82-83 on transformation frequency. Overnight cultures of UA159 and ∆SMU.82-83 were diluted 1:20 in fresh THYE and incubated in the presence or absence of pDL277 (specR). Results show the mean %TF of three independent experiments ± standard error.

5.5 Discussion

A microbial population that is genetically competent is better equipped to withstand environmental changes and stressful conditions. The state of competence increases genetic variability, thereby increasing the likelihood of survival for some members of the population during an environmental perturbation (Claverys et al., 2006). Furthermore, access to eDNA provides the competent population with a source of DNA for genetic repair, or nutrients during starvation (Claverys et al., 2006). In S. mutans, the state of competence is regulated by the competence peptide XIP (Fontaine et al., 2013; Mashburn-Warren et al., 2010). Since its discovery and implication in genetic transformation, XIP signaling has also been shown to regulate cell lysis and bacteriocin expression (Reck et al., 2015; Wenderska et al., 2012). Our study further examined the effects of XIP on the transcriptome of S. mutans using RNAseq. The cells were exposed to XIP during growth to mid-logarithmic phase to ensure sufficient exposure time for XIP to capture both direct and indirect effects of XIP on gene expression in S. mutans. This allowed for a global overview of the transcriptional changes associated with XIP signaling. Additionally, to ensure that any endogenous XIP production in WT mid-logarithmic cells did not mask any effects on the S. mutans transcriptome, we also examined the transcriptome of the comS deletion strain, which does not produce any endogenous XIP. Although a large overlap between the transcriptomes of UA159 and ∆SMcomS in response to 1µM XIP was observed, XIP treatment of ∆SMcomS resulted in the strongest upregulation of competence and DNA uptake loci (a transcriptional profile that matches competence induction), while the same treatment of the wild-type strain resulted in the highest increase in genes associated with bacteriocin expression and cell lysis (matching the lytic phenotype previously observed with higher XIP amounts (Wenderska et al., 2012) (Figure 5.1, Figure 5.3). Furthermore, the ∆SMcomS transcriptome in response to 1µM XIP contained increased number of genes involved in stress tolerance and pathogenesis. The differences observed between the two strains may be due to the amplification of the XIP signal in the wild type UA159 strain, which contains an undisrupted ComRS autoregulatory loop.

Overall, our results demonstrated that XIP induces a global change in gene expression in S. mutans that supports not only DNA uptake, repair mechanisms and

96 bacteriocin production, but also induces a global change in expression that favours genes involved in stress adaptation and virulence. We show that in addition to comCDE, 1 µM XIP also induced the transcription of hdrRM, brsRM, and vicRKX signaling systems (Figure 5.4). All of these systems have previously been implicated in one or more of the phenotypes associated with CSP signaling, including genetic transformation and bacteriocin production (Okinaga, Niu, et al., 2010; Okinaga, Xie, et al., 2010; Xie et al., 2010). The vicRKX operon is also an important regulator of acid tolerance, bacterial adherence and biofilm formation (Senadheera et al., 2012; Senadheera et al., 2005; Senadheera et al., 2007; Smith and Spatafora, 2012).

XIP treatment also resulted in the upregulation of the relQ (p)ppGpp synthase, and the repression of genes involved in cell growth and replication, a transcriptional profile that is consistent with the induction of the stringent response, usually activated in response to starvation (Figure 5.1, Figure 5.3). Collectively referred to as (p)ppGpp, this group of signaling molecules include the guanosine 3’-diphosphate, guanosine 5’- triphosphate, and guanosine 3’,5’-bispyrophosphate, and are responsible for global transcriptional changes that favour adaptation to a semidormant state, reducing chromosomal replication and growth of the population (Potrykus and Cashel, 2008). Furthermore, activation of the stringent response often results in the release of carbon catabolite repression (CCR). CCR mechanisms repress genes involved in the uptake and metabolism of “non-preferred” sugars to ensure the efficient uptake and utilization of the “preferred” carbohydrates. However, during starvation, the transport systems for non-preferred sugars are derepressed. Similarly to the stringent response, XIP treatment resulted in the downregulation of the ManL regulator of CCR that represses the expression of the lev operons involved in fructose uptake and metabolism. As expected, a decrease in manL, along with an increase in the levQRST four component signaling system that, in response to fructose levels, regulates LevDEFG fructose/mannose phosphotransferase system, resulted in a significant induction of the levDEFG operon (Figure 5.2) (Smith and Spatafora, 2012; Zeng and Burne, 2008, 2010; Zeng et al., 2006). Our transcriptome results therefore suggest XIP signaling induces a response that mirrors the stringent response to starvation.

In the gram-positive Bacillus subtilis genetic competence is tightly controlled and coordinated with the production of the (p)ppGpp alarmone and the stringent response (Erickson and Copeland, 1972; Inaoka and Ochi, 2002). This link between genome replication and natural transformation may provide a mechanism that ensures correct incorporation of the transformed DNA into the genome. Although the stringent response and genetic competence has previously been shown to be coregulated in S. mutans (Seaton et al., 2011), this is the first report that indicates the competence regulatory system ComRS may also regulate (p)ppGpp production. The effect of XIP on the accumulation of the (p)ppGpp messengers remains to be examined in future studies.

Our transcriptome data revealed 5 novel intergenic regions affected by XIP signaling (Figure 5.5). Of these the SMU.153-154 region was strongly upregulated by increasing concentrations of XIP and is located 3’ of bacteriocin synthetic genes, and therefore may be involved in bacteriocin induction. We further investigated the intergenic region SMU.82-83 located within a heat shock operon. This region was previously shown to be induced by heat stress (Latos, 2016) and our transformation assays reveal it may play a role in competence shut-off (Figure 5.6). The mechanism of competence regulation by SMU.82-83 remains to be examined in future experiments.

5.6 Conclusions

Our work demonstrates that in S. mutans, the competence-inducing XIP peptide not only controls DNA transformation and bacteriocin production, but also induces a response that resembles the stringent response to amino acid starvation. Further, we report five XIP-responsive intergenic regions expressing putative short RNAs (sRNAs), and provide the first evidence for a heat shock-regulated intergenic region that negatively regulates genetic competence. Furthermore, this work provides a greater understanding of how global gene expression patterns change in response to the peptide XIP. Collectively, these results provide further evidence that multiple stress response mechanisms are linked through the genetic competence signaling pathway in S. mutans.

5.7 Acknowledgements

We would like to thank Martha Cordova for her assistance with RNA extraction. I.B.W. was supported by the Ontario Graduate Scholarship. D.G.C. is a recipient of the NIH grant R01DE013230 and CIHR-MT15431. This research was also supported, in part, by NIH R01DE014711 to G.S.

Chapter 6: Summary, future directions and significance

6.1 Summary

This dissertation examined the biological function of the ComRS signaling system, and its regulation, in the cariogenic Streptococcus mutans. The contributions of this dissertation are summarized as a part of the current model for ComRS signaling, depicted in Figure 6.1. Briefly we have demonstrated a novel role for the ComRS regulatory system in the regulation of competence-associated cell lysis (Chapter 3). Our work revealed that excess amount of XIP induced cell lysis in as much as 82% of the population in a comR and comX dependent mechanism (Wenderska et al., 2012). This coordination of competence development and cell lysis may ensure availability of eDNA for DNA uptake by the competent subpopulation. In accordance with recent literature, our transcriptome analysis revealed that aside from ComX-regulated lysin LytF that has previously been shown to contribute to the competence-associated cell lysis (Dufour et al., 2013), XIP signaling also regulates key bacteriocin genes and bacteriocin regulators, including the ComCDE, HdrRM and BrsRM signaling systems (Chapter 5, Reck et al., 2015). Induction of bacteriocins may allow S. mutans to further scavenge eDNA from closely related species in the oral biofilm.

Our transcriptome analysis further suggests that XIP signaling plays a role in stress response that mimics the expression profile of the stringent response due to starvation (Chapter 5). A downregulation in genes involved in cell growth and replication was observed, including protein biosynthesis genes, genes encoding purines, pyrimidines, nucleotides and nucleosides, as well as tRNA. In accordance with the stringent response, XIP treatment further appeared to release carbon catabolite repression, resulting in the upregulation of the Lev systems involved in fructose sensing, transport and metabolism.

The (p)ppGpp alarmones of the stringent response are regulated by the genes of the rcrRPQ operon (Seaton et al., 2011). Deletions in this operon not only affect the

100 amount of (p)ppGpp produced but also display drastic competence phenotypes (Seaton et al., 2011, 2015). Our work demonstrates that the rcrRPQ operon contributes to genetic competence by regulating comS expression and the extracellular amounts of the mature peptide XIP (Chapter 4). We were one of the first groups to identify and quantify naturally produced XIP in S. mutans (Wenderska et al., 2012), and have further utilized this to characterize the role of RcrRPQ in XIP production. RcrR positively regulates comS expression and the resulting amounts of extracellular XIP. The RcrP ABC transporter negatively regulates comS expression and competence development, perhaps by expelling an unidentified competence stimulatory factor. Our work further suggests RcrQ as the putative exporter for the ComS prepeptide in S. mutans. The exporter for ComS has been unknown in all streptococcal species that utilize the ComRS system for competence induction. Whether a similar mechanism of ComS export exists in other streptococcal species has yet to be determined.

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Figure 6.1 Overview of the current model for ComRS signaling in S. mutans including specific contributions from chapters of this dissertation. Where appropriate references have been made to other studies with key contributions to the ComRS model. Key aspects of the model requiring further research have been indicated by a question mark or a dashed arrow.

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6.2 Future directions

This dissertation was largely descriptive in nature and paves the road for future mechanistic studies elucidating the role of RcrRPQ in comS regulation, and the role for ComRS in stress response, (p)ppGpp synthesis and growth regulation. We have demonstrated that the comS regulation by the RcrR regulator of the MarR-family may be indirect, possibly through the uncharacterized ComX-mediated positive feedback loop (Wenderska et al., 2012). Future studies examining the role of ComX in RcrR- mediated regulation of comS would provide insight into the competence signaling pathway. Furthermore, MarR-family of regulators are known to alter their binding affinity by specific ligands. Identification and characterization of these ligands would provide further insight into competence regulation by RcrR.

Our work demonstrated that the RcrP ABC transporter is a negative regulator of competence development. Further studies should aim at the identification of the substrate for the RcrP exporter and characterization of its role in competence regulation. Using LCMS/MS, comparison of the spectra of secreted factors by wild-type UA159 and the rcrP deletion strain will allow for the identification of the substrate for RcrP. The identified substrate should further be tested for its role in XIP-associated phenotypes. Quantitative RT-PCR and protein interactions with key regulators of competence, including the ComR-XIP complex and the ComX sigma factor, will also help determine the mechanism of action for the RcrP substrate in competence regulation.

Interestingly, we have observed that the competence negative phenotype of ΔSMrcrQ can only be complemented by exogenously added XIP in the absence of a functional comS gene. This suggests that the high expression of comS may be interfering with XIP activity. Future studies should examine whether the full-length ComS peptide has a functional role in competence regulation beyond its role as a prepeptide for the mature XIP.

Our transcriptome analysis revealed that XIP signaling results in the stringent response, however its effect on the intracellular levels of the (p)ppGpp alarmones have

103 not been investigated. Therefore further studies are needed to examine in detail the role for XIP signaling in the regulation of the intracellular (p)ppGpp pool and the stringent response.

Our transcriptome data represents the overall response to XIP treatment in the entire population and speaks nothing to the distinct subpopulations that may be present. From our studies and others in the field, we now know that XIP signaling carries out several biological functions, including genetic competence, cell lysis, bacteriocin production and the induction of the stringent response. Future research at a single cell level should aim at identifying these subpopulations in order to better understand the coordination of these events within the population. It is possible that the competence- associated cell lysis occurs in the same cells expressing high levels of bacteriocins as a mechanism of their release. Examining the coexpression of LytF and bacteriocin genes using fluorescent reporters would provide insight whether the increase in bacteriocin expression and the expression of the lysin occurs in the same subpopulation. Furthermore, the stringent response may be specifically induced in the competent subpopulation to allow for proper coordination of DNA replication and cell lysis. In Bacillus subtilis, in which competence development is also coregulated with (p)ppGpp synthesis, the competent subpopulation exhibits a decreased rate of DNA and stable RNA synthesis (Erickson et al., 1972; Inaoka et al., 2002). Examining these processes at a single level will determine whether a similar mechanism exists in S. mutans.

6.3 Significance

Streptococcus mutans is a leading etiological agent of dental caries, a global epidemic and one of the most common chronic diseases worldwide. S. mutans resides in the oral biofilm, where it produces organic acids from dietary sugars causing the dissolution of tooth enamel. In this biofilm mode of growth S. mutans has been shown to be highly competent for the uptake of extracellular DNA. The process of transformation of genetically competent cells increases chromosomal diversity enhancing population survival under stressful conditions. In S. mutans competence is regulated by the

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ComRS peptide signaling system. Our work has demonstrated that aside from its role in genetic transformation the ComRS signaling system plays a role in cell lysis, bacteriocin regulation, and stress response. Furthermore we presented evidence towards the regulation of ComRS by the stringent response regulatory operon rcrRPQ, and identified a putative exporter for ComS, thereby providing insight into the XIP signaling pathway. Understanding the peptide signaling systems that govern adaptation mechanisms in S. mutans, including horizontal gene transfer and the stringent response, can provide us with the knowledge to design more effective therapeutic strategies for the prevention of caries.

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Supplemental Tables

Table S5.1. Genes up- or down-regulated in S. mutans UA159 in response to 1 µM XIP.

Gene ID Fold Change Annotation

Cell killing and bacteriocin production: SMU.1910c 26.0 hypothetical protein SMU.1912c 24.2 hypothetical protein SMU.1913c 19.0 putative immunity protein; BlpL-like; immA SMU.1914c 31.9 hypothetical protein; nlmC; mutacin V SMU.1908c 22.1 hypothetical protein; immunity protein SMU.1909c 21.7 hypothetical protein SMU.1903c 20.0 putative bacteriocin secretion protein; bsmL SMU.1904c 18.3 hypothetical protein SMU.1905c 18.4 hypothetical protein SMU.1906c 17.9 bacteriocin-related protein SMU.150 10.4 nlmA; non-lantibiotic mutacin IV A SMU.151 14.0 nlmB; non-lantibiotic mutacin IV B SMU.152 8.9 hypothetical protein SMU.153 7.8 hypothetical protein SMU.423 10.2 nlmD; possible bacteriocin SMU.836 8.9 hypothetical protein SMU.925 2.5 cipI; bacteriocin immunity protein SMU.1917 3.6 comE; response regulator SMU.1916 2.8 comD; histidine kinase SMU.1915 2.9 comC; competence stimulating peptide SMU.168 2.8 transcriptional regulator toxin-antitoxin SMU.166 2.8 putative toxin-antitoxin system SMU.167 3.1 putative toxin-antitoxin system SMU_2035 -2.0 possible bacteriocin self-immunity protein

Competence and DNA transformation: SMU.2086 4.5 competence damage-inducible protein A SMU.1983 5.6 competence protein comYD SMU.1984 5.5 competence protein comYC SMU.1981c 5.3 competence protein comG SMU.1001 5.2 dprA; DNA processing protein; Smf family SMU.1997 4.4 comX; competence-specific sigma factor SMU.625 3.9 comEA; competence protein SMU.626 3.8 comEC; competence protein

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SMU.498 3.4 comFA; late competence protein F SMU.499 4.0 comFC; late competence protein SMU.644 2.0 coiA; competence protein SMU.1981c 5.3 competence protein G SMU.1985 5.5 comGB; competence protein SMU.1987 7.6 comGA; late competence protein competence factor transporting ATP-binding / permease SMU.1881c -2.4 protein

Mobile and extrachromosomal element functions: SMU.1817c -3.7 transposon-related, maturase-related protein fragment SMU.1816c -3.2 transposon-related, maturase-related protein fragment transposase, IS1216, also reported on plasmid, SMU.226c -2.6 truncated SMU.106c -2.5 transposase fragment SMU.1332c -2.3 transposase SMU.1374 -2.1 IS30 transposase related protein SMU.1329c -2.1 transposase fragment SMU.1331c -2.1 transposase SMU.1355c -2.1 transposase SMU.1330c -2.0 transposase, IS1167, fragment SMU.94c 3.3 transposase fragment SMU.1024c -3.8 transposase fragment SMU.1029 -9.2 hypothetical – transposon SMU.93c -2.1 putative transposase SMU.1407c -2.0 transposase, ISSmu1

Pathogenesis and toxin production: SMU.610 -3.5 cell surface antigen SMU.1396 -2.4 glucan-binding protein C

Cell division: SMU.454 -2.1 cell division protein SMU.1394 -2.0 GTP-binding protein LepA SMU.1276c -2.0 cell division regulator SMU.551 -2.0 cell division protein FtsA SMU.1003 3.5 glucose-inhibited division protein

DNA metabolism, replication, recombination and repair: SMU.1055 8.2 DNA repair protein RadC SMU.1967 7.9 ssbA; single-stranded DNA-binding protein SMU.505 7.5 dpn; adenine-specific DNA methylase SMU.506 5.8 ssuRB; type II restriction endonuclease

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SMU.1002 3.6 topA; DNA topoisomerase I SMU.2085 3.4 recA; recombinase A SMU.64 2.7 ruvB; holliday junction DNA helicase SMU.60 2.1 alkD; DNA alkylation repair enzyme SMU.510c -3.3 deoxyribonuclease SMU.1258c -2.6 type II restriction endonuclease subunit SMU.342 -2.5 possible primase-related protein SMU.1313c -2.4 ATP-dependent DNA helicase SMU.469 -2.3 recombination protein U SMU.1859 -2.2 single-stranded DNA-binding protein SMU.1114 -2.1 DNA gyrase A subunit SMU.1650 -2.1 endonuclease III (DNA repair) SMU.1192 -2.0 DNA-polymerase III subunit alpha SMU.1714c -2.0 tyrosine recombinase SMU.2088 -2.0 Holliday junction DNA helicase SMU.1581 -2.0 DNA polymerase III subunits gamma and tau

Transport and binding: SMU.772 3.2 gbpD; bifunctional glucan-binding protein D and lipase SMU.1966c 2.7 levT; ABC transport ribose-binding protein SMU.651c 2.2 ABC transporter; periplasmic substrate-binding protein SMU.1963c 2.3 levQ; sugar-binding periplasmic protein SMU.864 5.6 ABC transporter; permease component SMU.1195 2.9 ABC transporter permease protein SMU.863 4.5 ABC transporter, ATP-binding protein SMU.1194 2.7 yurY; ABC transporter, ATP-binding protein SMU.1063 -5.0 amino acid ABC transporter, ATP-binding protein SMU.819 -2.0 large conductance mechanosensitive channel manganese transporter/possible HitA ferric iron-binding SMU.770c 4.8 periplasmic protein SMU.1175 -2.5 dagA; sodium:alanine (or glycine) symporter cylB; ABC transporter, permease and solute binding SMU.432 -3.3 protein SMU.1062 -5.0 glycine-betaine binding ABC transporter permease SMU.602 -2.5 sodium-dependent transporter SMU.408 -2.8 xanthine/uracil permease family protein SMU.1289c -2.7 voltage-gated chloride channel family SMU.819 -2.0 large conductance mechanosensitive channel SMU.1176 -2.3 cation efflux protein SMU.1852 -2.3 cation transporter, CorA family SMU.121 -2.3 MATE efflux family , DinF SMU.540 -2.2 peroxide resistance protein / iron binding protein SMU.723 -2.2 calcium-transporting ATPase

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SMU.71 -2.4 MATE efflux family protein SMU.872 -2.0 fructose-specific PTS system enzyme IIBC component SMU.396 -2.9 glycerol uptake facilitator protein SMU.1137 -2.2 phosphate ABC transporter, permease SMU.388 -2.2 possible branched-chain amino acid permease SMU.951 -2.1 amino acid permease proline/glycine betaine ABC permease and solute SMU.1095 -2.4 binding protein branched-chain amino acid ABC transporter, substrate- SMU.1669 -2.4 binding protein SMU.1121c -2.4 ribonucleoside ABC transporter, solute-binding protein glycine betaine/carnitine/choline ABC transporter, SMU.2118 -2.1 substrate-binding protein SMU.1177c -2.0 amino acid ABC transporter, amino acid-binding protein SMU.1447c -2.0 ABC transporter, substrate-binding protein SMU.817 -2.0 amino acid ABC transporter, substrate-binding protein SMU.1136 -2.0 phosphate ABC transporter, permease SMU.242c -2.6 glutamine ABC transporter, solute binding protein SMU.1164c -2.9 ABC transporter, ATPase component SMU.1810 -2.6 ABC transporter, membrane spanning permease ABC-type multidrug / protein/ lipid transport system, SMU.922 -2.6 ATPase component SMU.1550c -2.6 ABC transporter, membrane spanning permease SMU.1668 -2.5 branched chain amino acid ABC transporter, permease SMU.567 -2.5 amino acid (glutamine) ABC transporter permease SMU.1119c -2.4 ribonucleoside ABC transporter permease SMU.1163c -2.4 ABC transporter, ATPase component glycine betaine / carnitine / choline ABC transporter SMU.2117 -2.4 permease SMU.1166c -2.4 ABC transporter permease SMU.1667 -2.4 branched chain amino acid ABC transporter, permease SMU.1118c -2.4 ribonucleoside ABC transporter permease SMU.257 -2.3 oligopeptide ABC transporter, permease SMU.256 -2.3 oligopeptide ABC transporter, permease SMU.1216c -2.3 ABC transporter, amino acid permease ABC transporter, permease protein;possible ferrichrome SMU.996 -2.3 transport system ABC transporter, permease, possibly bacteriocin SMU.656 -2.1 associated SMU.1179c -2.0 amino acid ABC transporter, permease SMU.1446c -2.0 ABC transporter, permease SMU.2148c -2.0 ABC transporter, membrane-spanning permease maltose / maltodextrin ABC transport system SMU.1570 -2.0 (permease)

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SMU.1569 -2.0 maltodextrin ABC transport system permease ABC-type multidrug/protein/lipid transport SMU.923 -2.3 system,ATPase component SMU.1063 -4.9 amino acid ABC transporter, ATP-binding protein SMU.1315c -2.6 ABC transporter, ATP-binding protein ABC transporter, ATP-binding protein; possible SMU.1695 -2.5 molybdenum transport system SMU.1178c -2.5 amino acid ABC transporter, ATP-binding protein SMU.1096 -2.5 ABC transporter, ATP-binding protein inorganic ion ABC transporter,ATP-binding protein; SMU.997 -2.4 possible ferrichrome transport system branched chain amino acid ABC transporter, ATP- SMU.1666 -2.4 binding protein SMU.1120 -2.3 ribonucleoside ABC transporter, ATP-binding protein SMU.241c -2.3 amino acid ABC transporter, ATP-binding protein SMU.1551c -2.3 ABC transporter, ATP-binding protein glycine betaine / carnitine / choline ABC transporter, SMU.2116 -2.3 ATP-binding protein, opuCA SMU.568 -2.2 glutamine ABC transporter, ATP-binding protein SMU.1431c -2.1 ABC transporter, ATP-binding protein SMU.1519 -2.1 glutamine ABC transport, ATP-binding protein ABC-type transport system ATP-binding protein SMU.1571 -2.1 (maltose) SMU.1006 -2.0 ABC transporter, ATP-binding protein SMU.2149c -2.0 ABC transporter, ATP-binding protein SMU.1445c -2.0 ABC transporter, ATP-binding protein branched chain amino acid ABC transporter, ATP- SMU.1665 -2.0 binding protein SMU.1167c -2.0 ABC transporter ATP-binding protein SMU.258 -2.0 oligopeptide ABC transporter, ATP-binding SMU.1811 -2.0 cdd4-like bacteriocin component, ScnF homolog SMU.2006 -2.3 preprotein translocase SMU.1348c -2.2 ABC transporter ATP-binding protein SMU.1365c -2.0 Permease-FtsX-like permease SMU.1366c -2.0 ABC transporter ATP-binding Protein

Signal transduction and transcriptional regulation: SMU.507 4.2 transcriptional regulator, DeoR family SMU.1193 3.9 transcriptional regulator, GntR family SMU.168 2.8 transcriptional regulator SMU.1965c 2.7 levS, histidine kinase SMU.1964c 2.4 levR, two-component response regulator SMU.1960c 2.5 levE; fructose-specific Enzyme IIB component SMU.1957 2.4 levG; fructose-specific Enzyme IID component

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SMU.1961c 2.2 levD; fructose-specific Enzyme IIA component SMU.1958c 2.3 levF; fructose-specific Enzyme IIC component SMU.2038 -2.4 pttB; phosphotransferase system SMU.1878 -3.9 manM; mannose PTS system component IIC SMU.1053 6.0 possible redox-sensing transcriptional repressor Rex SMU.1877 -3.8 manL; mannose PTS system component IIAB SMU.1879 -4.2 manD; mannose PTS system component IID SMU.1064c -3.0 transcriptional regulator SMU.1065c -3.3 transcriptional regulator, GntR family SMU.1165c -3.0 transcriptional regulator (TetR/AcrR family) SMU.1419 -2.7 transcriptional regulator SMU.921 -2.5 transcriptional regulator SMU.433 -2.5 transcriptional regulator SMU.702c -2.5 transcriptional regulator SMU.2058 -2.4 transcriptional regulator transcriptional regulator, LysR family (possible SMU.2060 -2.3 RUBISCO transcriptional regulator, RscR) SMU.1025 -2.2 transcriptional regulator SMU.1097c -2.2 transcriptional regulator transcriptional regulator, MepR protein (possible SMU.2108c -2.1 regulator of MepA) SMU.1745c -2.1 transcriptional regulator, MarR family SMU.1168 -2.0 transcriptional regulator (TetR/AcrR family) SMU.124 -2.0 transcriptional regulator, MarR family SMU.2001 -2.2 DNA-directed RNA polymerase, alpha subunit SMU.1990 -2.0 DNA-directed RNA polymerase, beta subunit SMU.336 -2.1 ribonuclease P protein component SMU.1607 -2.0 exoribonuclease R SMU.577 -2.5 sensor histidine kinase SMU.1547c -2.4 response regulator SMU.1548c -2.4 sensor histidine kinase SMU.576 -2.3 response regulator SMU.660 -2.1 histidine kinase phosphotransferase system, trehalose-specific IIBC SMU.2038 -2.4 component (EIIBC-tre) SMU.115 -2.1 PTS system, fructose-specific IIA component arginine transcriptional repressor (arginine hydroxymate SMU.2097 -2.3 resistance protein) SMU.292 -2.1 transcriptional regulator, probable AraC family SMU.105 -2.0 SCR operon transcriptional repressor SMU.2134 -2.1 transcriptional regulator, TetR/AcrR family transcriptional regulator, LysR family; probable SMU.1225 -2.1 metCcysK operon transcriptional activator

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Fatty acid and phospholipid metabolism: SMU.1743 -2.5 acyl carrier protein phosphatidate cytidylyltransferase (CDP-diglyceride SMU.1785 -2.1 synthase) SMU.1744 -2.3 3-oxoacyl-[acyl-carrier-protein] synthase III SMU.624 -2.2 1-acylglycerol-3-phosphate O-acyltransferase SMU.1417c -2.1 acyl-ACP thioesterase SMU.988 -2.0 cardiolipin synthase

Energy metabolism: SMU.671 3.3 citrate synthase SMU.672 2.9 isocitrate dehydrogenase SMU.670 2.6 aconitate hydratase; aconitase A SMU.646 2.2 hydrolase (possible phosphoglycolate phosphatase) SMU.352 2.4 ribulose-phosphate-3-epimerase SMU.1978 4.9 acetate kinase SMU.2042 -2.2 dextranase ( 1,6-alpha-glucanhydrolase ) SMU.2047 -2.1 PTS system, enzyme II, A component SMU.1496 -2.1 galactose-6-phosphate isomerase SMU.2065 -2.0 UDP-glucose 4-epimerase SMU.1278c -2.0 phosphoglycolate phosphatase SMU.1123 -2.2 deoxyribose-phosphate aldolase SMU.291 -2.1 transketolase SMU.942 -2.0 3-hydroxy-3-methylglutaryl-CoA reductase SMU.1104c -2.1 phosphoglycerate mutase-like protein SMU.1191 -2.5 6-phosphofructokinase SMU.700c -2.3 phosphoglycerate mutase SMU.1190 -2.2 pyruvate kinase SMU.1010 -2.0 citrate lyase synthetase SMU.1420 -2.3 NADPH-quinone reductase SMU.1971c -2.1 thioredoxin family protein glycosyl hydrolases family 8 protein; possible beta- SMU.1432c -2.6 glucanase SMU.2037 -2.2 trehalose-6-phosphate hydrolase

Central intermediary metabolism: SMU.1180 -2.2 alkylphosphonate uptake protein SMU.318 -2.3 peptidase/aminoacylase/hippurate hydrolase SMU.1595 -2.3 carbonic anhydrase (carbonate dehydratase) SMU.1635 -2.0 UDP-N-acetylglucosamine pyrophosphorylase SMU.939 -2.3 isopentenyl-diphosphate delta-isomerase SMU.943c -2.0 hydroxymethylglutaryl-CoA synthase SMU.1322 2.3 acetoin reductase

112

N-acetyl-gamma-glutamyl-phosphate reductase (N- acetyl-glutamate-gamma-semialdehyde SMU.663 2.7 dehydrogenase) SMU.1437 -2.3 UDP-N-acetylglucosamine 2-epimerase

Purines, pyrimidines, nucleosides, and nucleotides: purine operon repressor/xanthine SMU.356 2.1 phosphoribosyltransferase SMU.1050 -2.0 prs; phosphoribosyl pyrophosphate synthetase purH; phosphoribosylaminoimidazolecarboxamide SMU.37 -3.8 formyltransferase / IMP cyclohydrolase SMU.32 -4.6 purB; amidophosphoribosyltransferase SMU.59 -5.7 asl; adenylosuccinate lyase purE; phosphoribosylaminoimidazole carboxylase SMU.50 -5.6 catalytic subunit purM; phosphoribosylformylglycinamide cyclo-ligase SMU.34 -5.0 (AIRS) purK; phosphoribosylaminoimidazole carboxylase, SMU.51 -5.8 ATPase subunit SMU.35 -5.2 purN; phosphoribosylglycinamide formyltransferase SMU.48 -6.5 purD; phosphoribosylamine-glycine ligase SMU.30 -6.9 purL; phosphoribosylformylglycinamidine synthase purC; phosphoribosylaminoimidazole- SMU.29 -9.6 succinocarboxamide synthase SMU.1054 10.1 guaA; glutamine amidotransferase SMU.595 -2.4 dihydroorotate dehydrogenase (dihydroorotate oxidase) SMU.1223 -2.1 dihydroorotate dehydrogenase SMU.1050 -2.0 phosphoribosyl pyrophosphate synthetase SMU.1066 -2.1 GMP synthase SMU.2157 -2.1 inosine-5'- monophosphate dehydrogenase SMU.1122 -2.6 cytidine deaminase SMU.1215 -2.3 uracil-DNA glycosylase adenylate kinase (ATP-AMP transphosphorylase) SMU.2005 -2.1 (superoxide-inducible protein 16) SMU.1386 -2.0 uridine kinase SMU.668c -2.6 ribonucleotide reductase, large subunit SMU.667 -2.2 ribonucleotide reductase, small subunit SMU.2074 -2.1 anaerobic ribonucleoside-triphosphate reductase

Amino acid biosynthesis: SMU.54 -6.3 amino acid racemase chorismate mutase; possible prephenate SMU.531 -2.4 dehydrogenase SMU.538 2.2 tryptophan synthase, alpha subunit SMU.1312 -2.3 aspartate aminotransferase

113

SMU.54 -6.3 amino acid racemase SMU.70 -2.1 threonine synthase SMU.989 -2.1 aspartate-semialdehyde dehydrogenase SMU.449 -2.5 gamma-glutamyl kinase SMU.450 -2.4 gamma-glutamyl phosphate reductase phosphoribosyl-AMP cyclohydrolase/phosphoribosyl- SMU.1263 -2.3 ATP pyrophosphatase amidotransferase; possible imidazoleglycerol-phosphate SMU.1266 -2.4 synthase SMU.1268 -2.7 imidazoleglycerol-phosphate dehydratase SMU.1270 -2.4 histidinol dehydrogenase SMU.1271 -2.9 ATP phosphoribosyltransferase SMU.1273 -2.3 histidinol-phosphate aminotransferase SMU.233 -2.4 ketol-acid reductoisomerase SMU.231 -2.7 acetolactate synthase, large subunit (AHAS) SMU.232 -2.6 acetolactate synthase, small subunit SMU.1269 -2.4 phosphoserine phosphatase SMU.965 -2.0 homoserine dehydrogenase SMU.966 -2.0 homoserine kinase phosphoribosylformimino-5-aminoimidazole SMU.1265 -2.0 carboxamide ribotide isomerase SMU.1264 -2.0 cyclase HisF/ imidazoleglycerol-phosphate synthase SMU.536 2.0 phosphoribosylanthranilate isomerase

Biosynthesis of cofactors, prosthetic groups, and carriers: SMU.1827 -3.1 biotin biosynthesis protein bifunctional protein: folylpolyglutamate SMU.967 -2.1 synthase/dihydrofolate synthase SMU.970 -2.3 dihydroneopterin aldolase SMU.838 4.1 glutathione reductase ferrochelatase (heme synthetase) (protoheme ferro- SMU.2063 -2.2 lyase) SMU.917c -6.6 6-pyruvoyl tetrahydropterin synthase, PTPS SMU.954 -2.1 pyridoxal kinase SMU.85 -2.5 phosphomethylpyrimidine kinase

Cell envelope, biosynthesis and degradation of murein sacculus and peptidoglycan: SMU.1572 -2.1 UDP-N-acetylglucosamine 1-carboxyvinyltransferase SMU.20 -2.4 cell shape-determining protein MreC D-alanyl-D-alanine carboxypeptidase; penicillin-binding SMU.253 -2.1 protein SMU.599 -2.1 D-alanine-D-alanine ligase SMU.707c -2.9 endolysin SMU.1786 -2.1 undecaprenyl pyrophosphate synthetase

114

Adaptation to atypical conditions and detoxification: SMU.1059 -2.1 acid tolerance protein SMU.669c -3.6 glutaredoxin SMU.919c -7.6 ATPase, confers aluminum resistance SMU.1286c -2.1 multidrug resistance permease SMU.2109 -2.2 possible multiple drug efflux pump MepA SMU.745 -2.8 drug-export protein; multidrug resistance protein undecaprenyl-diphosphatase; bacitracin resistance SMU.244 -2.7 protein SMU.1057 -2.0 acid tolerance protein

Protein synthesis and fate: SMU.1477 -2.8 tRNA isopentenylpyrophosphate transferase SMU.788 -2.6 RNA methyltransferase, TrmA family SMU.1821c -2.4 glutamyl-tRNA (Gln) amidotransferase subunit C SMU.187c -2.3 probable tRNA-dihydrouridine synthase SMU.1820c -2.3 glutamyl-tRNA(Gln) amidotransferase subunit A SMU.1819 -2.3 glutamyl-tRNA (Gln) amidotransferase subunit B SMU.868 -2.1 tRNA (guanine-N1)-methyltransferase SMU.1139c -2.1 rRNA methyltransferase, NOL1/NOP2/sun family SMU.1707c -2.1 spoU-related rRNA/tRNA methylase SMU.1950 -2.0 ribosomal large subunit pseudouridine synthase D SMU.1510 -3.2 phenylalanyl-tRNA synthetase, beta subunit SMU.650 -3.1 alanyl-tRNA synthetase (alanine--tRNA ligase) SMU.1822 -3.0 aspartyl-tRNA synthetase SMU.773c -2.5 lysyl-tRNA synthetase SMU.558 -2.4 isoleucine-tRNA synthetase SMU.1272 -2.4 histidyl-tRNA synthetase SMU.1311 -2.4 asparaginyl-tRNA synthetase SMU.1512 -2.3 phenylalanyl-tRNA synthetase, alpha subunit SMU.1992 -2.2 tyrosyl-tRNA synthetase SMU.1770 -2.2 valyl-tRNA synthetase SMU.2158c -2.2 tryptophanyl-tRNA synthetase SMU.445 -2.2 glycyl-tRNA synthetase alpha subunit SMU.158 -2.1 cysteinyl-tRNA synthetase SMU.1943 2.1 leucyl-tRNA synthetase peptide chain release factor 3 ; translation elongation SMU.608 -2.8 and release factor SMU.2004 -2.2 translation initiation factor IF-1 SMU.697 -2.1 translation initiation factor IF-3 SMU.1326 -2.1 peptide chain release factor SMU.2031 -2.0 translation elongation factor Ts

115

SMU.2022 -2.8 50S ribosomal protein L22 SMU.957 -2.6 50S ribosomal protein L10 SMU.2021 -2.6 30S ribosomal protein S3 SMU.2020 -2.6 50S ribosomal protein L16 SMU.2003 -2.6 30S ribosomal protein S13, N-terminal fragment SMU.2024c -2.6 50S ribosomal protein L4, N-terminal fragment SMU.2025 -2.6 50S ribosomal protein L3 SMU.2002 -2.5 30S ribosomal protein S11 SMU.960 -2.5 50S ribosomal protein L7/L12 SMU.169 -2.5 50S ribosomal protein L13 SMU.2019 -2.4 50s ribosomal protein L29 SMU.2009 -2.4 30S ribosomal protein S5 SMU.2017 -2.4 50S ribosomal protein L14 SMU.2012 -2.4 30S ribosomal protein S8 SMU.2026c -2.4 30S ribosomal protein S10 fragment SMU.358 -2.3 30S ribosomal protein S7 SMU.2016 -2.3 50S ribosomal protein L24 SMU.1200 -2.3 30S ribosomal protein S1 SMU.2014 -2.3 30S ribosomal protein S14 SMU.2007 -2.3 50S ribosomal protein L15, N-terminal fragment SMU.2015 -2.3 50S ribosomal protein L5 SMU.1610 -2.2 50S ribosomal protein L33 SMU.120 -2.2 50S ribosomal protein L28 SMU.2008 -2.2 50S ribosomal protein L30 SMU.849 -2.2 50S ribosomal protein L27 SMU.698 -2.2 50S ribosomal protein L35 SMU.1626 -2.2 50S ribosomal protein L1 SMU.170 -2.2 30S ribosomal protein S9 SMU.2003a -2.2 50S ribosomal protein L36 SMU.1860 -2.1 30S ribosomal protein S6 SMU.2018 -2.1 30S ribosomal protein S17 SMU.2104a -2.1 50S ribosomal protein L32 SMU.1288 -2.1 50S ribosomal protein L19 SMU.2023c -2.1 30S ribosomal protein S19, C-terminal fragment SMU.1627 -2.0 50S ribosomal protein L11 SMU.357 -2.0 30S ribosomal protein S12 SMU.2011 -2.0 50S ribosomal protein L6 (BL10) SMU.818 -2.0 30S ribosomal protein S21 SMU.2000 -2.0 50S ribosomal protein L17 SMU.2032 -2.0 30S ribosomal protein S2 SMU.340 -2.0 50S ribosomal protein L34 SMU.500 2.0 ribosome-associated protein

116

aminotransferase (class V); possible iron-sulfur cofactor synthesis protein;pyridoxal-phosphate dependent SMU.1051 2.2 aminotransferase SMU.143c -2.1 polypeptide deformylase (PDF) SMU.188c -2.4 Hsp33-like chaperonin SMU.1787c -3.1 preprotein translocase, YajC subunit SMU.1973 -2.5 glutamyl aminopeptidase SMU.1592 -2.2 proline dipeptidase SMU.1030 -2.3 polyribonucleotide nucleotidyltransferase tRNA: SMU.t01 -2.8 SMU.t02 -4.6 SMU.t03 -2.9 SMU.t04 -2.9 SMU.t05 -2.1 SMU.t06 -2.1 SMU.t08 -2.4 SMU.t09 -2.9 SMU.t10 -2.3 SMU.t11 -2.2 SMU.t12 -4.1 SMU.t13 -2.2 SMU.t14 -2.6 SMU.t15 -2.5 SMU.t16 -4.0 SMU.t17 -3.0 SMU.t21 -2.4 SMU.t22 -3.1 SMU.t23 -2.2 SMU.t24 -2.4 SMU.t26 -2.4 SMU.t27 -3.1 SMU.t28 -2.3 SMU.t30 -3.3 SMU.t31 -2.6 SMU.t32 -2.5 SMU.t34 -2.2 SMU.t36 -2.8 SMU.t38 -2.6 SMU.t39 -3.2 SMU.t41 -3.1 SMU.t42 -3.4

117

SMU.t43 -2.1 SMU.t44 -2.0 SMU.t45 -2.4 SMU.t46 -2.4 SMU.t47 -2.9 SMU.t48 -3.1 SMU.t49 -3.1 SMU.t50 -2.6 SMU.t52 -2.3 SMU.t53 -2.8 SMU.t55 -2.7 SMU.t56 -2.6 SMU.t57 -3.0 SMU.t58 -2.4 SMU.t59 -2.2 SMU.t60 -2.7 SMU.t61 -3.4 SMU.t64 -2.7 SMU_t35 -2.0 SMU_t40 -2.0 SMU_t54 -2.0

Other: SMU.837 7.2 oxidoreductase, aldo/keto reductase family ythI ytxK; conserved hypothetical protein, SMU.1979c 6.5 methyltransferase domain SMU.1982c 6.0 conserved hypothetical protein SMU.769 5.6 conserved hypothetical protein SMU.1980c 5.6 conserved hypothetical protein argC; N-acetyl-gamma-glutamyl-phosphate reductase SMU.663 2.7 (N-acetyl-glutamate-gamma-semialdehyde dehydrogenase) SMU.508 3.2 conserved hypothetical protein (possible hydrolase) SMU_847c -2.4 hypothetical protein SMU.862 4.1 conserved hypothetical protein SMU.2076c 2.9 hypothetical protein SMU.2081 2.3 hypothetical protein SMU.355 2.5 CMP-binding factor 1 SMU.65 2.4 protein tyrosine-phosphatase SMU.1322 2.3 acetoin reductase SMU.1374 -2.1 IS30 transposase related protein SMU.329 2.2 conserved hypothetical protein SMU.807 2.2 conserved hypothetical protein (possible membrane

118

protein) SMU.673 2.1 conserved hypothetical protein SMU.354 2.0 conserved hypothetical protein SMU.67 2.0 acyltransferase SMU.2048 -2.3 hypothetical protein SMU.1395c -2.3 hypothetical protein SMU.222c -2.3 hypothetical protein (possible integrase fragment) SMU.223c -2.6 hypothetical protein SMU.224c -2.7 hypothetical protein SMU.225c -2.3 hypothetical protein SMU.277 -4.1 hypothetical protein SMU.278 -3.7 hypothetical protein SMU.279 -4.0 hypothetical protein SMU.281 -4.3 hypothetical protein SMU.283 -4.9 hypothetical protein SMU.284 -3.0 hypothetical protein SMU.285 -4.4 hypothetical protein SMU_429c -2.2 hypothetical protein SMU.600c -2.9 conserved hypothetical protein SMU.604 -2.4 conserved hypothetical protein SMU.621c -2.3 conserved hypothetical protein SMU.622c -2.5 conserved hypothetical protein SMU.958 -3.3 hypothetical protein SMU.959c -2.6 hypothetical protein SMU.1435c -2.7 hypothetical protein SMU.444 -2.5 hypothetical protein SMU.451 -3.2 hypothetical protein SMU.2033c -3.3 conserved hypothetical protein SMU.709 -3.0 conserved hypothetical protein SMU.703c -2.4 conserved hypothetical protein SMU.706c -2.1 conserved hypothetical protein SMU.711 -3.1 conserved hypothetical protein SMU.649 -3.4 conserved hypothetical protein SMU.1753c -3.5 conserved hypothetical protein SMU.1754c -4.6 conserved hypothetical protein SMU.1755c -4.1 conserved hypothetical protein SMU.1757c -4.3 conserved hypothetical protein SMU.1758c -4.0 conserved hypothetical protein SMU.1760c -4.0 conserved hypothetical protein SMU.1761c -4.2 conserved hypothetical protein SMU.1762c -3.5 conserved hypothetical protein SMU.1763c -3.5 conserved hypothetical protein

119

SMU.1764c -3.9 conserved hypothetical protein SMU.1262c -2.1 hypothetical protein SMU.1267c -2.5 hypothetical protein SMU.1310 -2.4 hypothetical protein SMU.1316c -3.0 hypothetical protein SMU.1317c -3.4 hypothetical protein SMU.940c -3.5 hemolysin III-related protein SMU.987 -3.0 cell wall surface anchor family protein SMU.1750c -3.7 hypothetical protein SMU.1752c -2.6 hypothetical protein SMU.1628 -2.2 conserved hypothetical protein SMU.1623c -2.1 conserved hypothetical protein SMU.1373c -2.8 hypothetical protein SMU.73 -4.0 conserved hypothetical protein SMU.1896c -4.1 hypothetical protein SMU.1895c -3.5 hypothetical protein SMU.1861c -2.0 hypothetical protein SMU.1794c -2.1 hypothetical protein SMU.18 -2.2 hypothetical protein SMU.1804c -2.2 hypothetical protein SMU.1813 -2.9 hypothetical protein, probable transposase fragment SMU.1818c -4.3 hypothetical protein SMU.1000 -4.6 hypothetical protein SMU.1373c -2.8 hypothetical protein SMU.1395c -2.3 hypothetical protein SMU.1552c -2.0 hypothetical protein SMU.1553c -2.7 hypothetical protein SMU.1587c -2.5 hypothetical protein SMU.1648c -2.2 hypothetical protein SMU.1766c -2.1 hypothetical protein SMU.1768c -2.1 hypothetical protein SMU.1946 -2.1 conserved hypothetical protein SMU.2064c -2.5 hypothetical protein SMU.2105 -2.1 hypothetical protein SMU.2106c -2.1 hypothetical protein SMU.2111c -2.2 hypothetical protein SMU.33 -5.8 hypothetical protein SMU.344 -2.2 hypothetical protein SMU.350 -2.4 hypothetical protein SMU.49 -6.9 hypothetical protein SMU.501 -4.3 hypothetical protein SMU.503c -2.2 hypothetical protein

120

SMU.545 -2.3 hypothetical protein SMU.55 -5.1 hypothetical protein SMU.58 -5.4 hypothetical protein SMU.594 -2.0 hypothetical protein SMU.605 -2.3 hypothetical protein SMU.606 -2.2 hypothetical protein SMU.620 -2.6 hypothetical protein SMU.722 -2.6 hypothetical protein SMU.748 -2.0 hypothetical protein SMU.771c 4.1 hypothetical protein SMU.791c -2.5 hypothetical protein SMU.999 -2.3 hypothetical protein SMU.45 -2.0 hypothetical protein SMU.1399 -2.0 hypothetical protein SMU.295 -2.0 hypothetical protein SMU.1771c -2.0 hypothetical protein SMU.1907 21.1 hypothetical protein SMU.1073 -7.5 formate--tetrahydrofolate ligase SMU.916c -7.6 conserved hypothetical protein SMU.914c -7.5 conserved hypothetical protein SMU.56 -6.2 Streptococcus-specific protein SMU.52 -5.8 conserved hypothetical protein SMU.31 -5.7 conserved hypothetical protein SMU.53 -5.5 conserved hypothetical protein SMU.915c -4.9 conserved hypothetical protein SMU.941c -4.8 conserved hypothetical protein SMU.2146c -4.6 conserved hypothetical protein SMU.72 -4.3 conserved hypothetical protein SMU.1545c -4.1 conserved hypothetical protein SMU.502 -4.1 conserved hypothetical protein SMU.36 -3.7 conserved hypothetical protein (eukaryotic-like) SMU.1876 -2.9 conserved hypothetical protein SMU.530c -2.8 conserved hypothetical protein SMU.721 -2.8 conserved hypothetical protein SMU.1026 -2.8 conserved hypothetical protein SMU.1701c -2.7 conserved hypothetical protein SMU.1377c -2.6 conserved hypothetical protein SMU.434 -2.6 conserved hypothetical protein SMU.1436c -2.6 conserved hypothetical protein SMU.720 -2.5 conserved hypothetical protein SMU.547 -2.5 conserved hypothetical protein SMU.1582c -2.5 conserved hypothetical protein

121

SMU.428 -2.5 conserved hypothetical protein, Cof family SMU.1482c -2.5 conserved hypothetical protein SMU.1211 -2.4 conserved hypothetical protein SMU.898 -2.4 conserved hypothetical protein conserved hypothetical protein; possible membrane SMU.2059c -2.4 protein SMU.1414c -2.4 conserved hypothetical protein SMU.227c -2.4 conserved hypothetical protein conserved hypothetical protein (possible integral SMU.2066c -2.4 membrane protein, ABC transporter) SMU.1484c -2.4 conserved hypothetical protein SMU.1375c -2.4 conserved hypothetical protein conserved hypothetical protein (phenylalanyl-tRNA SMU.1505c -2.3 synthetase fragment) SMU.1853 -2.3 conserved hypothetical protein conserved hypothetical protein (possible SMU.1483c -2.3 acetyltransferase) SMU.243 -2.3 conserved hypothetical protein SMU.343 -2.3 conserved hypothetical protein SMU.1314 -2.3 conserved hypothetical protein SMU.1321c -2.3 conserved hypothetical protein SMU.543 -2.3 conserved hypothetical protein SMU.1319c -2.3 conserved hypothetical protein SMU.471 -2.3 conserved hypothetical protein SMU.1703c -2.3 conserved hypothetical protein SMU.1871c -2.2 conserved hypothetical protein SMU.2162c -2.2 conserved hypothetical protein SMU.1284c -2.2 conserved hypothetical protein SMU.2147c -2.2 conserved hypothetical protein SMU.1968c -2.2 conserved hypothetical protein SMU.239c -2.2 conserved hypothetical protein SMU.521 -2.1 conserved hypothetical protein SMU.393 -2.1 conserved hypothetical protein conserved hypothetical protein (possible SMU.341 -2.1 deoxyribonuclease) SMU.523 -2.1 conserved hypothetical protein, VanZ-like family conserved hypothetical protein (possible alpha / beta SMU.737 -2.1 superfamily hydrolase) SMU.156 -2.1 conserved hypothetical protein SMU.1773c -2.1 conserved hypothetical protein SMU.948 -2.1 conserved hypothetical protein SMU.848 -2.1 conserved hypothetical protein conserved hypothetical protein (possible SMU.1406c -2.0 oxidoreductase)

122

SMU.1290c -2.0 conserved hypothetical protein SMU.1109c -2.0 conserved hypothetical protein SMU.159 -2.0 conserved hypothetical protein SMU.1546 -2.0 conserved hypothetical protein conserved hypothetical protein (possible myo-inositol- SMU.1140c -2.0 1(or 4)-monophosphatase) SMU.1704 -2.0 conserved hypothetical protein SMU.392c -2.0 conserved hypothetical protein, HI0933-like SMU.1036 -2.0 conserved hypothetical protein SMU.1171c -2.0 conserved hypothetical protein conserved hypothetical protein; possible SMU.1392c -2.0 acetyltransferase Permease-ABC-type antimicrobial peptide transport SMU.1347c -2.0 system, Permease component SMU.333 -2.0 conserved hypothetical protein SMU.125 -2.0 conserved hypothetical protein SMU.1153c -2.0 conserved hypothetical protein SMU.1784c -2.0 conserved hypothetical protein, peptidase family M50 SMU.497c -2.0 conserved hypothetical protein SMU.475 -2.0 conserved hypothetical protein conserved hypothetical protein (possible transcriptional SMU.751 -2.0 regulator) SMU.407 -2.0 conserved hypothetical protein SMU.1052 2.1 conserved hypothetical protein SMU.1196c 2.8 conserved hypothetical protein SMU.63c 3.3 conserved hypothetical protein SMU.1072c -7.6 acyltransferase SMU.1450 -3.0 amino acid permease SMU.1511c -2.7 acetyltransferase; possible transcriptional repressor GTP-binding protein (tyrosine phosphorylated protein SMU.546 -2.7 A); possible elongation factor SMU.1434c -2.6 glycosyl transferase SMU.701c -2.5 integral membrane protein SMU.623c -2.5 polysaccharide deacetylase SMU.1061 -2.4 DNA-binding protein SMU.1700c -2.3 LrgB-like protein; possible murein hydrolase regulator SMU.1100c -2.3 permease SMU.2043c -2.2 D-tyrosyl-tRNA deacylase metallo-beta-lactamase superfamily proteinhetical SMU.1444c -2.2 protein SMU.368c -2.2 metallo-beta-lactamase superfamily protein SMU.348 -2.1 histidine triad (HIT) hydrolase SMU.1558c -2.1 acetyltransferase, GNAT family SMU.1969c -2.1 probable transcriptional regulator

123

SMU.1555c -2.0 ribonuclease BN-like family protein SMU.337 -2.0 inner membrane protein uncharacterized phosphatase; possible PAP2 family SMU.1702c -2.0 protein SMU.1029 -9.2 hypothetical - transposon SMU.189 -2.6 hypothetical protein

124

Table S5.2. Genes up- or down-regulated in ∆SMcomS in response to 1 µM XIP.

Gene ID Fold Change Annotation

Cell killing and bacteriocin production: SMU.1910c 4.6 hypothetical protein SMU.1912c 4.0 hypothetical protein SMU.1913c 3.0 putative immunity protein; BlpL-like; immA SMU.1914c 3.1 hypothetical protein; nlmC; mutacin V SMU.1908c 4.2 hypothetical protein; immunity protein SMU.1909c 5.1 hypothetical protein SMU.1902c 6.5 hypothetical protein SMU.1903c 6.0 putative bacteriocin secretion protein; bsmL SMU.1904c 5.5 hypothetical protein SMU.1905c 5.0 hypothetical protein SMU.1906c 6.5 bacteriocin-related protein SMU.150 2.0 nlmA; non-lantibiotic mutacin IV A SMU.151 2.1 nlmB; non-lantibiotic mutacin IV B SMU.152 2.3 hypothetical protein SMU.423 4.0 nlmD; possible bacteriocin SMU.836 71.0 hypothetical protein SMU.299c 3.4 ip; bacteriocin peptide precursor SMU.1917 7.8 comE; response regulator SMU.1916 8.9 comD; histidine kinase SMU.1915 2.2 comC, competence stimulating peptide SMU.168 5.9 transcriptional regulator toxin-antitoxin SMU.166 5.4 putative toxin-antitoxin system SMU.167 6.4 putative toxin-antitoxin system

Competence and DNA transformation: SMU.1001 525.1 dprA; DNA processing protein; Smf family SMU.1981c 936.7 competence protein comG SMU.1983 408.9 competence protein comYD SMU.1984 1185.2 competence protein comYC SMU.1997 61.5 comX; competence-specific sigma factor SMU.498 351.0 comFA; late competence protein F SMU.625 183.8 comEA; competence protein SMU.626 394.6 comEC; competence protein SMU.499 277.3 comFC; late competence protein SMU.644 216.0 coiA; competence protein SMU.1987 625.8 comGA; late competence protein SMU.2086 10.0 cinA; competence damage-inducible protein A SMU.1985 783.0 comGB; competence protein

125

DNA metabolism, replication, recombination and repair: SMU.02 2.4 dnaN; DNA polymerase III, beta subunit SMU.1002 8.9 topA; DNA topoisomerase I SMU.1055 27.5 radC; DNA repair protein SMU.1056 3.4 DNA repair protein RadC SMU.1174 2.8 pcrA; ATP-dependent DNA helicase SMU.1192 2.4 dnaE; DNA-polymerase III subunit alpha SMU.123 2.2 polC; DNA-polymerase III, alpha subunit SMU.1472 3.0 recJ; single-strand DNA-specific exonuclease SMU.1500 2.0 rexB; ATP-dependent exonuclease subunit B SMU.1714c 2.0 xerD; tyrosine recombinase SMU.1967 215.1 ssbA; single-stranded DNA-binding protein SMU.2085 6.5 recA; recombinase A SMU.505 9.4 dpn; adenine-specific DNA methylase (DpnIIB) SMU.506 7.4 ssuRB; type II restriction endonuclease SMU.64 12.0 ruvB; holiday junction DNA helicase SMU.821 2.1 dnaG; DNA primase SMU.510c -2.5 deoxyribonuclease

Transport and binding: lantibiotic related antibiotic efflux protein/macrolide SMU.109 2.2 permease SMU.1658 -3.5 ammonium transporter, NrgA protein manganese transporter/possible HitA ferric iron-binding SMU.770c 8.0 periplasmic protein SMU.1013c 2.3 Mg2+/citrate complex transporter SMU.980 3.2 beta-glucoside-specific EII permease SMU.772 8.9 bifunctional glucan-binding protein D and lipase SMU.263 2.8 amino acid permease /putrescine antiporter SMU.1568 -3.8 maltose / maltodextrin-binding protein spermidine/putrescine ABC transporter, SMU.976 1.9 spermidine/putrescine-binding protein SMU.1900 2.1 ABC transporter SMU.998 2.4 ABC transporter, ferrichrome-binding protein amino acid ABC transporter, amino acid substrate- SMU.933 2.8 binding protein SMU.1963c 3.5 sugar-binding periplasmic protein SMU.1966c 4.3 levT, ABC transport ribose-binding protein, periplasmic maltose / maltodextrin ABC transport system SMU.1570 -3.1 (permease) SMU.1569 -2.7 maltodextrin ABC transport system permease SMU.2160 2.0 possible permease, ABC transporter protein SMU.909 2.0 malate permease

126

SMU.1093 2.1 ABC transporter permease protein SMU.934 2.1 amino acid ABC transporter, permease protein SMU.183 2.2 manganese ABC transporter permease element SMU.1149 2.2 ABC transporter, membrane spanning ABC transporter, permease, possibly bacteriocin SMU.657 2.3 associated SMU.864 2.4 ABC transporter, permease component ABC transporter, permease protein;possible ferrichrome SMU.996 2.4 transport system SMU.1928 2.4 protein secretion ABC transport permease SMU.975 2.4 spermidine/putrescine ABC transporter, permease ABC transporter, permease protein (possible taurine SMU.653c 2.5 transport system permease) SMU.1067c 2.6 ABC transporter, permease protein SMU.906 2.9 ABC transporter, ATP-binding / permease protein ABC transporter, permease, possibly bacteriocin SMU.656 3.3 associated SMU.1166c 3.4 ABC transporter permease ABC-type multidrug/protein/lipid transport system, SMU.905 2.8 ATPase component glycine betaine / carnitine / choline ABC transporter, SMU.2116 -2.1 ATP-binding protein, opuCA SMU.654 2.0 ABC transporter, ATP-binding protein SMU.1551c 2.0 ABC transporter, ATP-binding protein inorganic ion ABC transporter,ATP-binding protein; SMU.997 2.2 possible ferrichrome transport system SMU.1068c 2.2 ABC transporter, ATP-binding protein SMU.1041 2.4 ABC transporter, ATP-binding protein SMU.863 2.4 ABC transporter, ATP-binding protein SMU.1927 2.8 PsaA protein/ ABC transporter, ATP-binding protein SMU.1148 2.8 ABC transporter, ATPase component SMU.1167c 3.1 ABC transporter ATP-binding protein SMU.1899 4.4 ABC transport fragment

Signal transduction and transcriptional regulation: SMU.1926 2.0 transcriptional regulator PsaR SMU.1168 2.0 transcriptional regulator (TetR/AcrR family) SMU.977 2.6 transcription antiterminator LicT (fragment) SMU.1977c 2.8 transcriptional regulator SMU.1995c 3.1 zinc transport transcriptional repressor SMU.261c 3.4 transcriptional regulator SMU.592c 3.9 transcriptional regulator SMU.507 4.9 transcriptional regulator, DeoR family SMU.514 6.6 transcriptional regulator, AcrR family

127

SMU.1733c 2.1 SNF helicase SMU.1474c 2.8 ribonuclease Z SMU.994 2.2 ribonuclease HII SMU.1517 2.0 vicR, two-component response regulator SMU.1516 2.0 vicK, two-component sensor histidine kinase SMU.1964c 3.9 levR, two-component response regulator SMU.1965c 4.1 levS, histidine kinase SMU.1877 -2.0 manL, mannose PTS system component IIAB phosphotransferase system phosphohistidine-containing SMU.674 -2.8 protein PTS system IIB component, required for cellobiose SMU.1600 -2.2 uptake and metabolism SMU.1958c 2.8 levF, fructose-specific Enzyme IIC component SMU.1960c 3.0 levE, fructose-specific Enzyme IIB component SMU.1957 3.3 levG, fructose-specific Enzyme IID component SMU.1961c 3.4 levD, fructose-specific Enzyme IIA component SMU.65 11.7 protein tyrosine-phosphatase vicX, gtfB/C regulator, metallo-beta-lactamase SMU.1515 2.4 superfamily SMU.1053 10.5 possible redox-sensing transcriptional repressor Rex SMU.1919 2.3 sakacin A production response regulator glnR, transcriptional regulator; glutamine synthetase SMU.363 -2.8 repressor

Energy metabolism: SMU.671 -4.2 citrate synthase SMU.670 -4.0 aconitate hydratase; aconitase A SMU.672 -3.9 isocitrate dehydrogenase SMU.1495 2.0 galactose-6-phosphate isomerase SMU.981 3.0 beta-glucosidase, BglB protein SMU.2028 4.2 fructosyltransferase SMU.982 4.2 beta-glucosidase, BglB protein SMU.646 13.6 hydrolase (possible phosphoglycolate phosphatase) sugar-phosphate isomerase (ribose 5-phosphate SMU.2142 3.3 isomerase) SMU.352 4.0 ribulose-phosphate-3-epimerase SMU.1170 2.3 cytochrome c-type biogenesis protein SMU.1043c 2.0 phosphate acetyltransferase SMU.1422 2.1 acetoin dehydrogenase E1 component SMU.1423 2.8 acetoin dehydrogenase E1 component dihydrolipoamide acetyltransferase (acetoin SMU.1421 2.9 dehydrogenase E2 component) SMU.1424 3.7 dihydrolipoamide dehydrogenase SMU.1978 17.9 acetate kinase

128

SMU.665 2.5 acetylglutamate kinase

Central intermediary metabolism: SMU.132 2.2 amino acid amidohydrolase (hippurate amidohydrolase) SMU.1322 4.2 acetoin reductase SMU.664 3.0 ornithine acetyltransferase / N-acetylglutamate synthase N-acetyl-gamma-glutamyl-phosphate reductase (N- acetyl-glutamate-gamma-semialdehyde SMU.663 3.2 dehydrogenase) SMU.264 2.0 agmatine deiminase

Fatty acid and phospholipid metabolism: enoyl-acyl carrier protein(ACP) reductase; dioxygenase SMU.1335c 2.1 related to 2-nitropropane dioxygenase

Cell division: SMU.1713c 2.0 segregation and condensation protein A SMU.1003 10.2 glucose-inhibited division protein

Pathogenesis, toxin production and resistance: SMU.515 7.3 mycA; 67 kDa myosin-crossreactive antigen SMU.610 -6.0 spaP; cell surface antigen SMU.1340 2.0 bacA; bacitracin synthetase 1/ tyrocidin synthetase III

Adaptation to atypical conditions and detoxification: SMU.1046c 2.8 relQ; relA yjbM; GTP pyrophosphokinase SMU.1286c 2.9 multidrug resistance permease SMU.1338c 2.2 mefE; ABC transport macrolide permease

Mobile and extrachromosomal element functions: SMU.1398 2.4 irvR; repressor protein - phage associated SMU.1329c 2.4 paaB; transposase fragment SMU.1351 2.0 transposase fragment SMU.1372c 2.0 transposase, IS861, IS3 family SMU.149 2.0 transposase fragment (IS605/IS200-like) SMU.590c 4.0 transposase fragment SMU.1024c 2.0 transposase fragment SMU.767 2.6 transposase SMU.1380 3.1 Pseudo-Spn1 transposase SMU.766 4.1 transposase

Purines, pyrimidines, nucleosides, and nucleotides: SMU.1054 20.3 guaA; glutamine amidotransferase

129

purC; phosphoribosylaminoimidazole- SMU.29 -4.2 succinocarboxamide synthase SMU.30 -4.6 purL; phosphoribosylformylglycinamidine synthase SMU.32 -3.7 purB; amidophosphoribosyltransferase purM; phosphoribosylformylglycinamide cyclo-ligase SMU.34 -3.3 (AIRS) SMU.35 -2.4 purN; phosphoribosylglycinamide formyltransferase purH; phosphoribosylaminoimidazolecarboxamide SMU.37 -2.5 formyltransferase / IMP cyclohydrolase SMU.48 -2.6 purD; phosphoribosylamine-glycine ligase purE; phosphoribosylaminoimidazole carboxylase SMU.50 -2.7 catalytic subunit purK; phosphoribosylaminoimidazole carboxylase, SMU.51 -3.0 ATPase subunit purR; purine operon repressor/xanthine SMU.356 4.0 phosphoribosyltransferase

Amino acid biosynthesis: SMU.532 2.6 trpE; anthranilate synthase, component I SMU.533 4.4 trpG; anthranilate synthase, component II SMU.534 2.7 trpD; phosphoribosyl anthranilate transferase SMU.535 4.1 trpC; indole-3-glycerol phosphate synthase SMU.536 6.1 trpF; phosphoribosylanthranilate isomerase SMU.537 6.2 trpB; tryptophan synthase, beta subunit SMU.538 15.8 trpA; tryptophan synthase, alpha subunit SMU.1657c -2.5 glnB; nitrogen regulatory protein PII SMU.364 -2.7 glnA; glutamine synthetase type 1 SMU.1381 2.9 leuD; 3-isopropylmalate dehydratase, small subunit SMU.1382 3.0 leuC; alpha-isopropylmalate isomerase large subunit SMU.1383 2.6 leuB; 3-isopropylmalate dehydrogenase SMU.1384 2.6 leuA; 2-isopropylmalate synthase SMU.1173 2.6 cysD; O-acetylhomoserine sulfhydrylase SMU.1920 2.2 pdgA; phosphoglycerate dehydrogenase SMU.666 2.4 N-acetylornithine aminotransferase SMU.262 4.9 ornithine carbamoyltransferase SMU.54 -2.4 amino acid racemase

Biosynthesis of cofactors, prosthetic groups, and carriers: SMU.838 5.8 gor; glutathione reductase SMU.1045c 2.6 ppnK; NAD(+) kinase (ATP-NAD kinase) SMU.353 4.2 thiamine pyrophosphokinase

Cell envelope, biosynthesis and degradation of murein sacculus and peptidoglycan: SMU.21 2.0 cell shape-determining protein MreD

130

Protein synthesis and protein fate: SMU.1132 2.0 pepN; lysyl-aminopeptidase SMU.539c 45.1 prepilin peptidase type IV SMU.645 15.6 PepB oligopeptidase SMU.1610 3.6 rpmG; 50S ribosomal protein L33 SMU.340 2.6 rpmH; 50S ribosomal protein L34 SMU.500 2.1 yfiA; ribosome-associated protein SMU.1044c 2.6 rluE; ribosomal large subunit pseudouridine synthase trmU; tRNA (5-methylaminomethyl-2-thiouridylate)- SMU.2143c 2.6 methyltransferase SMU.r06 -3.0 SMU.r03 -3.0 SMU.r05 -2.4 SMU.r02 -2.3 tRNA: SMU.t22 -4.6 SMU.t03 -3.6 SMU.t02 -3.4 SMU.t07 -3.0 SMU.t21 -2.9 SMU.t01 -2.9 SMU.t61 -2.8 SMU.t05 -2.3 SMU.t42 2.2 SMU.t30 2.3 SMU.t09 2.4 SMU.t37 2.4 SMU.t59 2.5 SMU.t08 2.6 SMU.t63 3.6

Other: SMU.1073 -2.5 formate--tetrahydrofolate ligase SMU.31 -4.3 conserved hypothetical protein SMU.1403c -2.7 conserved hypothetical protein SMU.1405c -2.6 conserved hypothetical protein SMU.53 -2.5 conserved hypothetical protein SMU.1968c -2.5 conserved hypothetical protein SMU.36 -2.4 conserved hypothetical protein (eukaryotic-like) SMU.673 -2.4 conserved hypothetical protein SMU.502 -2.3 conserved hypothetical protein

131

SMU.1764c -2.3 conserved hypothetical protein SMU.1402c -2.3 conserved hypothetical protein SMU.1803c -2.3 conserved hypothetical protein SMU.1404c -2.1 conserved hypothetical protein SMU.52 -2.1 conserved hypothetical protein SMU.1763c -2.0 conserved hypothetical protein SMU.560c -2.0 conserved hypothetical protein SMU.845 2.0 conserved hypothetical protein conserved hypothetical protein (possible transcription SMU.345c 2.0 regulator) SMU.844 2.0 conserved hypothetical protein SMU.1923c 2.0 conserved hypothetical protein SMU.1956c 2.0 conserved hypothetical protein SMU.1701c 2.0 conserved hypothetical protein conserved hypothetical protein (possible membrane SMU.807 2.1 protein) SMU.746c 2.1 conserved hypothetical protein SMU.1048 2.1 conserved hypothetical protein SMU.523 2.1 conserved hypothetical protein, VanZ-like family conserved hypothetical protein (possible ppGpp- SMU.173 2.1 regulated growth inhibitor) SMU.1080c 2.1 conserved hypothetical protein SMU.1642c 2.1 conserved hypothetical protein SMU.929c 2.2 conserved hypothetical protein conserved hypothetical protein; Streptococcus-specific SMU.217c 2.2 protein SMU.1336 2.2 conserved hypothetical protein SMU.290 2.2 conserved hypothetical protein SMU.627 2.2 conserved hypothetical protein SMU.1442c 2.2 conserved hypothetical protein SMU.473 2.3 conserved hypothetical protein SMU.1678 2.4 conserved hypothetical protein conserved hypothetical protein (possible membrane SMU.835 2.4 protein) SMU.820 2.4 conserved hypothetical protein SMU.1070c 2.4 conserved hypothetical protein SMU.1975c 2.4 conserved hypothetical protein SMU.1172c 2.5 conserved hypothetical protein SMU.757 2.5 conserved hypothetical protein SMU.1856c 2.5 conserved hypothetical protein SMU.1976c 2.6 conserved hypothetical protein conserved hypothetical protein; possible ABC SMU.1042 2.6 transporter, permease SMU.1884c 2.8 conserved hypothetical protein

132

conserved hypothetical protein (possible alpha/beta SMU.1280c 2.9 superfamily hydrolase) conserved hypothetical protein (possible arsenate SMU.2084c 2.9 reductase) SMU.630 3.1 conserved hypothetical protein SMU.1400c 3.1 conserved hypothetical protein SMU.1475c 3.1 conserved hypothetical protein SMU.631 3.2 conserved hypothetical protein SMU.932 3.2 conserved hypothetical protein SMU.1621c 3.2 conserved hypothetical protein SMU.591c 3.2 conserved hypothetical protein SMU.1284c 3.6 conserved hypothetical protein SMU.790 3.7 conserved hypothetical protein SMU.354 4.5 conserved hypothetical protein SMU.508 4.5 conserved hypothetical protein (possible hydrolase) SMU.2080 5.0 brsR, conserved hypothetical protein conserved hypothetical protein; possible SMU.66 6.9 phosphatidylinositol-4-phosphate 5-kinase SMU.63c 8.5 conserved hypothetical protein SMU.769 19.2 conserved hypothetical protein conserved hypothetical protein, methyltransferase SMU.1979c 92.5 domain SMU.1982c 715.7 conserved hypothetical protein SMU.1980c 1618.5 conserved hypothetical protein SMU.1969c -2.6 probable transcriptional regulator SMU.987 -2.0 cell wall surface anchor family protein SMU.679 2.0 oxidoreductase, aldo/keto reductase family SMU.834 2.1 glycosyltransferase SMU.1476c 2.1 GTP-binding protein SMU.1337c 2.1 alpha/beta superfamily hydrolases SMU.641 2.3 oxidoreductase SMU.574c 2.3 effector of murein hydrolase SMU.346 2.4 NADH dehydrogenase; NAD(P)H nitroreductase SMU.1918 2.4 membrane-associated protein DedA SMU.1169c 2.5 thioredoxin family protein isopentenyl monophosphate kinase (4-diphosphocytidyl- SMU.1996 2.5 2-C-methyl-D-erythritol kinase) SMU.993 2.6 GTP-binding protein oxidoreductase, short chain dehydrogenase/reductase SMU.1473c 2.8 family SMU.643 3.3 acetyl esterase/ sugar hydrolase SMU.647 3.4 O-methyltransferase SMU.355 4.9 CMP-binding factor 1 SMU.67 5.8 acyltransferase

133

SMU.837 29.7 oxidoreductase, aldo/keto reductase family SMU.1895c -5.2 hypothetical protein SMU.1896c -4.3 hypothetical protein SMU.18 -4.1 hypothetical protein SMU.1804c -3.8 hypothetical protein SMU.33 -3.6 hypothetical protein SMU.1000 -3.0 hypothetical protein SMU.278 -2.7 hypothetical protein SMU.281 -2.6 hypothetical protein SMU.279 -2.5 hypothetical protein SMU.1752c -2.5 hypothetical protein SMU.503c -2.4 hypothetical protein SMU.501 -2.3 hypothetical protein SMU.49 -2.3 hypothetical protein SMU.284 -2.3 hypothetical protein SMU.958 -2.2 hypothetical protein SMU.55 -2.1 hypothetical protein SMU.277 -2.0 hypothetical protein SMU.1399 -2.0 hypothetical protein SMU.594 -2.0 hypothetical protein SMU.214c 2.0 hypothetical protein SMU.739c 2.0 hypothetical protein SMU.748 2.1 hypothetical protein SMU.1855 2.1 high density responsive membrane protein, HdrM SMU.1369 2.1 hypothetical protein SMU.1256c 2.1 hypothetical protein SMU.813 2.1 hypothetical protein (possible transcriptional regulator) SMU.185 2.1 hypothetical protein SMU.1047c 2.1 hypothetical protein SMU.1504c 2.1 hypothetical protein SMU.216c 2.1 hypothetical protein SMU.811 2.2 hypothetical protein SMU.1310 2.2 hypothetical protein SMU.219 2.2 hypothetical protein SMU.1069c 2.2 hypothetical protein SMU.215c 2.2 hypothetical protein SMU.642 2.2 hypothetical protein SMU.1373c 2.3 hypothetical protein SMU.1891c 2.3 hypothetical protein SMU.176 2.3 hypothetical protein SMU.1862 2.4 hypothetical protein SMU.1091 2.6 hypothetical protein (possible cell wall protein)

134

SMU.220c 2.6 hypothetical protein SMU.379 2.7 hypothetical protein SMU.1553c 2.8 hypothetical protein SMU.992 2.8 hypothetical protein SMU.2083c 3.1 hypothetical protein SMU.791c 3.1 hypothetical protein SMU.812 3.4 hypothetical protein SMU.1368 3.9 hypothetical protein SMU.2081 4.2 brsM, hypothetical protein SMU.513 5.1 hypothetical protein SMU.68 5.5 hypothetical protein SMU.1907 5.6 hypothetical protein SMU.2076c 6.5 hypothetical protein SMU.771c 7.1 hypothetical protein SMU.1378 10.3 hypothetical protein SMU.189 -5.6 hypothetical protein SMU.93c -2.2 hypothetical protein SMU.2053c 2.4 hypothetical protein (peptide) SMU.437c 2.7 Pseudo gene SMU.1379 3.7 hypothetical protein-HTH XRE domain SMU.41 6.0 hypothetical protein

135

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