AMMONIA-GENERATING MECHANISMS OF MUTANS STREPTOCOCCI By ANN RYAN GRISWOLD A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2006
To my parents, who provided the tools; and to Marcus, who provided the rest.
iii ACKNOWLEDGMENTS First and foremost, I would like to tha nk Dr. Burne and my supervisory committee members, Drs. Paul Gulig, Jeannine Brady and Michael Kilberg. Their vested interest in my project and valuable suggestions al ong the way were much appreciated. An enormous amount of laughter, sweat, and t ears colored the lines of text in this book and danced behind the black and white figu res. Without this inspiration and the people who provided it, nothing w ould have been possible. For this, I would especially like to thank my teacher, Dr. Margaret Che n, and my student, (Dr.) Max Jameson-Lee. Their invaluable presence showed me that teaching and being taught are often one and the same. I would also like to thank the enti re Burne lab, past and present, for answering my incessant questions and for being extrao rdinarily charming every day. Without them, the past five years could have easily turned into ten. In addition, I would like to formally acknowledge the coaches in my life, who have provided me with the perseverance to accomplis h my goals. Specifically, I would like to thank my father, who never let me settle fo r a half-hearted effort; Coach Lou, who begat miracles simply by standing next to the trac k, shouting that achievement is 90% mental and only 10% physical; Mr. Lopez, my Taekwondo instructor, who always insisted that I try harder, yet always proclaimed that I was â€œPurrrrrFECKT;â€ and most recently, Dr. Burne, who gave me the opportunity to learn that nothing good comes easy. Finally, I would like to thank my family and friends â€” especially Mom, Jackie, Jerry, Sandy, Stan, Pop Pop, and the love ly ladies of 2105 Allegany â€” for their
iv unconditional love and support wh en I needed it most. Above all, I am grateful to my Marcus, who made this entire journey fun and worthwhile.
v TABLE OF CONTENTS page ACKNOWLEDGMENTS.................................................................................................iii LIST OF TABLES...........................................................................................................viii LIST OF FIGURES...........................................................................................................ix ABSTRACT......................................................................................................................xii CHAPTER 1 INTRODUCTION........................................................................................................1 Mutans Streptococci and Thei r Niche in the Oral Cavity.............................................1 Streptococci...........................................................................................................1 Ecological Determinants of the Oral Cavity.........................................................2 Formation of the Dent al Plaque Biofilm...............................................................3 Epidemiology of Dental Caries a nd the Role of Sugar Metabolism............................4 Alkali Generation by Acid -Sensitive Bacteria.............................................................6 Urease....................................................................................................................7 The Arginine Deiminase System...........................................................................7 Streptococcus rattus ............................................................................................11 Acid Tolerance Mechanisms of Cariogenic Bacteria.................................................12 Streptococcus mutans ..........................................................................................12 F1F0 proton-translocating ATPase................................................................13 Membrane permeability...............................................................................14 Macromolecule repair..................................................................................14 Identification of Genes En coding a Putative ADS in S. mutans UA159.............15 Summary.....................................................................................................................16 Specific Aims..............................................................................................................16 2 MATERIALS AND METHODS...............................................................................19 Bacterial Strains and Growth Conditions...................................................................19 DNA Manipulations....................................................................................................20 Streptococcus rattus FA-1...................................................................................20 Streptococcus mutans UA159.............................................................................21 RNA Manipulations....................................................................................................25 Streptococcus rattus FA-1...................................................................................26
vi Streptococcus mutans UA159.............................................................................26 Protein Manipulations.................................................................................................28 Construction of N-terminal 6X-His-tagged AdiR...............................................28 AdiR Protein Purification....................................................................................28 Gel Shift Assays..................................................................................................29 Expression of S. rattus Flp in S. mutans ..............................................................30 Construction of Promoter Fusions and CAT Assays..................................................30 Biochemical Assays....................................................................................................31 Acid Killing Assays....................................................................................................33 Nucleotide Sequence Accession Numbers.................................................................34 3 ISOLATION AND CHARACTERIZATION OF THE ARGININE DEIMINASE OPERON IN S. rattus FA-1.......................................................................................44 Introduction.................................................................................................................44 Results........................................................................................................................ .45 Isolation of the ADS Genes of S. rattus FA-1.....................................................45 Nucleotide Sequence Analysis of the ADS Genes..............................................46 Localization of parcA and Reporter Gene Fusions................................................50 Expression of AD in S. rattus ..............................................................................51 Summary.....................................................................................................................53 4 REGULATION OF ARGININE DEIMINASE EXPRESSION IN S. rattus FA-1...61 Introduction.................................................................................................................61 Results........................................................................................................................ .62 Identification of the AdiR Binding Site Within parcA..........................................62 Identification of Additional Regulat ory Genes Linked to the ADS in S. rattus FA-1.................................................................................................................65 Genetic Manipulation of S. rattus FA-1..............................................................67 Identification of an ADS -Like Gene Cluster in S. mutans UA159.....................70 Regulation of S. rattus parcA by S. mutans AdiR12 and Flp................................73 Summary.....................................................................................................................76 5 ANALYSIS OF AN AGMATINE DEIMINASE GENE CLUSTER IN S. mutans UA159.........................................................................................................................86 Introduction.................................................................................................................86 Results........................................................................................................................ .88 Analysis of the Sequence of the Agmatine Deiminase Gene Cluster.................88 Agmatine Deiminase Expression in S. mutans ....................................................92 Agmatine Deiminase Specificity.........................................................................94 Ammonia Production from Agmatine.................................................................95 Identification of the AgDS in Related Viridans Streptococci.............................97 Summary...................................................................................................................100
vii 6 REGULATION AND PHYSIOLOGIC SI GNIFICANCE OF THE AGMATINE DEIMINASE SYSTEM OF S. mutans UA159........................................................115 Introduction...............................................................................................................115 Results.......................................................................................................................116 Role of the S. mutans SMU.261c Protein..........................................................116 Localization of paguB and paguR...........................................................................117 Carbon Catabolite Repression...........................................................................118 Regulation of the AgDS by Growth Phase and Environmental Stress..............120 Agmatine Deiminase a nd Biofilm Ecology.......................................................122 Summary...................................................................................................................128 7 SUMMARY AND FUTURE DIRECTIONS...........................................................154 Arginine Deiminase System.....................................................................................154 Agmatine Deiminase System....................................................................................156 Role of Ammonia Generation in Mutans Streptococci.............................................159 LIST OF REFERENCES.................................................................................................161 BIOGRAPHICAL SKETCH...........................................................................................178
viii LIST OF TABLES Table page 2-1. Primers used............................................................................................................. .35 2-2. Plasmids used............................................................................................................37 2-3. Strains used............................................................................................................. ...39 2-4. Primers used to sequence the S. rattus agu operon...................................................41 2-5. Oligos used in gel shift experiments..........................................................................42
ix LIST OF FIGURES Figure page 1-1. Arginine deiminase system.........................................................................................17 1-2. Comparison of the ADS and AgDS pathways............................................................18 2-1. Complementation of the adiR12 mutation in S. mutans .............................................43 3-1. The S. rattus FA-1 arc operon....................................................................................54 3-2. Upstream sequence of S. rattus FA-1 arcA ................................................................55 3-3. Primer extension analysis of S. rattus arcA ................................................................56 3-4. Reverse transcriptase PCR analysis of mRNA from S. rattus ....................................57 3-5. CAT specific activity of the S. rattus FA-1 arcA promoter.......................................58 3-6. Arginine deiminase enzyme activity of S. rattus FA-1..............................................59 3-7. Arginine deiminase enzyme activity of S. rattus FA-1 chemostat.............................60 4-1. Alignment of the S. rattus and S. mutans AdiR1 proteins..........................................77 4-2. Gel shift analysis of purified His6x-AdiR from S. rattus ...........................................78 4-3. Alignment of the S. rattus and S. mutans Flp proteins...............................................79 4-4. Identification of putative ADS regulators in S. mutans UA159.................................80 4-5. Reverse transcriptase P CR of mRNA isolated from S. mutans ..................................81 4-6. A) Homologies of S. mutans Flp, AdiR1 and AdiR2.................................................82 4-7. Integration of the S. rattus parcA cat fusion into wild-type S. mutans and six engineered strains.....................................................................................................83 4-8. CAT specific activity of S. rattus parcA in S. mutans wild-type and four engineered strains.......................................................................................................................84
x 4-9. CAT specific activity of S. rattus parcA in S. mutans wild-type and two engineered strains.......................................................................................................................85 5-1. The putative AgDS gene cluster in S. mutans UA159.............................................102 5-2. Conserved regions of actual and putative agmatine deiminase................................103 5-3. Reverse transcriptase PCR analysis of mRNA from S. mutans ...............................104 5-4. Northern blot analysis of tota l RNA isolated from wild-type and aguB.................105 5-5. Slot blot analys is of mRNA isolated from wild-type S. mutans ...............................106 5-6. A) Agmatine deiminase enzyme activity..................................................................107 5-7. AgD activity of wild-type S. mutans UA159 grown under inducing conditions and assayed for production of Ncarbamoylputrescine from various primary amines.108 5-8. Ammonia production from agmatine at various pH values......................................109 5-9. Survival of wild-type S. mutans UA159 (WT) and an AgDS-defective strain harboring a polar mutation in aguB (B-) in 0.1 M glycine buffer, pH 2.8.............110 5-10. A) AgD enzyme activity in selected oral streptococci...........................................111 5-11. A) Northern blot of RNA isolated from S. rattus FA-1..........................................112 5-12. AgD enzyme activity of S. rattus FA-1..................................................................113 5-13. AgD enzyme activity of S. rattus FA-1 grown in TV medium containing 2 % glucose or galactose...............................................................................................114 6-1. Predicted structure of the S. mutans AguR protein..................................................129 6-2. AgD enzyme activity of S. mutans UA159 and aguR , in response to increasing concentrations (mM) of agmatine..........................................................................130 6-3. The aguRB intergenic sequence and primer extension analysis...............................131 6-4. Construction of paguB reporter gene fusions..............................................................132 6-5. AgD enzyme activity of S. mutans UA159, and of otherwise-isogenic ccpA and ccpB strains............................................................................................................133 6-6. AgD enzyme activity in relation to growth domain.................................................134 6-7. AgD activity in mid-exponentia l phase or stationary phase.....................................135
xi 6-8. AgD activity in co ntinuous cultures of S. mutans UA159 maintained at pH 5 or pH 7........................................................................................................................136 6-9. Real-Time RT-PCR of aguA expression (copies/ l) in continuous cultures of S. mutans UA159 maintained at pH 5 or pH 7...........................................................137 6-10. Ag D activity in response to growth at 37 C or 42 C...........................................138 6-11. AgD enzyme activity of S. mutans UA159 grown in TV me dium containing 25 mM galactose and 10 mM agmatine......................................................................139 6-12. AgD enzyme activity of S. mutans UA159 after incubation with 400 mM NaCl..140 6-13. Growth of S. mutans UA159, aguB , and aguC strains...........................................141 6-14. Growth of S. mutans wild-type, aguB , aguC and aguR strains..............................142 6-15. Growth of an aguRB double mutant strain.............................................................143 6-16. Growth of S. mutans and S. rattus strains..............................................................144 6-17. Growth of selected oral streptococci......................................................................145 6-18. Growth of S. mutans UA159 and aguB in the presence and absence of agmatine in TV medium contai ning 25 mM glucose.............................................................146 6-19. Growth of S. mutans UA159 and aguB in the presence and absence of agmatine in TY medium containi ng 25 mM galactose..........................................................147 6-20. Growth of S. mutans UA159 and aguB in the presence and absence of agmatine in TY medium contai ning 25 mM glucose.............................................................148 6-21. Growth of S. mutans UA159 and aguB in the presence and absence of agmatine in BHI medium.......................................................................................................149 6-22. Map of the putative oligopeptide ABC transport system.......................................150 6-23. Growth of the oppA and aguD mutants..................................................................151 6-24. Real Time RT-PCR (copies/ l) of the oppA , oppB , nitroreductase and aguA genes.......................................................................................................................152 6-25. Proposed role of the AgDS in virulence.................................................................153
Abstract of Dissertation Pres ented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy AMMONIA-GENERATING MECHANISMS OF MUTANS STREPTOCOCCI By Ann Ryan Griswold May 2006 Chair: Robert A. Burne Major Department: Medical Sciencesâ€“Immunology and Microbiology Streptococcus mutans is the etiological agent of dent al caries, a prevalent infectious disease and a cause of bacteria l endocarditis. Virulence of S. mutans results from continued growth and production of acid at pH values well below 5, generating a hostile environment for beneficial oral flora. Interestingly, the closest relative of S. mutans is Streptococcus rattus, a comparatively acid-se nsitive bacterium associated with dental health. Survival of acidification by S. rattus is enhanced thro ugh ammonia production via the arginine deiminase system (ADS). The ADS consists of arginine deiminas e (ArcA), ornithine carbamoyltransferase (ArcB), and carbamate kinase (ArcC), which convert arginine to ammonia, ATP, and CO2. S. rattus ADS enzymes are transcribed in one operon that also contains an arginine:ornithine antiporter (ArcD) a nd a transpeptidase (ArcT). A putative 70 arc promoter (parcA) was located by primer extension. Th e ADS is activated by AdiR in the presence of arginine and is regulated by carbon catabolite repressi on, apparently through
a CcpA-dependent pathway. Gel shift assays identified a putative AdiR binding site within the first 80 bases of parcA. Interestingly, a gene cluster with simila rity to the ADS genes was discovered in S. mutans , an organism considered incapabl e of arginine catabolism. The S. mutans aguR-aguBDAC genes were found to encode an agmatine deiminase system (AgDS) consisting of agmatine deiminase (AguA), put rescine carbamoyltran sferase (AguB), and carbamate kinase (AguC), which convert ag matine to putrescine, ammonia, ATP, and CO2. The AgDS is activated by low pH and heat, as well as by agmatine via AguR. Additionally, the AgDS is subject to carbon catabolite repression via a CcpA/CcpBindependent mechanism. Ag matine inhibited growth of S. mutans , suggesting a role for the AgDS in agmatine detoxification. Examination of nearly identical system s in closely-related but physiologically diverse mutans streptococci revealed mu ltifaceted roles for ammonia production. In S. rattus , the ADS primarily generates ATP and al kalinizes the extracellular environment during growth at low pH. In contrast, the physiological role of the AgDS in S. mutans is complex, conveying bioenergetic advantages by enhancing pH and generating ATP, as well as by detoxifying agmatine produced by acid-sensitive organisms in response to acidification by S. mutans .
1 CHAPTER 1 INTRODUCTION Mutans Streptococci and Thei r Niche in the Oral Cavity Streptococci The bacterial genus Streptococcus is comprised of Gram-positive, microaerophilic, non-motile, chain-forming cocci. The name â€œstreptococcusâ€ originated from the Greek word streptos, which describes a chain-like structure that is easily bent or twisted. Streptococcus species are organized into groups acco rding to their hemolytic properties. Many oral streptococci, including the muta ns streptococci, belong to the larger, -hemolytic, viridans group. The name â€œviridansâ€ originated from the Latin word viridis , which describes the green halo of -hemolysis that forms around viridans streptococci grown on blood agar plates. The mutans streptococci can be furthe r distinguished from other viridans streptococci by their ability to (1) fermen t mannitol and sorbitol , (2) produce glucans from sucrose, and (3) sustain growth at low pH (Carlsson, 1967). Members of the mutans group (with their natural hosts in parentheses) include: Streptococcus mutans (human), Streptococcus sobrinus (human), Streptococcus cricetus (human, animal) , Streptococcus rattus (human, rat), Streptococcus ferus (rat), Streptococcus downeii (monkey) and Streptococcus macacae (monkey). The oral streptococci occupy a variety of niches in the oral cavity, including the tongue, cheeks, teeth and saliva (Frandsen et al. , 1991). The mutans streptococci, in particular, are most commonly isolated from the dental plaque of their respective hosts
2 (Bowden et al. , 1979). The preferred habitat of each species within the oral cavity is dictated by a variety of ecological determinants, as described below. Ecological Determinants of the Oral Cavity The oral cavity is composed of severa l anatomical structures, each of which support distinct microbial communities. The normal flora of the tongue, for example, is different than the flora typically found on the tooth surface (Frandsen et al. , 1991; Loesche, 1982). Even multiple teeth in the same person or multiple surfaces on the same tooth can harbor different microbial populations (Loesc he, 1982). The ecological determinants that compel microbial species to preferentially colonize specific sites in the mouth are complex, but generally include the abi lities to 1) adhere to a particular surface (Gibbons and van Houte, 1975), 2) metaboli ze the available nutrients (Bibby, 1976), and 3) withstand the environmental stresses and in hibitory factors encountered in the habitat (van Houte, 1976). For example, the mucosal surfaces of the mouth are exposed to air, and thus attract a larger numb er of microaerophilic or faculta tive anaerobic bacteria than the highly anaerobic folds of the cheeks and crevices of the tongue (Loesche et al. , 1983). In general, saliva contains the most he terogeneous group of bacteria, due to washoff from all of the surfaces in the mouth (Loesche, 1982). However, because of the constant flow of saliva and the frequency of swallowing, most bacteria are unable to remain in the saliva permanently and are for ced to re-attach and re-colonize the surfaces from which they were derived in order to pe rsist in the oral cavity (Ellen and Burne, 1996; Gibbons, 1989). Moreover, the flow of saliv a also results in the physical removal of unattached bacteria from the surfaces of the mouth. Therefore, the ability to adhere to a surface is a major ecological dete rminant for the oral microbiota.
3 Some bacteria, such as the mutans strept ococci, are well suited for attachment to the tooth, a non-shedding structure with a la rge surface area that favors accumulation of bacteria. The biofilm formed by microorga nisms and their extracellular products is commonly referred to as dental plaque. Oral streptococci ar e abundant in dental plaque, typically comprising around 23% of the total cultivable mi croflora, while the proportion of mutans streptococci range s from 2-5% in samples from healthy individuals (Bowden et al. , 1975). A variety of other ba cteria are also commonly isol ated from dental plaque, including an abundance of Gram-positive (e.g. Actinomyces spp) and Gram-negative (e.g. Prevotella spp.) rods, as well as Neisseria spp., Veillonella spp., Fusobacterium spp., and Lactobacillus spp. Formation of the Dent al Plaque Biofilm As mentioned above, the accumulation of bacteria and extracellular components on the tooth surface is referred to as the dent al plaque biofilm. A biofilm is generally defined as a community of microorgani sms attached to a surface (Costerton et al. , 1987). Dental plaque biofilms are three-dimensional matrices formed by the structured accumulation of a diverse community of mi croorganisms and extracellular material derived from the surrounding environmen t and from the bacteria (Costerton et al. , 1987). Bacteria growing in biofilms often exhib it enhanced resistan ce to host defenses, antimicrobials, and environmental st ress (Caldwell and Costerton, 1996). Dental plaque biofilm formation begins with the acquired pell icle, which develops when acidic glycoproteins from saliva adhere to the tooth surface and form a thin film. Early plaque colonizers, su ch as streptococci and Actinomyces spp. (Kolenbrander and London, 1993), adhere to the acquired pellicle via hydrophobic or lectin -like interactions, followed by co-aggregation with late-coloni zing bacteria, such as obligate anaerobes
4 (Kolenbrander and London, 1993). Bacteria in the dental plaque biofilm become irreversibly bound in the presence of sucros e, which is used to form extracellular adherent glucans and fructans (Ellen and Burne, 1996). Epidemiology of Dental Caries and the Role of Sugar Metabolism Dental caries is one of the most prevalen t infectious diseases in both industrialized and developing countries. The United States Centers for Disease Control has determined that dental caries is by far the most co mmon chronic disease of childhood, surpassing the second-most common childhood disease, as thma, by over 8-fold (Beltran-Aguilar et al. , 2005). More than 91% of Americans over th e age of 20 have expe rienced some degree of dental caries in their pe rmanent teeth (Beltran-Aguilar et al. , 2005). As a result, the economic burden of dental caries in the Un ited States is overwhelming: over $40 billion is spent on treatment and prevention of dental caries each year, according to estimates by the American Dental Association. Hence, a gr eat deal of research has been devoted to understanding the pathogenesis of this costly and ubiquitous disease. Epidemiological studies indica te that the prevalence of caries increased drastically in the 20th century when sucrose became a majo r component of the diet (Gustafsson et al. , 1954; Marthaler, 1967; Scheinin et al. , 1976; Zita et al. , 1959). Sucrose is a low molecular weight di-saccharide that can be easily sequestered and metabolized by many oral bacteria. Throughout the late 1800s and early 1900s, evidence accumulated to link bacterial sugar metabolism to acid producti on and tooth decay. In 1860, Pasteur first noted that bacteria are capable of producing lactic acid via suga r fermentation (Pasteur et al. , 1860). Seven years later, Magitot exposed teeth to sugar fermentation mixtures and showed that dental car ies could be induced in vitro via exposure to the acidic by-products of sugar metabolism (Magitot, 1867). Mag itotâ€™s findings were expanded in 1890, when
5 Miller demonstrated that bacteria present in human saliva can produce lactic acid from dietary components such as bread and sugar, an d that the lactic acid is made in sufficient quantities to play a significant role in tooth decay. Furt hermore, Miller proposed that tooth decay occurs in two steps: 1) decalci fication of tooth enamel by exposure to acid, and 2) dissolution of the softened, decalcified tissue by parasitic digestion. Miller concluded that any bacteria capable of both acid production and protein digestion could contribute to tooth de cay (Miller, 1890). The first in vivo study to correlate ba cterial sugar metabolism and acid production was performed in 1940, when Stephan describe d a causal relationshi p between the rapid decline in plaque pH observed after a suga r rinse and the presence of a mixed population of plaque bacteria (Stephan, 1940; Stephan and Hemmens, 1947). The decline in pH was linked to bacterial production of lactic acid, an end product of sugar metabolism via the Embden-Meyerhof Parnas (EMP) pathway (S andham and Kleinberg, 1970a, b). When acid production subsided, Stephan observed a gr adual rise in plaque pH that eventually reached a plateau. Moreover, the plateau (or resting pH) of caries-active dental plaque was found to be more acidic than th at of caries-free plaque (Margolis et al. , 1988), further supporting a connection between ac id and dental caries. The factors responsible for the gradual rise in pH were therefore of major inte rest to researchers, gi ven the apparent role of acid production in dental caries. Subsequent research revealed that the ri se in pH observed by Stephan was due to ammonia production from arginine or urea by a subset of acidsensitive organisms present in saliva and dental plaque (Wijeyeweera and Kleinberg, 1989b). This observation gave rise to the not ion that, while many plaque ba cteria metabolize sugar into
6 acidic end-products, not all sugar-metabol izing bacteria are implicated in the pathogenesis of dental caries. Thus, the Ste phan curve illustrates the dynamic that exists between acid-tolerant and acid-sensitive ba cteria residing in th e plaque biofilm. Following dietary intake, plaque bacteria rapidly metabolize sugar into acidic end products, effectively lowering the pH and pe rpetuating an acidic environment that is antagonistic to colonization by acid-sensitiv e flora (Sandham and Kleinberg, 1970a, b). After the available sugar has been metaboli zed, acid-sensitive arginolytic and ureolytic bacteria mediate a gradual re turn to resting pH by producin g ammonia from arginine and urea, respectively (Wijeyeweer a and Kleinberg, 1989b). Dental caries result when prolonged acidification of dent al plaque invokes a shift in the microbial species composition of the bi ofilm, whereby the acid-sensitive bacteria begin to die off and the acid-to lerant bacteria continue to survive, proliferate, and produce acid (Wijeyeweera and Kleinberg, 1989b). When dental plaque reaches a critical pH of around 5.5, tooth enamel undergoes decalc ification and dissolu tion, resulting in dental caries (Afonsky, 1961; Loesche, 1986). Alkali Generation by Acid-Sensitive Bacteria Acid-sensitive oral bacteria have develope d a variety of strate gies to cope with environmental acidification, in cluding mounting of an adapti ve acid tolerance response and production of alkali. The alkali-generating mechanism us ed by salivary bacteria is the hydrolysis of urea by urease, while plaque bacteria rely on catabo lism of arginine via the arginine deiminase system (ADS) (Burne and Marquis, 2000). Ammonia generation by these mechanisms maintains pH across the cytoplasmic membrane and raises the pH of the external environment, helping to st abilize healthy plaque biofilm communities.
7 Urease During environmental acidification, a subset of salivary bacteria rely on the urease enzyme to hydrolyze urea to ammonia and CO2 (Sissons et al. , 1989). Urea levels in saliva are relatively high, ranging from 3 to 10 mM in most individuals (Kopstein and Wrong, 1977). Although a small number of speci es express urease activity, these species are capable of rapidly acquiring and metabolizing the available urea (Sissons and Cutress, 1988). As a result, ureolysis is thought to contribute a large amount of ammonia to salivary and plaque fluid (Wijeyeweera and Kleinberg, 1989b). Much of the research on urease activity in oral bacteria has focused on S. salivarius and Actinomyces naeslundii, two abundant members of the oral flora. Previous work in the Burne laboratory has shown that ureolysis by oral bacteria can significantly al kalinize both saliva and dental plaque, and effectively hinder the progression of dent al caries (Chen et al. , 2002; Clancy et al. , 2000; Morou-Bermudez and Burne, 2000; Shu et al. , 2003). The Arginine Deiminase System Physiological role of the ADS. The ADS has been identified in a broad range of prokaryotes (Abdelal, 1979). The physiologi cal role of the system and mode of regulation varies among species, but an ove rall theme involves ge neration of ATP in accordance with equimolar hydrolysis of substrat e, allowing the use of arginine as a sole energy source (Crow and Thomas, 1982). Saliva contains approximately 50 M free arginine. In addi tion, many oral bacteria possess peptidases capable of cleaving argi nine from salivary proteins, such as the histidine-rich peptides, which contain appr oximately 10 to 20 mol-percent arginine (Curran et al. , 1998; Rogers et al. , 1988; Van Wuyckhuyse et al. , 1995). Thus, the ADS is a convenient means for acid-sensitive plaque bacteria to generate ATP as an energy
8 source, particularly when the pH is too lo w for glycolytic enzymes to function (CasianoColon and Marquis, 1988) (Burne and Mar quis, 2000). Plaque bacteria known to hydrolyze arginine via the ADS include Streptococcus gordonii, S. sanguis , Streptococcus parasanguis , Streptococcus milleri , S. oralis , S. mitis biovar 2, S. rattus and Lactobacillus fermentum (Dong et al. , 2002; Ferro et al. , 1983; Floderus et al. , 1990; Griswold et al. , 2004a; Hiraoka et al. , 1986; Marquis et al. , 1987; Poolman et al. , 1987). In addition to ATP generation, ammonia generation via the ADS neutralizes the bacterial cytoplasm (Casiano-Colon and Mar quis, 1988) and alkalinizes the surroundings, further protecting acid-sensitive oral bacteria from lethal acidification (Casiano-Colon and Marquis, 1988) . The ADS is a major source of ammonia in dental plaque, which has been measured at levels in excess of 52 mM (Edgar and Higham, 1990; Margolis et al. , 1988; Tatevossian and Gould, 1976; Wijeye weera and Kleinberg, 1989a). Other pathways that contribute a mmonia to dental plaque in clude ureolysis, Stickland fermentation (1 Ala + 2 Gly + H20 3 Acetate + CO2 + 3 NH3), and deamination of amino acids (Curtis et al. , 1983). In non-oral streptococci, the ADS may play a role in virulence. An AD homologue, SAGP (streptococcal acid glycoprotein) was identified in S. pyogenes , and was subsequently found to be capable of inhi biting proliferation of human T lymphocytes (Degnan et al. , 1998). Mutation of SAGP redu ced the acid tolerance of S. pyogenes , suggesting that the ADS system may help this organism survive within acidic phago-lysosomes (Degnan et al. , 2000). In addition, a homologue of SAGP was identified in Streptococcus suis , a pathogen implicated in meningitis in humans, as well as meningitis, septicemia, and arthritis in pigs. The SAGP homologue, AdiS, was
9 identified during a screen for cell-surface pr oteins up-regulated in response to heat (Winterhoff et al. , 2002), suggesting that the S. suis ADS may be induced during colonization of deep tissues or upon entering the bloodstream. Genetic structure. Genes encoding the ADS are commonly arranged as an operon, although the gene order varies among bacteria (Barcelona-Andres et al. , 2002; Dong et al. , 2002; Maghnouj et al. , 1998; Vander Wauven et al. , 1984; Zuniga et al. , 1998). Typically, the operons contain arcA , encoding arginine deiminase (AD), which hydrolyzes arginine to generate citr ulline and ammonia. A second gene, arcB , encodes a catabolic ornithine carbamoyltransferase (cOTC) , which converts citrulline to ornithine and carbamoylphosphate. Finally, arcC encodes a catabolic carbamate kinase (CK), which transfers a phosphate group from carba moylphosphate to ADP, generating ATP, CO2 and ammonia. Many ADS operons also incl ude one or more additional genes, which may encode an arginine-ornithine antiporter ( arcD ), a putative aminopeptidase ( arcT ), or transcriptional regulators of the Crp/Fnr ( arcR ) and/or ArgR/AhrC families ( argR ) (Dong et al. , 2002; Zuniga et al. , 1998). ADS regulation. In most bacteria studied thus far, the ADS appears highly regulated at the genetic level (Barcelona-Andres et al. , 2002; Dong et al. , 2002; Maghnouj et al. , 1998; Zuniga et al. , 2002). Molecular char acterizations of the arc operons of other oral st reptococci, particularly S. gordonii and S. sanguis, indicate that AD activity in is induced by arginine and is subject to carbon catabolite repression (CCR), with lower activity observed in the presence of the repressing sugar, glucose (Dong et al. , 2002; Ferro et al. , 1983). In AT-rich, Gram-positive bacteria, CCR is mediated by the trans -acting catabolite control protein A (CcpA), which binds to
10 cis -acting catabolite response elements ( cre ) in the presence of preferred carbohydrate sources to regulate the expression of catabolic genes and operons (Saier et al. , 1996). In S. gordonii , the trans -acting ArcR protein activates the arc genes in response to arginine (Dong et al. , 2002). In the absence of ArcR, AD ac tivity is decreased and the effects of catabolite repression and s ubstrate induction are dimini shed. In addition, the S. gordonii arc operon is induced in respons e to anaerobic growth by flp, an Fnr-like protein (Dong et al. , 2004). However, anaerobic induction does not appear to play a major role in arc expression of other related micr oaerophilic bacteria, such as S. rattus and E. faecalis (Burne et al. , 1991; Simon et al. , 1982). Incidentally, the E. faecalis arc operon does not appear to be linked to any fnr -like regulators, possibly explaining the insensitivity of this system to oxygen tension (Barcelona-Andres et al. , 2002). However, in bacteria capable of both aerobic respiration and anaerobic ferm entation, aeration is the primary regulator of arc expression. For example, in Psuedomonas and Bacillus , ArcR regulates the ADS by binding to a Crp consensus sequence and activating the arc operon during conditions of oxygen limitation, while the system is st rictly repressed du ring aerobic growth (Gamper et al. , 1991; Maghnouj et al. , 1998). In these bacteria , arginine induction is mediated by ArgR, a homolog of AhrC that was originally identified in Bacillus subtilis as a repressor of the arginine biosynthe tic genes (Mountain and Baumberg, 1980). In B. licheniformis, ArgR activates the ADS by binding to a conserved ARG box upstream of the arcA promoter (Maghnouj et al. , 2000). Multiple transcrip tional activators, usually ArcR in combination with Flp or ArgR/AhrC, are linked to the arc operons of most bacteria, including S. gordonii, E. faecalis and B. licheniformis , (Barcelona-Andres et al. , 2002; Dong et al. , 2004; Maghnouj et al. , 1998; Maghnouj et al. , 2000).
11 Streptococcus rattus The ADS-using bacterium, S. rattus , was originally isolated from the dental plaque of a laboratory rat, although it is estimated that this strain colonizes de ntal plaque in up to 50% of the human population (Bratthall, 1972). Like other ADS-pos itive oral bacteria, S. rattus has not been linked to caries development. However, S. rattus is unique because it is the only ADS-utilizing bacteria to bel ong to the mutans group of streptococci (Coykendall, 1974). Biochemical characteri stics, as well as 16S rRNA analyses (Kawamura et al. , 1995) and DNA hybridization studies (Coykendall et al. , 1971), confirm that S. rattus is a member of this taxonomic gr oup, and is the closest relative of S. mutans , the etiological agent of dental caries. Despite this taxonomic relationship, S. rattus is the least cariogenic and ac iduric of the mutans group. Previous research concerning the ADS in S. rattus FA-1 focused on protection against acid killing and modulation of ADS enzyme activity by aeration and carbohydrates. The ADS can protect S. rattus from acid killing at pH 3.5 to 4.0, which are clinically relevant levels (Casiano-Col on and Marquis, 1988). The optimal pH for ammonia production from argini ne is around pH 3.5, although arginolysis can occur at pH values as low as 2.1 in intact cells (Marquis et al. ), allowing S. rattus to raise both the cytoplasmic and external pH to neutral levels (Marquis et al. , 1987). Compared to other ADS-positive oral streptococci, such as S. sanguis and S. gordonii , the ADS of S. rattus FA-1 is considerably more acid-tolerant and is much less sensitive to aeration and catabolite repression (Burne et al. , 1991; Curran et al. , 1998). Thus, regulation of this operon may differ significantly from nonmutans ADS-positive streptococci. A molecular characterization of the arc operon of S. rattus FA-1 may provide a clearer
12 understanding of the evolution a nd diversity of the mutans stre ptococci in relation to pH adaptation and cariogenicity. Acid Tolerance Mechanisms of Cariogenic Bacteria A large amount of research has focused on dissecting the various mechanisms of acid tolerance employed by cariogenic bacteria , as knowledge of these mechanisms is key to the development of novel anti-caries st rategies. Of the seve ral hundred bacterial species known to colonize dental plaque, S. mutans, Streptococcus sobrinus , and Lactobacillus casei are among the most notorious ly acidogenic (Loesche, 1986) (Socransky et al. , 1982). S. mutans and S. sobrinus can sustain growth and carry out glycolysis at pH values below 5, lowering th e extracellular plaque pH to 4 and below (de Soet et al. , 2000). Streptococcus mutans Streptococcus mutans was originally isolated by Clarke in 1924 from carious lesions in humans (Clarke, 1924), and by the 1960s was recognized as a major etiologic agent of dental caries (Keyes, 1960) (F itzgerald and Keyes, 1960) (Loesche, 1986). However, S. mutans colonizes both healthy and cari ogenic plaque, implying that an additional factor is required fo r initiation of disease. As discussed above, dietary sucrose plays an important role in caries formation. S. mutans uses invertase to convert sucrose into glucose and fructose, which are furthe r broken down into lactic acid (Kuramitsu, 1973). In addition, sucrose can be converted, via glucosyltransferase enzymes, into glucans, which are glucose polymers that mediate attachment of S. mutans to the tooth surface (Kuramitsu, 1976). Thus, high-sucrose diets lead to acidificati on of dental plaque by favoring the outgrowth of S. mutans . Streptococcus mutans has been estimated to
13 comprise anywhere from 0 to 50% of the total dental plaque biofilm, depending on the cariogenic state of the sample (Gibbons and van Houte, 1975). Several determinants of acid tolerance in S. mutans have been characterized, including a membrane-bound F1F0-ATPase, reduction in permeability of the cell membrane to protons, induction of DNA re pair pathways, and up-regulation of stress-specific pr oteins (Bender et al. , 1986; Cox et al. , 2000; Hahn et al. , 1999; Hanna et al. , 2001; Kremer et al. , 2001; Lemos et al. , 2001; Lemos and Burne, 2002; Quivey et al. , 2001). These systems allow S. mutans to carry out glycolysis until dental plaque becomes very acidic without suffering lethal effects from the accumulation of glycolytic end products in the medium. F1F0 proton-translocating ATPase In order to maintain a transmembrane pH gradient of 0.5 to 1 unit during growth in acidic conditions, S. mutans relies on extrusion of intracellular protons via an acid-inducible F1F0-ATPase (Bender et al. , 1986). The pH optima of the S. mutans F-ATPase is 6.0, similar to that for S. sobrinus (Bender et al. , 1986; Nascimento et al. , 2004). However, the pH optima of F-ATPase s in acid-sensitive bacteria, such as S. sanguis and S. salivarius, are 7.5 and 7.0, respectively (Bender et al. , 1986). Therefore, the ATPases of acid-senstive bacteria appear ill-equipped to maintain pH during extreme acidification of dental plaque , as compared to the ATPases of S. mutans and S. sobrinus . Induction of F-ATPase activity at lo w pH may thereby contribute to the competitive advantage of S. mutans over acid-sensitive plaque bacteria (Quivey et al. , 2001).
14 Membrane permeability The cell membrane provides the first barri er to extracellular acid by reducing the influx of protons into the cell. Consequen tly, membrane biosynthesis and assembly are critical for survival at low pH because of their role in the integrity of the cell structure. In response to acid, S. mutans alters its membrane com position, increasing production of mono-unsaturated and long-chain fatty acids, which could alte r the proton permeability of the lipid bilayers or indirec tly affect secretion and functi on of enzymes involved in acid tolerance, particularly the F1F0-ATPase (Quivey et al. , 2000). Mutation of dltC , a gene involved in D-alanyl-lipotei choic acid synthesis (Boyd et al. , 2000), and dagK , a gene involved in phospholipid metabo lism (Yamashita, 1993), render S. mutans incapable of surviving at low pH. In addition, mutation of the ffh gene, encoding a homologue of the eukaryotic signal recognition particle , significantly hinders ability of S .mutans to withstand environmental acidification (Kremer et al. , 2001). Interestingly, this mutation also reduces F1F0-ATPase activity, suggesting that Ffh assists with the membrane rearrangement required for in sertion of additional ATPase complexes during growth in acidic conditions. Macromolecule repair The ability to repair denatured DNA and prot eins is essential for continued growth of bacteria in acidic environments. One of the major consequences of growth at low pH is increased protonation of pur ines and pyrimidines, and subsequent cleavage of the glycosyl bond, which results in AP (apurinic or apyrimidinic) sites (Lindahl and Nyberg, 1972). In S. mutans , these sites are repaired by acid-inducible AP endonucleases, which resemble the E. coli exonuclease III (Hahn et al. , 1999). In addition to AP endonucleases, the DNA repair protein RecA is required for repair of stalled replication
15 forks and regulation of the SOS response. Mutation of S. mutans recA increases sensitivity to killing at pH 2.5 (Cox et al. , 2000), but acid adaptation alleviates this sensitivity, suggesting that other acid-inducible DNA repair enzymes, such as AP endonucleases, can compensate fo r the loss of RecA (Quivey et al. , 1995). The S. mutans UvrA protein also assists with acid to lerance by excising regions of damaged DNA following exposure to acid or UV radiation (Hanna et al. , 2001). At low pH, molecular chaperones are required to stabilize, repair , and protect DNA repair enzymes and other proteins (Lemos et al. , 2001; Lemos and Burne, 2002). DnaK and GroEL are among the best characterized chaperones involved in the S. mutans ATR. These chaperones also function in the heat shock response. A DnaK knockdown that reduced protein levels by 50% displayed increased acid sensi tivity and diminished ability to lower the pH when incubated with glucose in pH drop experiment s. In addition, the Clp and HtrA proteins are involved in target ing misfolded or damaged proteins for degradation. Mutation of the ClpP peptidase not only re duced acid tolerance of S. mutans , but also reduced biofilm formation and genetic transformability (Lemos and Burne, 2002). Identification of Genes Encoding a Putative ADS in S. mutans UA159 In oral streptococci that are genera lly considered less acid tolerant than S. mutans , such as S. gordonii and S. sanguis , generation of ammonia by the ADS protects against environmental acidification (Bur ne and Marquis, 2000). Whereas S. mutans metabolizes a more extensive range of carbohydrates than many Gram-positive bacteria (Ajdic et al. , 2002), it does not have the ability to hydrolyze argi nine via the ADS (Perch et al. , 1974). In fact, absence of a functional ADS is the major distinguishing factor between S. mutans and its closest relative, S. rattus (Coykendall, 1974). However, the recent publication of the S. mutans UA159 genomic sequence led to the di scovery of a gene cluster with
16 striking similarity to known ADS genes (Ajdic et al. , 2002). Considering the role of acid tolerance in virulence of S. mutans , the role of this system in arginine catabolism is worthy of investigation. Summary The ability to tolerate environmental acidi fication is essentia l for oral bacteria associated with dental health, as well as for those associated with disease. However, the acid tolerance mechanisms used by cario genic and non-cariogenic bacteria differ drastically in their impact on oral healt h. Many acid-sensitive generate alkali from arginine or urea to raise the extracellular pH a nd create a more habitable environment. In contrast, cariogenic bacteria such as S. mutans employ acid tolerance mechanisms to maintain a pH of 0.5-1 to allow continued glycolys is (and acid productio n) at low pH. Thus, the mutans streptococci is an interes ting group to study from the perspective that it contains both the etiological agent of dental caries ( S. mutans ) and an apparently noncariogenic bacterium ( S. rattus ). The intriguing discovery of an ADS-like gene cluster in S. mutans presents a unique opportu nity to examine the role of ammonia-generating mechanisms in the virulence and acid:b ase physiology of muta ns streptococci. Specific Aims Identify the genes encoding the ADS of S. rattus FA-1 and examine the ADS-like gene cluster of S. mutans UA159, confirm ORF expressi on, map the promoters and measure enzyme activity Examine the regulation of the ADS in S. rattus and the AD-like system in S. mutans , specifically focusing on the roles of the putative regulatory proteins SMU.261c and ArcR, and the effects of low pH and carbon catabolite repression on expression of the operons Compare analogous ammonia generating mechanisms used by S. rattus FA-1 and S. mutans UA159
17 Figure 1-1. Arginine deiminase system Arginine Citrulline NH3 Ornithine Carbamoylphosphate Pi NH3 + CO2 ATP ADP cOTC Anti p orte r C K AD
18 Figure 1-2. Comparison of the ADS and AgDS pathways. Agmatine can be produced from arginine by arginine decarboxylase. AD, arginine deiminase; OTC, ornithine carbamoyltransferase; CK , carbamate kinase; AgD, agmatine deiminase; PTC, putrescin e carbamoyltransferase. CO2Ar g inine Decarbox y lase NH3 + CO2 AD ATPCK Agmatine Carbamoylputrescine NH3 AgD Putrescine PTCPi AgDS Arginine AD Citrulline NH3 Ornithine OTC Pi ADS Carbamoylphosphate
19 CHAPTER 2 MATERIALS AND METHODS Bacterial Strains and Growth Conditions S. rattus FA-1 and S. mutans UA159 were grown in brain heart infusion (BHI, Difco Laboratories, Detroit, MI) broth at 37oC, in 5% CO2 and 95% air. To monitor ADS expression, wild-type S. rattus FA-1 was grown in trypton e-vitamin (TV) based broth (Burne et al. , 1999) containing 0.2 or 2% glucose or ga lactose, with or without 10 mM arginine. To monitor AgDS expressi on, wild-type and mutant strains of S. mutans UA159 were grown in TV medium (Burne et al. , 1999) containing 0.5% glucose or galactose, with or without 10 mM agmatine. Recombinant E. coli strains were maintained on L agar supplemented, when indicated, with 25 g mL-1 of chloramphenicol (Cm) or 50 g mL-1 of kanamycin (Km). Chemical reagents were obtained from Sigma (St. Louis, MO). For studies on pH-dependent regulati on of AgDS expression, steady-state continuous cultures of S. mutans were grown in a Biostat i Twin Controller chemostat (B. Braun Biotech, Inc., Allentown, PA) in a tr yptone-yeast extract (TY) medium (Wexler et al. , 1993) supplemented with 25 mM glucose at a dilution rate ( D ) of 0.3h-1. Cultures were maintained at pH 5 or pH 7 by the addi tion of 2 M KOH. Where indicated, cultures were pulsed with 3 mM agmatine for one hour prior to sampling. Growth curves of S. mutans UA159 and mutant strains were generated using a Bioscreen C (Oy Growth Curves AB Ltd., Hels inki, Finland). Optical density at 600 nm was recorded every 30 minutes, with shaking for 10 seconds before each reading.
20 DNA Manipulations Plasmid DNA was isolated from E. coli by the method of Birnboim and Doly (Birnboim and Doly, 1979). Plasmid DNA used in sequencing reactions was further purified using the QIAprep Spin Plasmid Kit (Qiagen Inc., Valencia, CA). Cloning and electrophoretic anal ysis of DNA fragments were carried out according to established protocols (Ausubel et al. , 1989). Southern hybridization and high-stringency washes were performed as previously described (LeB lanc and Lee, 1982). Restriction and DNA modifying enzymes were purchased from Li fe Technologies Inc. (Rockville, MD) or New England Biolab (Beverley, MA). Primer sequences are listed in Table 2-1. Genetic properties of all plasmids and bacterial stra ins described below are available in Tables 2-2 and 2-3, respectively. Streptococcus rattus FA-1 Genomic DNA from S. rattus FA-1 was isolated as pr eviously described (Chen et al. , 1996). To prepare sub-genomic DNA libraries of S. rattus FA-1, chromosomal DNA was digested to completion using Xba I or Eco RV. The digested DNA fragments were separated on agarose gels and the Xba I fragments of 6to 8-kbp or Eco RV fragments of 4to 5-kbp were enriched by gel purifica tion using the Elu-Quik DNA Purification Kit (Schleicher & Schuell, Keene, NH.). The isolated DNA fragments were ligated onto Xba Ior Hin cII-digested, phosphatase-treated pSU20 (Bartolome et al. , 1991), respectively. The ligation mixt ures were used to transform E. coli DH10B and Cm resistant transformants were selected. To develop an arcB -specific probe, an internal fragment of the S. rattus FA-1 arcB gene, encoding cOTC, was generated by PCRs using degenerate primers based on alignments of known anabolic and catabolic OTCs (Dong et al. , 2002). Using the arcB
21 fragment as a probe, a la mbda clone containing the arcA gene and a partial arcB gene was identified. Specific primers internal to arcB were designed and the subsequent PCR product was used to screen a subgenomic library. Primer arcB -S encoded amino acid residues 3 through 8 of cOTC. Primer arcB -AS encoded the antisense sequence, corresponding to amino acid residues 127-132 of cOTC. Each reaction consisted of 25 cycles at a stringent annealing temperature (55oC). The product with the correct predicted size was gel purified prior to cloni ng into pCRII to generate pJZ22. Southern blot analysis (Southern, 1975) confirme d hybridization of the PCR product to S. rattus FA-1 chromosome at high stringency. Se quence analysis and BLAST searches were performed to confirm that the product shar ed high degrees of homology with other cOTCs. The agu operon of S. rattus FA-1 was sequenced using the primers in Table 2-4. Initial sequences were derived by PCR amplification of S. rattus chromosomal DNA using degenerate primers based on the S. mutans agu operon. Preliminary sequence analysis was conducted on the amplicon using primers from S. mutans . Additional primers were subsequently designed, based on the progressively available S. rattus agu sequence. Streptococcus mutans UA159 Genomic DNA was isolated from S. mutans UA159 as previously described (Chen and Burne, 1996). Recombinant PCR (Higuc hi, 1990)was used to construct a polar aguB insertional mutant ( aguB :: Km). The first half of aguB was amplified using primer pairs aguB -SXba I and aguB -ASEco RI, which inserted Xba I and Eco RI restriction sites. The remaining portion of aguB was amplified using primer pairs aguB -SEco RI and
22 aguB -ASSst I, which inserted Eco RI and Sst I restriction sites to facilitate cloning. The PCR fragment was then cloned onto pGEM7zf(+) a nd electroporated into E. coli DH10B. The construct was then digested with Eco RI and an kanamycin cassette harboring strong transcription/translat ion termination signals ( KmR) (Perez-Casal et al. , 1991) was inserted at the Eco RI restriction site to disrupt aguB . The construct was transferred into S. mutans , selecting for growth on BH I agar containing 1.0 mg mL-1 of Km, and correct integration was confirme d by Southern blot analysis. To develop an aguB -specific probe, an internal fragment of S. mutans UA159 aguB , encoding a putative putrescine carbamoy ltransferase, was PCR amplified using primer pair aguB -S and aguB -AS. Each PCR reaction consisted of 25 cycles at a stringent annealing temperature (55 C). Southern blot analys is confirmed hybridization of the PCR product to the S. mutans UA159 chromosome at high stringency. Recombinant PCR was used to construct a non-polar aguC insertional mutant ( aguC Km). The 5 -half of aguC was amplified using primer pairs aguC -SPst I and aguC -ASSma I, which inserted Pst I and Sma I restriction sites to facilitate cloning. The remaining portion of aguC was amplified using primer pairs aguC -SSma I and aguC -ASSst I, which inserted Sma I and Sst I restriction sites. The two primary PCR products were then used in a reaction with primers aguC -SPst I and aguC -ASSst I to generate a secondary PCR product corre sponding to the entire length of aguC . The PCR fragment was then cloned onto pGEM 5zf(+) and electroporated into E. coli DH10B. A promoterless kanamycin resistance cassette (KmR), from Tn 1545 and lacking a terminator (Perez-Casal et al. , 1991), was inserted at the Sma I restriction site to disrupt aguC . The construct was integrated into the S. mutans chromosome via natural
23 transformation (Perry and Kuramitsu, 1981) se lecting for growth on BHI agar containing 1.0 mg mL-1 of Km, and correct integration was c onfirmed by Southern blot analysis. The aguR deletion mutant ( aguR ::Em) was constructed by PCR ligation mutagenesis (Lau et al. , 2002). Primers aguR -S and aguR -ASHind III were used to amplify a 0.6-kbp region upstream of aguR . Primers aguR -SSst I and aguR -AS were used to amplify a 600-bp region downstream of aguR . The PCR products were digested with Hind III and Sst I, respectively, and ligated to the erythromycin resistance cassette (EmR) derived from Tn 916 delta E (Rubens and Heggen, 1988). The ligation mixture was introduced into S. mutans UA159 by natural transformation, and bacteria were plated on BHI agar containing 8 g mL-1 of Em. Double crossover mutants were confirmed by Southern blot and PCR. S. mutans aguD was mutated by allelic replacement with KmR ( aguD ::Km) using PCR ligation mutagenesis. Primers aguD -S and aguD Sma I-AS were used to amplify a 600-bp region upstream of aguD . Primers aguD Sma I-S and aguD -AS were used to amplify a 600-bp region downstream of aguD . The PCR products were digested with Sma I and ligated to KmR. The ligation mixture was introduced into S. mutans UA159 by natural transformation, and bacteria were plated on BHI agar containing 1 mg mL-1 of Km. Double crossover mutants were confirmed by PCR. S. mutans oppA was mutated by allelic replacement with KmR ( oppA :: Km) using PCR ligation mutagenesis. Primers oppA -540-S and oppA -1340Sma I-AS were used to amplify an 800-bp region upstream of oppA . Primers oppA -2980Sma I-S and oppA -3800-AS were used to amplify an 800-bp region downstream of oppA . The PCR products were digested with Sma I and ligated to KmR. The ligation mixture was
24 introduced into S. mutans UA159 by natural transformation, and bacteria were plated on BHI agar containing 1 mg mL-1 of Km. Double crossover mutants were confirmed by Southern blot and PCR. S. mutans adiR was mutated by disruption with a spectinomycin resistance cassette, generating adiR ::Sp. The 5 half of S. mutans adiR12 was amplified via recombinant PCR using primers U1640Aat II-S and U2060Sph I-AS, which inserted Aat II and Sph I sites, respectively. The 3 half of adiR12 was amplified using primers U2060Sph I-S and U2480Sst I-AS, which inserted Sph I and Sst I sites. The two primary PCR products were amplified in a secondary reacti on using the end primers, U1640Aat II-S and U2480Sst I-AS. The resulting product was digested with Aat II and Sst I, then ligated into pGEM-7Zf(+) at Aat II and Sst I sites, generating pAG61. The spectinomycin resistance cassette (SpR) from pGEM7-Sp (laboratory stock) was inserted into the engineered Sph I site, disrupting adiR12 and generating pAG62. p AG62 was introduced into S. mutans UA159 via natural transformation, and bacteria were plated on BHI ag ar containing 1 mg mL-1 of spectinomycin. The double cross mutant, AG63, was confirmed by PCR analysis. To complement the adiR12 mutation, the S. rattus FA-1 adiR12 structural gene, including 215 bases upstream and 100 bases downstream, was amplified using PCR primer pair F8040Bam HI-S and F8920Pst I-AS (Figure 2-1). The resulting PCR product was digested with the Bam HI and Pst I restriction enzymes and ligated into the corresponding restriction s ites in pGEM3z(+). SpR was inserted at a Sph I site. The adiR12 -SpR fragment was released from the plasmid by digestion with Bam HI and Hind III and was gel purified. To replace S. mutans adiR (including 215-bp upstream and
25 100-bp downstream) with S. rattus adiR and the flanking regions, three-way ligation mutagenesis was employed using the adiR -SpR fragment and the two arms flanking S. mutans adiR (Figure 2-1). The arm 5 to adiR12 in S. mutans was amplified using the PCR primer pair U2755Bam HI-S and U3355-AS. The arm 3 to adiR in S. mutans was amplified using primer pair U977S and U1577Hind III-AS. The arms were digested with Bam HI and Hind III, respectively, and ligated to the S. rattus adiR PCR product. The ligation mixture was transferred to S. mutans UA159 via natural transformation and cells were plated on BHI ag ar containing 1 mg mL-1 of spectinomycin. The double crossover mutant strain, AM82, was confirmed by PCR analysis. The S. mutans flp deletion mutant ( flp Em) was constructed by PCR ligation mutagenesis. Primers U2160-S and U2780Hind III-AS were used to amplify a 0.6-kbp region upstream of flp . Primers U3740Sst I-S and U4280-AS were used to amplify a 600-bp region downstream of flp . The PCR products were digested with Hind III and Sst I, respectively, and ligated to EmR. The ligation mixture was introduced into S. mutans UA159 by natural transformation and bacteria were plated on BHI agar containing 8 g mL-1 of Em. Double crossover mutant s were confirmed by PCR. The ligation mixture was also introduced into AG63 and AM82 via natural transformation to generate a double flp adiR12 mutant strain, AG68, and a double mutant strain complemented with S. rattus adiR , AM94, respectively. RNA Manipulations RNA was prepared from S. mutans UA159, MJL1, and S. rattus FA-1 cells and immediately treated with the RNAprotect reagent from Qiagen (Qiagen Inc., Valencia, CA). Total RNA was isolated using protocols described elsewhere (Chen and Burne,
26 1996). The RNA was further pu rified, treated with DNaseI using the RNeasy RNA Clean Up mini kit from Qiagen and stored at -80 C. Primer sequences ar e listed in Table 2-1. Streptococcus rattus FA-1 Primer extension analysis was used to map the arcA transcription initiation site. Primer PE arcA -S encoded the antisense sequence of arcA located 50-b downstream from the translational start site. Incuba tion of radiolabeled primers with 50 g of total RNA at 42C for 90 minutes was followed by revers e transcription and the products were separated by electrophoresis and disclose d by autoradiography. A DNA sequencing reaction using the same primer was included on the gel to allow identification of the start site. To determine if the arc genes of S. rattus FA-1 could be co-transcribed, reverse-transcriptase (RT) PCR was perf ormed using the Supe rScript First-Strand Synthesis System (Invitrogen, Carlsba d, CA). RNA was isolated from S. rattus FA-1 grown in TV + 2% galactose and 1% argi nine. PCR amplification of the cDNA was performed using various primer pairs (RT arcAB -S and RT arcAB -AS, RT arcDT -S and RT arcDT -AS, RT arcTadiR -S and RT arcTadiR -AS) flanking the intergenic region of arcAB , arcDT and arcTadiR , respectively. Streptococcus mutans UA159 To assess transcription of the flp , adiR , oppA , oppB , SMU.260 and agu genes in the S. mutans wild-type and/or aguR strains, the SuperScript Fi rst-Strand Synthesis System (Invitrogen, Carlsbad, CA) was used to c onduct reverse transcri ptase-PCR (RT-PCR) analysis. First-strand cDNA s ynthesis was generated from 1 g of total RNA using random hexamers as recommended by the s upplier (SuperScript First-Strand Synthesis
27 System for RT-PCR; Invitrogen, Carlsbad, CA). PCR amplification of the first strand aguB, flp and adiR cDNA was performed using primer pairs RT flp -S and RT flp -AS, RT adiR1 -S and RT adiR1 -AS, and RT adiR2 -S and RT adiR2 -AS, respectively. PCR amplification of the first strand oppA , oppB and SMU.260 cDNA was performed using primer pairs RT oppA -S and RT oppA -AS, RT oppB -S and RT oppB -AS, and RTsmu.260-S and RTsmu.260-AS. For analysis of aguA expression in response to growth at pH 5 versus pH 7, the aguA -specific primers, RT aguA -S and RT aguA -AS were used for both cDNA generation and first-strand amplificati on. All gene-specific RT-PCR primers were designed using Beacon Designer 2.0 software (P remier Biosoft International, Palo Alto, CA). Standard curves for each gene, prepared as described by Yin et al. (Yin et al. , 2001), were used in every run. A range of 101 to 108 copies was found to be adequate for all genes examined. Primer extension analysis was used to map the aguB and aguR transcription initiation sites. Primer Pe aguB -AS encoded the antisense sequence of aguB located 30 bases downstream from the translational start site. Primer PE aguR -AS encoded the antisense sequence of aguR located 25 bases downstream from the translational start site. Incubation of radiolabeled primers with 50 g of total RNA at 42C for 90 minutes was followed by reverse transcription and the pr oducts were separated by electrophoresis and disclosed by autoradiography. DNA sequencing reactions using the same primers were included on the gel to allow identif ication of the start sites. Comparisons of the amounts of AgDS mRNA in S. mutans UA159 cells grown under different growth conditions was done by sl ot blot analysis. Samples containing 1 or 2 g total RNA, as well as a 2 g sample of RNaseA-tre ated RNA as a negative
28 control, were UV-crosslinked to a 0.45 micron nitrocellulose membrane. The DNA probe was labeled using the Ambion Bright St ar Psoralen-Biotin nonisotopic labeling kit (Ambion Inc, Austin, TX). Hybridizations were carried out as recommended by the supplier at high stringency conditions. Protein Manipulations Construction of N-terminal 6X-His-tagged AdiR The adiR structural gene was PCR amplified from the S. rattus chromosome using the primer pair AdiRpET-PstI-S and AdiR pET-HindIII-AS, and digested with the Pst I and Hind III restriction enzymes. The resulti ng 440-bp fragment was purified from a 1.6% TAE agarose gel and ligated, in-frame, into Pst I and Hind III sites on the pET-45b(+) vector from Novagen (Madison, W I). The pET-45b(+) vector contains an N-terminal 6X-His-tag, located upstream of the multiple cloning site, and protein expression is controlled by the bacteri ophage T7 promoter. The pET-45b(+)/ adiR construct was electroporated into E. coli BL21 competent cells, as indicated by the manufacturer, and plated ont o LB agar containing 100 g mL-1 of ampicillin. A clone harboring the correct constr uct, MJL2, was identified by restriction digestion with Pst I and Hind III, followed by sequence analysis using the T7 Terminator Primer supplied by the manufacturer (Novagen, Madison, WI). To express His-tagged AdiR in E. coli , MJL2 was electroporated into the BL21-DE3 expression strain, which supplies the T7 RNA polymerase gene on the DE3 lysogen. A clone harbor ing the correct construct, MJL3, was isolated and confirmed usi ng restriction digestion analysis. AdiR Protein Purification To induce expression of AdiR, MJL3 wa s grown in 500 mL of LB medium containing 100 g mL-1 of ampicillin, at 37 C with shaking, to an OD600 of 0.5 and
29 induced with 1 mM iso-propylthio-D-galactopyranoside (IPTG). After three hours, cells were harvested by cen trifugation at 9000 x g at 4 C, for 30 minutes, and the pellet was stored overnight at C. Cell pellets were thawed on ice and resuspended in native lysis buffer containing 1 mM imidazole. Ce lls were sonicated on ice in 15 second intervals for a total of 2 minutes, fo llowed by centrifugation at 9000 x g at 4 C, for 30 minutes. The lysate was mixed 1:1 with Ni -NTA agarose resin (Qiagen, Valencia, CA) and was rocked continually at 4 C for 30 minutes. The lysate:resin mixture was then loaded onto a glass Econo-Column (1 x 10 cm ; BioRad, Hercules, CA). Flow through, wash and elution fractions were collected at 4 C. All buffers were prepared as the manufacturer instructed (Novagen, Madison, WI). Gel Shift Assays Overlapping 30-mer oligos span ning the 200 base region upstream of the S. rattus arcA transcription initiation site (Table 2-5) were synthesi zed by Integrated DNA Technologies (Coralville, IA). Oligos were end-labeled with [ -32P]-ATP using T4 polynucleotide kinase and were purified us ing the QIAquick Nucleotide Removal Kit (Qiagen). Binding reactions cont ained 0.5 ng DNA probe, 2 g nonspecific competitor DNA [poly (dIdC)], 300 g mL-1 of bovine serum albumin (BSA), 5 mM MgCl2, either 0 or 10 mM arginine-HCl, 1X binding buffer ( 10 mM Hepes, pH 7.9, 50 mM KCl, 1 mM EDTA, 5 mM DTT, and 10% glycerol), and 400 pmol (~6.8 g) purified His-tagged AdiR, in a total volume of 10 l. To confirm the specificity of probes VI and VII, 0.4 (1:1), 40 (1:100) or 400 (1:1000) pmol of th e cold (unlabeled) VI and VII oligos were added to the binding reactions in a series of competition experiments, bringing the total
30 volume to 15 l. All reactions were performed for 30 minutes at room temperature, then loaded directly onto a 4% polyacrylamide [( 30:1) acrylamide:bis-acrylamide] low ionic strength gel, and electrophoresed at 25 mA for approximately 4 hours. Expression of S. rattus Flp in S. mutans The flp structural gene was PCR amplified from the S. rattus chromosome using primer pair NicFlpBam HI-S and NicFlpPst I-AS and was digested with Bam HI and Pst I restriction enzymes. The resulting 6 58-bp product was purified from a 0.8% TAE agarose gel and ligated into the Bam HI and Pst I sites on the nicin-inducible expression vector, pMSP3535 (Bryan et al. , 2000). This vector contains the ColE1 origin of replication, in addition to re plication genes and antibiotic resistance markers compatible with Gram-positive bacteria. Protein expression is regulated by the pnisA promoter, which requires nicin for activation. The resu lting construct was electroporated into E. coli DH10B competent cells and plated on LB agar containing 100 g mL-1 of ampicillin and 300 g mL-1 of erythromycin. A clon e containing the correct construct, pNicinFlp, was confirmed by restriction enzyme digestion with Bam HI and Pst I, as well as sequence analysis. S. mutans AM72 was transformed with pNicinFlp, generating the nicin-inducible Flp-expressing strain, AM100. As a control, AM72 was transformed with the empty pMSP3535 vector, generating strain AM99. CAT activity was measured in AM99 and AM100 as described below, except that 25 ng mL-1 of nicin and 10 g mL-1 of erythromycin were added to the growth media. Construction of Promoter Fusions and CAT Assays The S. rattus FA-1 arcA promoter and deletion derivatives were amplified via recombinant PCR (Higuchi, 1990) using sense primers ( arcA Sac I-S-400,
31 arcA Sac I-S-150, and arcA Sac I-S-100) in conjunction with the antisense primer arcA Bam HI-AS. The S. mutans UA159 aguB promoter and deletion derivatives were amplified using sense primers ( aguB Xba I-S-41, aguB Sac I-S-100, and aguB Sac I-S-157) in conjunction with the antisense primer aguB Bam HI-AS. These primers allowed insertion of Sac I or Xba I and Bam HI restriction sites to facilitate cloning. The PCR products were ligated to the 5 end of a promoterless chloramphenicol acetyltransferase gene ( cat ) from Staphylococcus aureus (Ehrlich, 1978), and cloned onto the integration vector pMJB8A (Chen et al. , 2002). The constructs were then electroporated into E. coli DH10B, screened for the corr ect configurations and then introduced into S. gordonii DL-1, S. mutans UA159 and mutant strains by natural transformation. Cultures used in CAT assays were grown in TV medium containing 2% glucose or galactose, with or without 1% ar ginine, to an optical density at 600 nm = 0.6. Biochemical Assays S. rattus FA-1 was grown in TV medium supplemented with 0.2 or 2% glucose or galactose 1% argini ne at 37C in 5% CO2 and 95% air. AD activity was measured by monitoring production of citrul line from arginine as previo usly described (Archibald, 1944). Chloramphenicol acetyltransferas e assays were performed using the spectrophotometric method of Shaw (Shaw, 1979). Enzyme activities were normalized to protein concentration, which was determ ined by the method of Bradford (Bradford, 1976) using a kit (BioRad) with BSA as the standard. AgD activity was measured by colorimetric determination of N-carbamoylputrescine production from ag matine using the method of Archibald (Archibald, 1944). S. mutans and S. rattus strains were grown in BHI supplemented with 10 mM agmatine or TV medium containing 2% glucose or galactose, with or without
32 10 mM agmatine, to OD 600 nm = 0.6. Cells were harvested by centrifugation, washed once with 10 mM Tris-maleate buffer, pH 6.0, and resuspended in 1/ 10 of the original culture volume in the same buffer. The cel ls were permeabilized using 1/20 volume of toluene and two, one-minute freeze-thaw cycl es. The cell suspension was centrifuged and the pellet was resuspended in 500 l of 10 mM Tris-maleate, pH 6.0. A 50 l aliquot of the cell suspension was used in a 500 l reaction mixture, cont aining 10 mM agmatine or 50 mM arginine when specified. After 30 minutes, reactions were terminated by the addition of an equal volume of 10% trichlor oacetic acid and N-carbamoylputrescine was measured. The protein concentration of the cell suspension was determined as follows. A known volume of the cell suspension was mi xed with an equal volume of glass beads (0.1 mm) and homogenized using a Bead B eater. The samples were centrifuged for 10 min in a refrigerated microcentrifuge and the protein concentration of the lysate was measured using a protein assay (Bio-Ra d, Hercules, CA) based on the method of Bradford (Bradford, 1976), with bovine albumin serum as the standard. AgD activity was expressed as nmol N-carbamoylputrescine min-1 (mg protein)-1. Ammonia production from agmatine was measured in intact cells of S. mutans UA159 and S. rattus FA-1 grown to mid-exponential phase (OD600 0.6). The cells were collected by centrifugation, wa shed with potassium phosphate buffer, pH 7.0, and resuspended in 1/10 the original culture volume of the same bu ffer. A 10 l aliquot of the cell suspension was used in a reacti on mixture containing 50 mM potassium phosphate, pH 7.5, and 10 mM agmatine. The reaction was carried out at 37 C for 30 minutes. Initially, we attempted to m easure ammonia production using the Nessler reagant from Sigma (St. Louis, MO) but ad dition of agmatine to the reaction mixture
33 caused the Nesslerâ€™s reagant to precipitat e. Consequently, ammonia production was measured using an Ammonia Detection kit (Diagnostic Chemicals Limited, Charlottetown, Canada), which allows de termination of ammonia concentration by monitoring the rate of NADP production in a glutamate dehydrogenase-catalyzed reaction: NH4 + + -ketogluterate + NADH L-glutamate + NADP+ + H20. The protein concentration of cell suspensions was de termined as described above. Ammonia production was expressed as nmol NH4 + min-1 (mg protein)-1. Ammonia production by intact cells of S. mutans at various pH values were carried out as described above, except different bu ffers were used (glycine-HCl < pH 4.5; Tris-maleate > pH 4.5 to 7; potassium phospha te buffer > pH 7.0). Reactions were carried out for 60 minutes. Appropriate contro ls were conducted to en sure that inclusion of different buffers did not alter the ability to measure ammonia using the coupled assay. Acid Killing Assays Overnight cultures of S. mutans wild-type and aguB strains grown in TV media containing 25 mM galactose were reinocul ated 1:40 into fresh TV media containing 25 mM galactose with 0 mM or 10 mM ag matine. Cells were harvested at OD600 = 0.5, centrifuged for 10 minutes at 4 C, 4000 rpm. Pellets were washed with 1 mL of 0.1 M glycine buffer, pH 7, and centrifuged as be fore. Pellets were resuspended in 0.1 M glycine buffer, pH 2.8, containing 0 or 10 mM ag matine, an initial sample was taken, and the tubes were placed on a rocker at room temperature. Subsequent samples were taken at 30, 60 and 90 minutes. Serial dilutions of each sample were plated on BHI agar and incubated at 37 C with 5% CO2 for 48 hours.
34 Nucleotide Sequence Accession Numbers The complete sequences of the arc operon and adiR have been deposited in the GenBank database, and the accession number is AY396288. The complete sequences of the agu operon and aguR have been deposited in the GenBank database, and bear accession number BK004003.
35 Table 2-1. Primers used in this study Primer Sequencea Application arcB -S 5 -CAAGTATTTCAGGGACGC-3 arcB probe arcB -AS 5 -CATCTGTCAAGCCATTCC-3 arcB probe aguB -SXba I 5 -CAGATTATA TCTAGA CAGAGGATTT-3 Inactivation of aguB aguB -ASEco RI 5 -TACCAGCTGG GAATTC TTCTATCATTGTA-3 Inactivation of aguB aguB -SEco RI 5 -TACAATGATAGAA GAATTC CCAGCTGGTA-3 Inactivation of aguB aguB -ASSst I 5 -ACCGTCCAT GAGCTC ATCTGTAATCT-3 Inactivation of aguB aguB -S 5 -CAGATTATATCTAGACAGAGGATTT-3 aguB probe aguB -AS 5 -CATCTGTCAAGCCATTCC-3 aguB probe aguC -SPst I 5 -GTCTAGAGA CTGCAG TGCCAAAGCACA-3 Inactivation of aguC aguC -ASSma I 5 -TTTTCGCCAACCTCT CCCGGG ATCTACTTT-3 Inactivation of aguC aguC -SSma I 5 -AAGTAGAT CCCGGG AGAGGTTGGCGAAAA-3 Inactivation of aguC aguC -ASSst I 5 -GGCTTTTCCACT GAGCTC TGCTTCAAC-3 Inactivation of aguC aguR -S 5 -CGTTCTTTTCCTGCAGGACTCTCAAG-3 Inactivation of aguR aguR -ASHind III 5 -CGTAAATTG AAGCTT TTCCTAAACTGAC-3 Inactivation of aguR aguR -SSst I 5 -CTCCTTTAATTT GAGCTC AATATCTATAGT-3 Inactivation of aguR aguR -AS 5 -GATATCATCCAATCTAGAAAGAACAGTTG-3 Inactivation of aguR PE arcA -S 5 -GACGATGTAACATTACCTTCTT-3 Primer Extension RT arcAB -S 5 -AGCTAGGAAACTGCGTCCCT-3 RT-PCR RT arcAB -AS 5 -TTTAGACTCTTTACAGGACAGATT-3 RT-PCR RT arcDT -S 5 -TTTAGACTCTTTACAGGACAGATT-3 RT-PCR RT arcDT -AS 5 -TGAATATTCATCTGTTTACCCCTT-3 RT-PCR RT arcTadiR -S 5 -AGTGAGTTGTCTGAGTTTCTA-3 RT-PCR RT arcTadiR -AS 5 -TTTATCTTACTTTGGCGCAATA-3 RT-PCR RT aguB -S 5 -CAGATTATATCTAGACAGAGGATTT-3 RT-PCR RT aguB -AS 5 -TACCAGCTGGGAATCCTTCTATCATTGTA-3 RT-PCR PE aguB -AS 5 -TCCTCTGTCGTAATATAATCTGT-3 Primer Extension PE aguR -AS 5 -ATAGATTATAGATATAGATGAGTTC-3 Primer Extension RT aguA -S 5 -ATGCTTGGATTCGTGACTGTGG-3 RT-PCR RT aguA -AS 5 -AAGACCATCGACTAAGCCTCCC-3 RT-PCR arcA-Sac I-S400 5 -TTGCTCTA GAGCTC TCAAATGACAGAA-3 parcA 340 Amplification arcA Sac I-S150 5 -TTATAAATTC GAGCTC CAAAAAACGTGAA-3 parcA 80 Amplification arcA Sac I-S100 5 -TAAATAACAATTC GAGCTC GAAAAAAAT CTTA-3 parcA 50 Amplification arcA Bam HIAS 5 -TTTTGAGTCAT GGATCC TACTCCTTTCGAT-3 parcA Amplification RT flp -S 5 -ATCGCTTTCTAACTGGCTGGC-3 RT-PCR RT flp -AS 5 -TTGGGAGAGGCA GATGTTTTGG-3 RT-PCR RT adiR1 -S 5 -ATCGCTTTCTAACTGGCTGGC-3 RT-PCR RT adiR1 -AS 5 -TTGGGAGAGGCAGATGTTTTGG-3 RT-PCR RT adiR2 -S RT adiR2 -AS aguA -S 5 -AGCGTCTTGTTGATAGCCTTTC-3 5 -ATGAGCCCATTTGAAGAAGCC-3 5 -GCCTGGGGAGGCTTAGTCG-3 RT-PCR RT-PCR aguA probe
36 Table 2-1. Continued Primer Sequencea Application aguA -AS 5 -CAGTGAATATTGCCACCACC-3 aguA probe AdiRpETPst I-S 5 -GGGACTGCAGATGAATAAATTATTGC-3 AdiR Expression AdiRpETHind III-AS 5 -CAAATGAAGCTTTAAGCTGGACAG-3 AdiR Expression NicFlpBam HI-S 5 -CTAAAGAT GGATCC CAAATGTTGCG-3 Flp Expression NicFlpPst I-AS 5 -GTTTCCGTCAATCC CTGCAG TAAAT-3 Flp Expression U1640Aat II-S 5 -GGAGAGAG GACGTC CTTAG-3 Inactivation of adiR12 U2060Sph I-AS 5 -CCTTTTGGTTATTTG GCATGC TGAAGAAA-3 Inactivation of adiR12 U2060Sph I-S 5 -TTTCTTCA GCATGC CAAATAACCAAAAGG-3 Inactivation of adiR12 U2480Sst I-AS 5 -GAAGTATGGTAT GAGCTC TATATGTAGGA-3 Inactivation of adiR12 F8040Bam HI-S 5 -GTTATCTATGGGT GGATCC TGTCAT-3 adiR Complementation F8920Pst I-AS 5 -CCAGTTTCTCCGCC CTGCAG TCTGCT-3 adiR Complementation U977-S 5 -GACTCAACTCACGTTTTTGTGTG-3 adiR Complementation U1577Hind IIIAS 5 -GATGCTATCAAGGC AAGCTT AATTTTAGA-3 adiR Complementation U2755Bam HI-S 5 -CGTAAAGGGTATA GGATCC TCAGAACAG-3 adiR Complementation U3355-AS 5 -GATAAGGAGAGTGATAAGATGTTA-3 adiR Complementation U2160-S 5 -CCAAAACATCTGCCTCTCCCAA-3 Inactivation of flp U2780Hind IIIAS 5 -TAGATTGTCAGAG AAGCTT GCTGTTC-3 Inactivation of flp U3740-SstI-S 5 -CTAATCTCAAATC GAGCTC AAAAGGTTC-3 Inactivation of flp U4280-AS 5 -CAATTAATTCTGCCTCCTTATGTC-3 Inactivation of flp RT oppA -S 5 -TCAGGTCTTGCTGTTGAGTCTC-3 RT-PCR RT oppA -AS 5 -TGAGGAGTCGGTGATTCTAAGG-3 RT-PCR RT oppB -S 5 -ATGGGTTTGATTGTAGGTGCTC-3 RT-PCR RT oppB -AS 5 -AGGTCGGCAGAATGGTTTGAG-3 RT-PCR RTsmu.260-S 5 -CGTCGCAGTATCTATGCCTTAG-3 RT-PCR RTsmu.260-AS 5 -CTTGAGAGTCTTGCCCGAAAAG-3 RT-PCR aguD -S 5 -GTGTTGAACGTCACCAAAGTG-3 Inactivation of aguD aguD Sma I-AS 5 -CTTTCCTTCCATA CCCGGG TTCCTTTCTT-3 Inactivation of aguD aguD Sma I-S 5 -TAGGTGAACTTATTATTATCAT CCCGGG AAT-3 Inactivation of aguD aguD -AS 5 -CGTCCCTTGGCATCTGTCTG-3 Inactivation of aguD oppA -540-S 5 -GAGTCGGGATAGATATGATCG-3 Inactivation of oppA oppA -1340Sma I-AS 5 -CTTTTCTTTATTGCTATCAT CCCGGG TCCT-3 Inactivation of oppA oppA -2980Sma I-S 5 -CTTCACCTATGCTTAT CCCGGG TAACTTTT-3 Inactivation of oppA oppA -3800-AS 5 -CCATAGGACCCACTAAAGTCA-3 Inactivation of oppA aguB Xba I-S41 5 -TGTAATCGTTTACA TCTAGA AGTTTATAGT-3 PaguB -41 Amplification aguB Sac I-S100 5 -GGGTTTATTTTT GAGCTC CGGTTTATTTC-3 PaguB -100 Amplification aguB Sac I-S157 5 -GTAATTATATCATTCA GAGCTC CTTTTATTGA-3 PaguB -157 Amplification aguB Bam HIAS 5 -TTTTTTCATCAT GGATCC CTCCTCTATTTTT-3 PaguB Amplification a The introduced restriction recognition sites within primers are indicated in boldface characters.
37 Table 2-2. Plasmids used in this study Plasmid Phenotype or description Reference or source pSU20 Cmr; general cloning vector Bartolome et al. (1987) pUC18/pUC19 Apr; general cloning vector Invitrogen pUC18erm Apr, Emr; pUC18 carrying the Emr cassette from pCER1000 Claverys, et al. (1995) pCRTMII Apr, Kmr; general cloning vector Invitrogen pJZ22 Apr, Kmr; pCRTMII carrying 0.5-kbp fragment encoding internal fragment of arcB of S. rattus This study pvT924 Kmr; plasmid encoding the polar kanamycin gene from Perez and Casal (1991) pALH123 Kmr; plasmid encoding the non-polar kanamycin gene from Tn1545 Perez (1991) pGEM-3Zf(+) Apr; general cloning vector Promega pGEM-5Zf(+) Apr; general cloning vector Promega pGEM-7Zf(+) Apr; general cloning vector Promega pGEM7-Spc Apr, Spr; pGEM-7Zf(+) carrying the Spr cassette from pSN13 Lab stock pC194 Cmr; plasmid encoding a promoterless cat gene originally isolated from Staphylococcus aureus Horinouchi and Weisblum (1982) pMC286 Apr; promoterless cat gene was cut from pC194 and cloned into the Bam HI and Sph I sites of pGEM-3Zf(+) Lab stock pMJB8 Apr, Kmr; integration vector for S. gordonii DL-1 that allows insertion of foreign DNA at the gtfG locus Chen et al. (2002) pMC341B Kmr, Emr; integration vector for S. mutans UA159 that allows insertion of foreign DNA at the mtlA locus Lab stock pET45b(+) Apr; N-terminal 6x His-tagged protein expression vector Novagen pAG1 Apr; pGEM-7Zf(+) with S. mutans aguB inserted at Xba I and Sst I recognition sites This study pAG5 Apr; pGEM-5Zf(+) with S. mutans aguC inserted at Pst I and Sst I recognition sites This study pAG6 Apr; pGEM-5Zf(+) with S. mutans oppB inserted at Pst I and Sst I recognition sites This study pAG9 Apr, Kmr; pAG1 with Kmr cassette inserted at Eco RI recognition site This study pAG13 Apr, Kmr; pAG5 with Kmr cassette inserted at Sma I recognition site This study pAG14 Apr, Kmr; pAG6 with Kmr cassette inserted at Sma I recognition site This study pAG61 Apr; pGEM3 with S. mutans adiR12 inserted at Hind III and Sst I recognition sites This study pAG62 Apr; pAG61 with Spr cassette inserted at Sph I recognition site This study pAM3 Apr; S. rattus parcA 340 inserted into pMC286 at Bam HI and Sst I recognition sites This study pAM4 Apr, Cmr; S. rattus parcA 80 inserted into pMC286 at Bam HI and Sst I recognition sites This study pAM9 Cmr, Kmr; S. rattus parcA 80 inserted into pMJB8a at Sma I and Sph I recognition sites This study pAM12 Apr, Kmr; pAM3 with Kmr cassette inserted at blunt Sst I recognition site This study pAM17 Apr, Kmr; S. rattus parcA 340 inserted into pmjb9a at Sma I site This study
38 Table 2-2. Continued Plasmid Phenotype or description Reference or source pAM23 Apr, Cmr; S. rattus parcA 50 inserted into pMC286 at Bam HI and Sst I recognition sites This study pAM34 Cmr, Kmr; S. rattus parcA 50 inserted into pmjb8a at Sma I and Sph I recognition sites This study pAM48 Cmr, Kmr; S. rattus parcA 80 inserted into pMC341B at Sst I and Sph I recognition sites This study pAM50 Cmr, Kmr; S. rattus parcA 50 inserted into pMC341B at Sst I and Sph I recognition sites This study pAM71 Apr; S. rattus flp inserted into pGEM-7Zf(+) at Hind III and Eco RI recognition sites This study pAM75 Apr, Emr; pAM71 with Emr cassette inserted at Sst I and Eco RI recognition sites This study pAM77 Apr; S. rattus parcA 386 inserted into pMC286 at Sst I and Bam HI recognition sites This study pAM78 Apr, Kmr; S. rattus parcA 386 inserted into pMC340B at Sst I and Sph I recognition sites This study pAM80 Apr; S. rattus adiR12 inserted into pGEM-3Zf(+) at Bam HI and Pst I recognition sites This study pAM81 Apr, Spr; pAM80 with Spr cassette inserted at Sph I recognition site This study pMSP3535 Apr, Emr; Nisin-controlled protein expression vector for Grampositive bacteria Bryan et al. (2000) pNicinFlp Apr, Emr; S. rattus flp structural gene inserted into pMSP3535 This study pDL278 Spr; E. coli-Streptococcus spp. shuttle vector LeBlanc (1992)
39 Table 2-3. Strains us ed in this study Strain Phenotype or description Reference or source E. coli strains: DH10B General cloning strain Invitrogen BL21 General purpose host Novagen BL21 DE3 General purpose protein expression host Novagen MJL2 BL21/pET45b(+) expressing S. rattus AdiR This study MJL3 BL21-DE3/MJL2 This study S. rattus strain: FA-1 Wild-type host ATCC 19645 S. gordonii strains: DL1 Wild-type host ATCC 49818 AM13 S. rattus parcA 80 integrated into gtfG locus of DL1 This study AM19 S. rattus parcA 340 integrated into gtfG locus of DL1 This study AM43 S. rattus parcA 50 integrated into gtfG locus of DL1 This study S. mutans strains: UA159 Wild-type host ATCC 700610 TW01 ccpA ::Km Wen and Burne (2002) TW02 ccpB ::Em Wen and Burne (2002) JLCtsR ctsR ::Km Lemos and Burne (2002) SM11 hrcA ::Km Lemos et al. (2001) SM12 dnaK ::Km Lab stock, unpublished JLClpP clpP ::Km Lemos and Burne (2002) MJL1 aguR ::Em This study AG17 aguB :: Km This study AG21 aguC ::Non-polar Km This study AG35 S. mutans paguB-41 integrated into mtlA locus of UA159 This study AG37 S. mutans paguB-100 integrated into mtlA locus of UA159 This study AG39 S. mutans paguB-157 integrated into mtlA locus of UA159 This study AG41 S. mutans paguB-41 integrated into mtlA locus of MJL1 This study AG43 S. mutans paguB-100 integrated into mtlA locus of MJL1 This study AG45 S. mutans paguB-157 integrated into mtlA locus of MJL1 This study AG49 aguR ::Em , aguB :: Km This study AG52 S. rattus parcA 386 integrated into mtlA locus of AG50 This study AG53 S. rattus parcA 386 integrated into mtlA locus of UA159 This study AG56 oppA :: Km This study AG57 oppB :: Km This study AG58 SMU.260 (putative nitroreductase):: Km This study AG59 aguD ::non-polar Km This study AG63 adiR12 ::Sp This study AG64 S. rattus parcA 80 integrated into mtlA locus of AG63 This study AG65 flp ::Em This study AG66 S. rattus parcA 80 integrated into mtlA locus of AG65 This study
40 Table 2-3. Continued Strain Phenotype or description Reference or source AG68 S. rattus parcA 80 integrated into mtlA locus of flpadiR12 This study AM64 S. rattus parcA 80 integrated into mtlA locus of UA159 This study AM72 flp ::Sp This study AM82 adiR12 ::FA-1 adiR12 + Sp This study AM83 S. rattus parcA 80 integrated into mtlA locus of AM82 This study AM93 S. rattus parcA 80 integrated into mtlA locus of AM76 This study AM94 S. rattus parcA 80 integrated into mtlA locus of flpadiR12 ::FA-1 adiR12 + Sp This study AM99 AM72 harboring pMSP3535 This study AM100 AM72 harboring pNicinFlp, nicin inducible This study
41 Table 2-4. Primers used to sequence the S. rattus agu operon Primera Sequence Loci 5-S 5 -TATTTCCAATTTACGGGTGTTCT-3 aguR 100-S 5 -GTGAATGTGAGTTTTTACTGTGC-3 aguR 560-S 5 -TTGATTTGGTAGGTAATAGAGGT-3 aguR 1050-S 5 -GGCTTTGTAAAAAAGGCATAAAC-3 aguR-aguB 1760-S 5 -GAATGGAATTTGTTCACTTTGGA-3 aguB 2300-S 5 -CGCAATCATGTCTGTCCTAAAC-3 aguB-aguD 3080-S 5 -GCTTTTGGTATTGGCGTCTCA-3 aguD 3540-S 5 -CGTCCTTTTAAGGTTAGTGGC-3 aguD-aguA 4260-S 5 -GTGTCTGTTACATCTTAGTCGG-3 aguA 4840-S 5 -CAGTGAAAATGGAGAAAATGTATG-3 aguA-aguC 5340-S 5 -GAGGTTGGCGAAAAGTAGTTG-3 aguC 560-AS 5 -TACCTCTATTACCTACCAAATCAA-3 aguR 1050-AS 5 -GTTTATGCCTTTTTTACAAAGCC-3 aguR 1760-AS 5 -GTCCAAAGTGAACAAATTCCA-3 aguR-aguB 2300-AS 5 -GTTTAGGACAGACATGATTGCG-3 aguB 3080-AS 5 -GAGACGCCAATACCAAAAGCA-3 aguB-aguD 3540-AS 5 -GCCACTAACCTTAAAAGGACG-3 aguD 4260-AS 5 -CCGACTAAGATGTAACAGACAC-3 aguD-aguA 4840-AS 5 -CATACATTTTCTCCATTTTCACTG-3 aguA 5340-AS 5 -CAACTACTTTTCGCCACCTC-3 aguA-aguC 5780-AS 5 -CCGTAATTTGTGTGCCACTTC-3 aguC aNumber denotes the position of the primer relative to the 5 end of the agu operon
42 Table 2-5. Oligos used in gel sh ift experiments with purified AdiR Designation Sequence Positiona I 5 -ATGATTCTCAAAATCGAAAGGAGTAGTTAACATG-3 +3 to -32 II 5 -TTGTAATAAATGAGAAATTAATGATTCTCAAAATCGAAAG-3 -12 to -52 III 5 -AAAAACCTCTAAAATGGAATTTGTAATAAATGAGAAATTA-3 -32 to -72 IV 5 -TCATTTATAAAAAACCTCTAAAATGGAAT-3 -52 to -80 V 5 -AATGCGAAAAAAATCTTATTTCATTTATA-3 -72 to -100 VI 5 -ATAACAATTCAAATGCGAAAAAAATCTTATT-3 -80 to -111 VII 5 -CATCAAAAAACGTGAAATAAATAACAATTCA-3 -100 to -131 a Position of each oligo relative to the ATG start codon of S. rattus arcA (positions +1 to +3).
43 Figure 2-1. Complementation of the adiR12 mutation in S. mutans with the adiR12 genes from S. rattus , generating strain adiR12 ::Fa diR12.
44 CHAPTER 3 ISOLATION AND CHARACTERIZATION OF THE ARGININE DEIMINASE OPERON IN S. rattus FA-1 Introduction The ADS is considered a critical factor in oral biofilm pH homeostasis that may inhibit emergence of cariogenic flora (Burne and Marquis, 2000). This pathway is used by many acid-sensitive bacteria a ssociated with dental health to survive acidification of dental plaque by S. mutans and other cariogenic, acid-tole rant bacteria. As detailed earlier, the ADS consists of three enzymes. Arginine is hydrolyzed by AD (ArcA) to generate citrulline and amm onia. Citrulline is then converted to ornithine and carbamoylphosphate via ornithine carbamoyltr ansferase (ArcB). Finally, carbamate kinase (ArcC) transfers a phosphate from carbamoylphosphate to ADP, yielding ATP. Ammonia production from this pathway protects bacteria from lethal acidification and ATP production provides a source of energy for the cells. Previous research has demonstrated that S. rattus FA-1 hydrolyzes arginine via the ADS. Streptococcus rattus is similar to other ADS-positive oral bacteria in that it is acidsensitive compared to S. mutans in the absence of arginine and has not been linked to caries development in humans. However, S. rattus is a member of the mutans group of streptococci and is mo st closely related to S. mutans , which is noted for exceptional cariogenicity and acid tolerance. In f act, a major characteristic distinguishing S. rattus from S. mutans is the ability to catabolize argini ne via the ADS (Coykendall, 1974). The
45 purpose of this study was to identif y the genes encoding the ADS in S. rattus and initiate a molecular characterization of the arc operon. Results Isolation of the ADS Genes of S. rattus FA-1 Prior to my arrival in Dr. Burneâ€™s laboratory, a subgenomic DNA library was created by digesting chromosomal DNA from S. rattus FA-1 to completion using various restriction enzymes. The digested DNA frag ments were separated on a 0.8% agarose gel and screened for the presence of arcB by hybridization with a 0.35-kbp PCR product internal to the S. rattus arcB gene. Results indicated that the arcB gene was contained on an Xba I fragment of approximately 7-kbp. To clone this fragment, a subgenomic DNA library of Xba I fragments was constructed in th e intermediate copy-number plasmid, pSU20 (Bartolome et al. , 1991). The library was screen ed by colony hybridization under stringent conditions with an arcBspecific probe. A positive clone, containing a 7-kbp DNA insert (pJZ29), was identified. Southern blot analysis confirmed that the 7-kbp Xba I DNA fragment originated from S. rattus and demonstrated that the fragment was continuous on the chromosome. Results of DNA sequence analysis performed on the 7-kbp Xba I fragment indicated that this fr agment contained the 3 portion of a partial open reading frame (ORF) that shared homology with other known arcA genes, followed by five complete ORFs. To obtain genomic DNA fragments containing the complete ORF, and potentially identify other genes tightly linked to the ar ginine catabolism cluster, a subgenomic DNA library of Eco RV fragments was constructed and screened with a 1-kbp DNA fragment containing the 3 portion of arcA . A 4-kbp EcoR V fragment was subs equently isolated,
46 and a Southern blot analysis under stringent conditions confirmed that the fragment originated from S. rattus . Nucleotide Sequence Anal ysis of the ADS Genes Using a series of nested deletions genera ted by exonuclease III, the complete sense and antisense nucleotide sequences of the 7-kpb Xba I fragment and 4-kpb EcoR V fragment were determined. The nucleotid e sequences were translated and found to encode six ORFs arranged in an apparent oper on (Figure 3-1). Each ORF began with an ATG start codon and was preceded by a putativ e Shine-Dalgarno sequence. Between the fifth and sixth ORF, a stable stem:loop struct ure that could possibly act as a terminator was identified. Based on similarity to known arc genes, the ORFs were designated arcA, B, C, D, T and adiR. My first task was to perform BLAST searches using the predicted amino acid sequences of the six ORFs to identify seque nce similarity to ot her known proteins. S. rattus AD, encoded by arcA , was 85, 84, 80 and 79% identical to the AD enzymes of Streptococcus agalactiae, Streptococcus pyogenes, Streptococcus pneumoniae and S. gordonii , respectively. S. rattus AD also shared homology with the AD proteins of Enterococcus faecalis , Clostridium perfringens , Bacillus licheniformis , Lactobacillus sakei and Staphylococcus aureus . Several conserved regi ons were identified in S. rattus ArcA, including the signature arginine deiminase motifs SEIGKLKKVML (aa 11-21), FTRD (aa 164-167), EGGD (aa220-223), and MHLDTVF (aa 274-280) (Knodler et al. , 1998). S. rattus cOTC, encoded by arcB , shared 88, 86, 85 and 85% identity with the cOTCases of S. pyogenes, S. agalactiae, S. gordonii and S. pneumoniae, respectively. Additional homologies were obs erved with cOTC enzymes from E. faecalis , L. sakei , B.
47 licheniformis and S. aureus . Conserved carbamoylphosphate binding and catalysis motifs, STRTR and HPTQ, were identifie d at amino acid residues 57-61 and 135-138, respectively (Houghton et al. , 1984). The conserved ornith ine binding site (LHCLP) was identified at positions 270-274. Carbamate kinase, encoded by S. rattus arcC , was 73 and 72% identical to that of S. gordonii and S. pneumoniae , respectively. Homologies we re also observed with the carbamate kinases of Listeria monocytogenes , S. pyogenes , S. agalactiae , S. mutans , L. sakei , and B. licheniformis . Aside from the highly conserve d arginine residues at amino acid residues 157 and 160, no other cons erved motifs were identified in S. rattus ArcC, as was the case for S. gordonii and E. faecalis (Dong et al. , 2002; Marina et al. , 1998). The arginine/ornithine antiporter encoded by S. rattus arcD shared 49 and 42% identity with those of L. sakei and S. aureus , respectively. Conservation was also observed with the arginine/ornithine antiporters of S. agalactiae , C. perfringens and Pseudomonas putida . Twelve predicted transmembrane helices were identified in ArcD of S. rattus using the dense alignment surface method from DAS-Transmembrane Prediction Server (http://www.sbc.su.se/~miklos/DAS ) (Cserzo et al. , 1997). S. rattus arcT , encoding a putative peptidase, shared 58% identity with ArcT of L. lactis and 59% identity with SMU.816, a putative aminotransferase of S. mutans UA159. Additional homologies were obser ved with ArcT proteins of Lactobacillus lactis and L. sakei , as well as a Lactobacillus plantarum aminotransferase. The amino acid sequence of an appare nt regulatory protein linked to the arc operon was compared to those of regulatory proteins controlling arginine metabolism in other bacteria. The most significant level of similarity was observed with the putative S.
48 mutans UA159 ArgR (80% identity). The sec ond highest identity was with ArgR of S. agalactiae (58%), followed by putative arginine repressors in S. pyogenes and S. pneumoniae and AhrC of L. lactis . The AhrC protein was originally identified in Bacillus subtilis as a repressor of the arginine biosynthetic genes (Mountain and Baumberg, 1980). An AhrC homologue , ArgR, has been identified in B. licheniformis, where it both represses the anabolic orn ithine carbamoyltransferase and activates arc operon expression in the pres ence of arginine (Maghnouj et al. , 1998). In contrast to the B. licheniformis ArgR, it is not known whether the re gulatory protein associated with the arc operon in S. rattus also regulates argini ne biosynthetic genes. Additionally, only a very low level of similarity was shared between S. rattus AdiR and known Crp/Fnr-like proteins, such as S. gordonii ArcR (30%) and B. licheniformis ArcR (no similarity). To avoid generating confusion of this apparent S. rattus ADS regulatory protein with the global regulator ArcR or regul ators repressing argi nine biosynthesis (ArgR/AhrC), we have designated this protein AdiR to reflect its putative role as a regulator of the a rginine d ei minase gene cluster. We analyzed the predicted amino acid sequence of S. rattus AdiR to determine if any conserved DNA or arginine binding residues were present. The conserved SR--RE motif thought to be involved in DNA contact by the Escherichia coli ArgR (Tian and Maas, 1994) was found at amino acid residues 44-49 of the S. rattus AdiR, consistent with the theory that the N-terminus of ArgR is involved in DNA binding. Conserved amino acid residues thought to be impor tant for arginine binding in the E. coli ArgR (Burke et al. , 1994; Van Duyne et al. , 1996) were identified near the C-terminus at
49 positions 101 (alanine) and 124 (aspartic acid). In addition, a conserved glycine residue involved in oligomerization was id entified at position 122 (Van Duyne et al. , 1996). Most ADS genes appear to be hi ghly regulated (Barcelona-Andres et al. , 2002; Dong et al. , 2002; Maghnouj et al. , 1998; Zuniga et al. , 2002). Three transcriptional activators of the E. faecalis arc operon have been identi fied, including ArcR and two ArgR/AhrC-type regulators (Barcelona-Andres et al. , 2002). In E. faecalis , ArgR activates arc operon expression in the presence of arginine by bindi ng to a conserved ARG box located upstream of the arcA promoter (Maghnouj et al. , 1998). Transcriptional regulation of ADS by multiple protei ns has also been proposed for B. licheniformis , as putative binding sites for both ArgR/AhrC-t ype and Crp/Fnr-type proteins have been identified 5 to the transcriptional initiation site (Maghnouj et al. , 1998). ArcR, a Crp/Fnr-type transcriptional regulator, activates the arc genes by binding to a Crp-like consensus sequence upstream of the arcA promoter. We analyzed the sequence upstream of S. rattus arcA for possible ArgR or ArcR DNA bi nding sites. Potential binding sites for both ArgR and ArcR (Grandori et al. , 1995; Tian and Maas, 1994) were identified 200 and 52 bases upstream of the arcA transcriptional start site, respectively (Figure 32). The presence of a putative Arg box ( AATGAATTTATA AGTT AA ) and an imperfect palindrome possibly co nstituting a Crp binding site ( AATG C GA -N6TC TT ATT ) (bases in bold match the E. coli consensus sequences) (Maas, 1994) suggest that the ADS in S. rattus may be under the control of a Crp/Fnr family member, in addition to an ArgR/Ahr C-type protein (Maghnouj et al. , 1998); most likely the S. rattus AdiR protein.
50 Localization of parcA and Reporter Gene Fusions Primer extension analysis was used to map the arcA promoter region. A single band was observed corresponding to a G residue 49 bases upstream of the arcA start codon (Figure 3-3). Examination of the upstr eam sequence revealed a putative promoter that was most similar to 70-type promoters. The -10 region ( TA A AAT ) shared 5 out of 6 bases with the consensus sequen ce (bold), whereas the -35 region (A T TT CA ) identified 17 bases upstream of the -10 re gion shared only 3 bases with the consensus (bold). To determine which arc genes could be transcribed from the arcA promoter, RTPCR analysis was performed on RNA isolated from S. rattus FA-1 grown in TV + 2% galactose + 1% arginine (Fi gure 2-4). RT-PCR results suggested the presence of a polycistronic arcABCDT transcript. No transcript could be detected between the intergenic region of arcT and adiR , suggesting that adiR is transcribed from a separate promoter. Expression of the arc operon in Lactobacillus sakei (Zuniga et al. , 1998) , as well as some oral streptococci (Dong et al. , 2002), is under the contro l of carbohydrat e catabolite repression (CCR). ADS expression in these ba cteria appears to be up-regulated in the presence of arginine (Poolman et al. , 1987) and repressed by glucose (Curran et al. , 1998; Simon et al. , 1982). In AT-rich gram-positive bacteria, CCR is mediated by the trans acting catabolite control protei n A (CcpA), which binds to cis -acting catabolite response elements ( cre ) in the presence of preferred ca rbohydrate sources to regulate the expression of catabolic genes and operons (Saier et al. , 1996). Two potential CcpAdependent cre were identified at ( TGAAAT AA ATAACA ) and ( TGTAATCGCTTTC T) relative to the arcA transcriptional start site, with bases
51 matching the consensus shown in bold (Hueck et al. , 1994). The presence of these elements is consistent with the observation that the ADS in S. rattus FA-1 is regulated by CCR (Burne et al. , 1991). To assess the functionality of parcA and the putative cre sites found upstream of the promoter region, parcA and deletion derivatives were fused to a cat gene from S . aureus . Three arcA promoter fusions were constructed: (1) 340 bases upstream of the arcA transcriptional start site , including both putative cre sites, (2) 80 bases upstream, including one cre site, and (3) 50 bases upstream, lacking both cre sites. All of the promoter fusions were construc ted such that expression of cat was driven by the cognate arcA RBS. Since an efficient system of genetic transformation has not yet been established for S. rattus FA-1, the promoter fusions were integrated into the gtfG gene on the chromosome of the naturally competent, ADS-positive organism, S. gordonii DL-1 using a previously describe d integration vector (Chen et al. , 2002). Results showed that the intact parcA was functional in S. gordonii , and deletion of the putative cre sites resulted in increased promoter activity relative to the wild-type, providing evidence that expression from parcA is regulated via CcpA-dependent catabolite repression (Figure 3-5). Expression of AD in S. rattus The identification of putative cre sites upstream of parcA, as well as the ADS regulation patterns observed in S. gordonii , prompted us to inve stigate the role of catabolite repression in S. rattus FA-1 arc regulation. Wild-type S. rattus FA-1 was grown in TV broth containing glucose or ga lactose, with or without supplemental arginine, to mid-exponential phase and AD activity was measured. Galactose was included in the analysis because it is not repressive for AD expression in S. gordonii (Dong et al. , 2002). In wild-type S. rattus grown in 2% carbohydrate, peak AD activity
52 was found in cells grown in galactose and argi nine, while the expression level decreased by 75% in cells grown in glucose, regardless of the presence of argini ne (Figure 3-6). In agreement with previous studies (Burne et al. , 1991; Curran et al. , 1998), catabolite repression of ADS in S. rattus FA-1 was not evident when the amount of glucose or galactose present in the growth medium was reduced to 0.2%. This differs from CCR of the ADS in other closely relate d oral streptococci, including S. gordonii , where inclusion of as little as 0.2% glucose in the growth medium strongly repr esses ADS expression. The different susceptibilities of ADS e xpression to CCR may reflect different physiological roles of the system in these ba cteria. Specifically, susceptibility to CCR at lower carbohydrate concentrations would indicate that the ADS in S. gordonii may be primarily involved in energy generation and not acid tolerance, since the system may be repressed when conditions exist that favor low pH, (e.g., higher carbohydrate availability). In contrast, a lack of repression of the ADS in the presence of comparatively high levels of sugar may allow S. rattus to derive ATP from arginine and carbohydrate concurrently, and th e ammonia generated could e nhance the growth in, or protect the organisms from, a low pH environment. As shown in Figures 3-5 and 3-6, there is some induction of the S. rattus ADS by arginine, especially at low carbohydrate concentrations, but th e induction is not of the magnitude that is seen in other organisms. However, when S. rattus was grown to steady-state in a chemostat, AD enzyme activity is clearly induced in the presence of 50 mM arginine (Figure 3-7). In agreement with previous work, the S. rattus ADS does not appear to be up-regulated at pH 5 versus pH 7. Although the S. rattus AD is not acidinducible, previous studies have sh own that generation of ATP and NH3 from arginine at
53 low pH is vital for maintenance of pH and generation of en ergy when the surrounding medium is too acidic for efficient glycolysis. Taken together, these observations lend credence to the idea that there are fundamental differences in regulation of ADS in S. rattus when compared to other bacteria. Further physiologi cal characterization of the S. rattus arc operon is ongoing to understand the contribution of the ADS to acid tolerance of mutans streptococci. Summary Using an arcB gene fragment obtained by degenerate PCRs, the FA-1 arc operon was identified in subgenomic DNA librarie s and sequence analysis was performed. Results showed that the genes encoding the arginine deiminase pathway in S. rattus FA-1 are organized as an arcABCDT-adiR , including the enzyme s of the pathway, an arginine:ornithine antiporter (ArcD) and a putative regulatory protein (AdiR). The arcA transcriptional start site was identified by primer extension and a 70-like promoter was mapped 5 to arcA . Reverse transcriptase PCR was used to establish that arcABCDT could be co-transcribed. Re porter gene fusions and AD assays demonstrated that the operon is regulated by substrat e induction and catabolite repr ession, the latter apparently through a CcpA-dependent pathway.
54 Figure 3-1. The S. rattus FA-1 arc operon. Restriction sites used to isolate the arc gene fragment from a subgenomic library ar e noted. The arrows indicate positions of primers used in RT-PCR. Xba I Xba I Eco RV MW pI 46.6 4.80 37.9 5.20 33.7 5.06 51.0 8.92 43.3 4.82 16.5 6.16 1 2 3 4 5 6 7 kbp 0 arcA arcB arcC arcD arcT R Eco RV
55 Figure 3-2. Upstream sequence of S. rattus FA-1 arcA , with putative binding consensus sequences for ArgR (ARG), CcpA (CRE ) and ArcR (CRP) underlined. Bases matching the consensus sequences are shown in bold. The putative ARG box shown to be the binding site for AdiR (see Chapter 4) is highlighted. The and promoter elements are boxed. Th e transcriptional in itiation site (TIS) and ribosomal binding site (RBS) are underlined. AGCTGCCTTAGTTTTAATAGTCTA AATGAATTTATA AGTT AA ATATT ATAAAAATTTTAATTAATATTTATAAAATATACAGGTTAAAGTTAGG TTTTATTCGA TGTAATCGCTTTC T TTTTTAGTTTTTATAAATTCTTTC ATCAAAAAACG TGAAAT AA ATAACA ATTCA AATG C GAAAAAAATC TT ATT TCATTTATAAAAAACCTCTAAAATGGAATTTG TAATAAATGA GAAATTAATGATTCTCAAAATCGAAAGGAGT AGTTAACATG CRE CRE -35 -10TIS RBSarcA >> ARG CRP
56 Figure 3-3. Primer extension analysis of S. rattus arcA . The arrowhead indicates initiation of transcription at a G residue, 49 bases 5 to the ATG translational start codon. T G C A
57 Figure 3-4. Reverse transcriptas e PCR analysis of mRNA from S. rattus grown in TV broth containing 2% galactose and 1% arginine. Primers specific to the arc intergenic regions were used to amp lify cDNA. Lane 1, molecular weight markers; lanes 2 4, arcAB intergenic region; lanes 5 7, arcDT intergenic region; lanes 8 10, arcTR intergenic region. The order within each triplicate (2-4, 5-7, 8-10) is: cDNA, chromosomal DNA, and a control in which a reaction containing mRNA with no reverse transcriptase was used in the amplification reaction. arcAB arcDT arcTR Lanes: 2,5,8 cDNA 3,6,9 chDNA 4,7,10 No RT arcA B C D T R 1 2 3 4 5 6 7 8 9 10
58 Figure 3-5. CAT specific activity of the S. rattus FA-1 arcA promoter (parcA 340) and derivatives in which one (parcA ) or both ( parcA 50) cre have been deleted. Bacterial cultures used in the CAT assa ys were grown in TV containing 0.2% or 2% carbohydrates, with or without 1% arginine. CA T activity is expressed as nmol/min/mg protein. Results shown are the average and standard deviations (error bars) of a minimum of 9 separate cultures for each strain and condition. One-way ANOVAs and pair-wise Student t-tests were used to identify significant differences (p < 0.05) in CAT activity. Similar letters indicate no significant di fference in CAT activity. a a a a Glucose Galactose 0 5 10 15 20 parcA 340 parcA 80 parcA 50 parcA 340 0.2% 2% 2% 2% Glucose + Arginine Galactose + Arginine e d b e a a b c e e b fCAT Specific Activity U (mg protein)-1
59 Figure 3-6. Arginine deiminase enzyme activity of S. rattus FA-1 grown in TV broth containing 0.2% or 2% ca rbohydrates, with or without 1% arginine. Activity is expressed as mol citrulline produced/min/mg protein. Results shown are the average and standard deviations (e rror bars) of a minimum of 9 separate cultures for each strain and conditi on. One-way ANOVAs and pair-wise Student t-tests were used to identify significant differences (p < 0.05) in AD activity. Similar letters indicate no significant difference in AD activity. 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 0.2 % 2 % Glucose Galactose Glucose + Arginine Galactose + Arginine 0.0 a b d e c ac f f AD Activity U (mg protein)-1
60 Figure 3-7. Arginine deiminase enzyme activity of S. rattus FA-1 chemostat cultures grown in TV medium containing 25 mM glucose, with or without 50 mM arginine. Steady state cultur es were maintained at pH 5 or pH 7 with the addition of 2 M KOH. Activity is expressed as mol citrulline produced/min/mg protein. Results show n are the average and standard deviations (error bars) of a minimum of 9 separate cultures for each strain and condition. One-way ANOVAs and pair-wise Student t-tests were used to identify significant differences (p < 0.05) in AD activity. Similar letters indicate no significant di fference in AD activity. 0 0.2 0.4 0.6 0.8 1 1.2 1.4 1.6 50 mM Arg: + + pH: 5 5 7 7 a b b c AD Activity U (mg protein)-1
61 CHAPTER 4 REGULATION OF ARGININE DEIMINASE EXPRESSION IN S. rattus FA-1 Introduction As described in the previous chapter, a putative transcriptional regulator, AdiR, is transcribed immediately downstream of the arcABCDT operon in S. rattus . In this chapter, the role of AdiR in arc gene regulation is investig ated. However, as detailed below, it was not feasible to construct a mutation of the adiR gene in S . rattus , as attempts to genetically manipulate this orga nism did not yield consistently successful results. Given these circumstances , the closely related bacterium, S. mutans UA159, was used as a host to study regulation of the S. rattus arc operon. Taxonomic studies have suggested that S. rattus diverged from S. mutans during evolution of the mutans group, but that the two bacteria remain very closely related (Kawamura et al. , 1995). In fact, S. rattus was considered a subspecies of S. mutans until 1977, when Coykendall proposed to elevate S. rattus to species status based on the observation that S. rattus produced citrulline from arginine, while S. mutans did not (Coykendall et al. , 1976) (Coykendall, 1974). Subsequent ly, the arginine dihydrolase (or "deiminase") test became the main biochemical criterion used to distinguish the two organisms. Thus, a definitive phenotypic difference resulting from the evolutionary divergence is that S. rattus retained a functional ADS, while S. mutans lost, or failed to acquire, the system. Although S. mutans does not possess the ADS, a clus ter of ADS-like regulators has been identified in the UA159 genomic sequen ce. The presence of these regulators has
62 several potential implications. Notably, if these regulators are capable of controlling ADS expression in a manner consistent with ADS regulation in other bacteria, an arginolytic strain of S. mutans could potentially be constructed and implanted in an animal model to determine if this strain can compete with other oral bacteria in a manner similar to wild-type S. mutans , while reducing the cariogenic ity of dental plaque via alkali production. Thus, the goals of this study were three-fo ld: (1) to locate the binding site of AdiR within parcA, (2) to determine if the putative ADS regulators in S. mutans have retained the ability to regulate the arc genes, and (3) to use S. mutans as a model to examine the roles of the S. rattus ADS regulators, specifically fo cusing on AdiR. Ultimately, the results of this study should shed li ght on the evolutionary divergence of S. rattus and S. mutans in relation to the ADS. Results Identification of the AdiR Binding Site Within parcA AdiR shares significant homology with th e global regulator, ArgR. In other bacteria ArgR (also referred to as AhrC) functions as a classic feedback regulator, repressing the arginine biosynt hetic genes in the presence of arginine (Glansdorff, 1987; Lim et al. , 1987; North et al. , 1989; Smith et al. , 1989). At the same time, ArgR activates expression of genes fo r arginine catabolism operons, such as arginine deiminase ( arc ), arginine succinyltransferase ( ast ) and arginase ( roc ) genes (Gardan et al. , 1997; Kiupakis and Reitzer, 2002; Maghnouj et al. , 1998; Park et al. , 1997). In E. coli , ArgR functions as a 98-kDa hexamer composed of identical 16.5-kDa monomers. The hexamer is stabilized when six argi nines bind at the trimer:trime r interface via ArgR residues alanine-103, aspartic acid-125, a nd aspartic acid-126 (Van Duyne et al. , 1996). An N-
63 terminal winged helix-turn-helix (HTH) do main mediates DNA binding via the serine-47 and arginine-48 residues, introdu cing a 70 bend in the DNA (Burke et al. , 1994; Grandori et al. , 1995; Tian and Maas, 1994). The presence of conserved DNA and arginine binding motifs in S. rattus AdiR (Figure 4-1) suggest this regulator may activate the arc operon in trans by binding to a conserved ArgR binding site within parcA. In other bacteria, ArgR activates the ADS in an arginine-dependent manner by binding to a conserved ar ginine binding site (ARG box) upstream of the arc operon and inducing transcription (Klingel et al. , 1995; Maas, 1994; Miller et al. , 1997; RodriguezGarcia et al. , 1997). The ARG box typically forms a weak palindrome with the consensus sequence TGMAT-wwww-ATKCA (M = A or C; W = A or T; K = G or T; less conserved bases shown in lower cas e), although the exac t sequence varies considerably within and am ong bacterial species (Makarova et al. , 2001). ARG boxes are typically found in pairs, arranged in tandem within the promoter regions of ArgRcontrolled operons. However, some exceptions have been noted in the literature. For example, a single ARG box is present in the promoter region of the Bacillus subtilis rocDEF operon, and no conserved ARG boxes are present in the Lactococcus lactis subsp. cremoris arcA promoter, despite tight control of the arc genes by ArgR/AhrC-type regulators (Larsen et al. , 2004; Miller et al. , 1997). In addition, the arcA promoter region of L. lactis subsp. lactis contains multiple ARG box half-sites that directly participate in DNA binding by ArgR (Larsen et al. , 2005). Multiple ARG box half-sites were also identified in the promoter regi ons of the arginine biosynthetic car and argF operons in Pseudomonas aeruginosa (Park et al. , 1997).
64 Sequence analysis of S. rattus parcA revealed potential ARG boxes at positions -200 (TGAAT TTAT A AG TT) and -57 (TGAA A-TAAA TAACA ) relative to the arcA transcription initiation site (underlined bases match the consensus). Of note, a variation of the -57 ARG box is present when the putative core motifs are separated by 9-bp instead of 4-bp (TGAA A-TAAA TAACA-ATTCA ), a distance that corresponds to almost one full turn of the DNA helix, potentially allowing the ArgR trimers to bind the same DNA â€œface,â€ as is the case with ARG boxes separated by 2-3-bp. To locate the AdiR bi nding site within parcA, the adiR structural gene was cloned onto the pET45b expression vector and AdiR was purified by Ni-NTA chromatography. Gel shift assays were performed to monitor AdiR binding to 32 P[ -ATP]-labeled probes corresponding to overlapping 30-mers spanning parcA (Figure 4-2) . Results showed that AdiR binds within the first 82 bases of parcA in an arginine-indep endent manner. This region contains the predicted 57 ARG consensus sequences (TGAA A-TAAA TAACA ; and the extended version TGAA A-TAAA TAACA-ATTCA ). A conserved CRP binding site (AAA CGTGAAATAAAT AACAATT ) (Maghnouj et al. , 2000) overlaps the ARG consensus sequences in this region, spanning ba ses -53 to -74, raising the possibility that the S. rattus arc operon is controlled by multiple transcriptional regulators. Of note, the S. gordonii arc operon is under the contro l of three regula tors: Flp, ArcR and CcpA. In S. gordonii , the ArcR binding site overlaps with one of the conserved CcpA binding sites by 12 bases, suggesting that modulation of arc expression by glucose and arginine may involve some physical interac tion of ArcR and CcpA (Zeng et al. , 2006). In addition, the ARG boxes upstream of arc operons in E. faecalis and B. licheniformis overlap with conserved CRP binding sites (Barcelona-Andres et al. , 2002; Maghnouj et al. , 1998),
65 furthering the possibility that Crpand ArgR-type regulator s coordinately regulate the arc genes. Identification of Additional Regula tory Genes Linked to the ADS in S. rattus FA-1 As discussed in Chapter 3, the arc operon was isolated from subgenomic DNA libraries on two fragments of 7-kbp and 4-kbp. The fragments were sequenced and found to encode arcABCDT and adiR . After characterizing the arc genes, subsequent analysis of the isolated fragments revealed two additio nal regulatory genes with possible ties to the ADS. Flp. On the 5 end of the available sequence, an ORF with significant homology to the F nr-l ike p rotein, Flp, was identified (Dong et al. , 2004; Lu et al. , 1999; Stockley et al. , 1998). Flp belongs to the Crp-Fnr superfamily of transc riptional regulators that modulate arc expression in response to oxygen tension in other bacteria (Dong et al. , 2004; Lu et al. , 1999; Stockley et al. , 1998). The paradigm of this superfamily, E. coli FNR (for â€œF umarate N itrate R eductaseâ€), possesses a [4Fe -4S] cluster that undergoes oxidation during growth in aerobic cond itions, rendering the pr otein inactive. Conversely, anaerobic growth l eads to reduction of the four conserved cysteine residues that serve as ligands for the [4Fe-4S] cluste r, allowing FNR to stim ulate genes involved in anaerobic respiration and repr ess genes involved in aerobesis, over 75 lo ci in all (Kiley and Helmut, 1999). Crp-Fnr regulators are typically com posed of 230 â€“ 250 amino acids and are characterized by the presence of both an N-te rminal sensory domain and a C-terminal HTH DNA binding domain (Korner et al. , 2003; Unden and Duchene, 1987). The sensory domain consists of an -helix involved in dimerization and an antiparallel -
66 barrel held in place by three c onserved glycine residues (Shaw et al. , 1983; Unden and Duchene, 1987; Weber and Steitz, 1987). Un like Crp, which binds cyclic-AMP in Gramnegative bacteria (Unden and Duchene, 1987; Weber and Steitz, 1987), Fnr is a metalloprotein that regulates genes in response to oxygen tension (Lazazzera et al. , 1996). The N-terminal 30 amino acids of Fn r contains a cluster of four conserved cysteine residues that act as ligands for a [4Fe4S] cluster. The C-terminal HTH motif of Crp-Fnr-type regulators contains conserved arginine180, glutamine-181, and arginine-185 resi dues involved in DNA contact (Lazazzera et al. , 1996). Previous studies sugge st that Crp-Fnr pr oteins bind as dimers to conserved Fnr sites (TTGAT-N4-ATCAA) and Crp s ites (aaaTGTGAtctagaTCACAttt) within the promoter regions of target genes. Differe nces in the consensus sequences recognized by the E. coli Fnr and Crp proteins are due to th ree conserved amino acid residues at positions 213, 217 and 221 in the HTH domain. Replacement of the Fnr residues (V-213, S-217 and G-221) with Crp residues (R-213, G-217 and K-221), conveys the ability to recognize and bind to conser ved Crp sites (Lazazzera et al. , 1996). The Flp protein of L. casei binds to a similar consensus sequence (CCTGA-N4-TCAGG) to modulate expression of genes involved in the redox response (Gostick et al. , 1998). S. rattus flp is encoded 1064-bp upstream of arcA . The predicted amino acid sequence of Flp was determined and found to c ontain an N-terminal sensory domain with three conserved glycine residues at positions 47, 59 and 70, as well as a C-terminal HTH motif, with conserved RE residues at positions 190 and 191 (Figure 4-3). Three cysteine residues were identified in Flp, in contrast to the four residues conserved in other Fnr homologs. Of note, only two cystei ne residues are present in the S. gordonii , L. lactis
67 and L. casei Flp proteins (Dong et al. , 2004; Gostick et al. , 1998; Scott et al. , 2000). S. rattus Flp shares highest homology with a putative Crp-Fnr type regulator in S. mutans UA159 (SMU.2094c; 88% ID), and lesser hom ology with a conserved hypothetical protein in Streptococcus pyogenes (1276; 54% ID), Fnr of B. licheniformis (41% ID), a putative Flp of E. faecalis (36% ID), ArcR of L. sakei (32% ID), and Flp of S. gordonii DL-1 (29% ID). Previous studies have shown that AD enzyme activity in S. rattus is slightly repressed during aerobic growt h, implying that this system is under the control of an oxygen-responsive regulator (Burne et al. , 1991). Furthermore, sequence analysis of parcA revealed an imperfect palindrome possibly constituting a Crp binding site (AATG CGA -N6-TC TTATT ) (underlined bases match the E. coli consensus sequence) (Maas, 1994) at positi on relative to the arcA transcriptional start site (Figure 3-2). Thus, it is possible that the S. rattus arc operon is activated by Flp during anaerobic growth, in addition to ac tivation by AdiR in the presence of arginine. AdiR2. On the 3 end of the available se quence, immediately following adiR , an ORF of unknown function was identified that shares homology with conserved hypothetical proteins in other bacteria. This ORF has been designated adiR-2 to reflect the tight linkage of this gene to the arc operon and adiR-1 in S. rattus . Conserved Crp or Fnr-type residues were not detected in AdiR2 using the NCBI Conserved Domain Database. Genetic Manipulation of S. rattus FA-1 To further explore the role of AdiR in ADS regulation, we attempted to construct a knockout of this gene in S. rattus . However, genetic manipulation of S. rattus is very difficult (Havarstein et al. , 1997; Perry and Kuramitsu, 1981). In other streptococci, such
68 as Streptococcus pneumoniae , Streptococcus mitis and Streptococcus oralis , secretion of the competence-stimulating peptide, ComC, activates a quorum sensing system composed of a histidine kinase, ComD, a nd a response regulator, ComE, that mediate induction of genetic competence (Havarstein et al. , 1997). Several strategies were used to introduce foreign DNA into S. rattus . To optimize the conditions required for tran sfer of foreign DNA into S. rattus , the streptococcal shuttle vector pDL278 was purified from S. gordonii DL-1 and used in the following experiments to decrease potential restric tion-modification problems. Initially, we attempted to induce natural competence in S. rattus by optimizing the protocol used for natural transformation of S. mutans . Bacteria were grown in BHI containing 10% horse serum and incubated with competence-stim ulating peptide (ComC) purified from S. mutans UA159. At various optical densities at 600 nm (OD600) ranging from 0.1 to 0.4., 200-700 ng pDL278 was added to the cultures, as indicated in the protocol for transformation of S. mutans UA159 (Perry and Kuramitsu, 1981). No spectinomycinresistant colonies formed on the BHI pl ates, even after extended incubation. Other groups have introduced DNA into re lated non-naturally competent bacteria, such as S. salivarilus, by growing the bacteria in me dia containing cell wall weakening agents, such as glycine or DL-threo nine, prior to elect roporation (Buckley et al. , 1999; Chassy, 1976). Glycine replaces D-alanine residues in the peptidoglycan layer of Grampositive bacteria, interfering with cell wall synthesis and assembly. In a similar manner, DL-threonine inhibits cell wall crosslinking and increases the efficiency of electroporation. The minimum inhibitory c oncentrations (MICs) of glycine and DL-
69 threonine were found to be 0.05 M and 0.1 M, respectively, for S. rattus grown in Todd Hewitt media containing 0.2% yeast extract (THY). The protocol used to electroporate S. salivarius was obtained and used as a starting point for manipulating S. rattus (Buckley et al. , 1999). In preparat ion for electroporation, S. rattus was grown in THY media containing either 0.05 M glycine or 0.1 M DLthreonine to various optical densities at 600 nm (OD600) ranging from 0.1 to 0.4. Bacteria were harvested and electroporated w ith 200-600 ng pDL276. As a control, S. rattus was electroporated without DNA. Cells we re plated on BHI agar containing 500 g mL-1 of spectinomycin and incubated for 48 hours at 37 C in 5% CO2. After 24 hours, fewer than five pinpoint colonies appeared on agar plates corresponding to S. rattus grown in 0.05 M of glycine to OD600 = 0.2 and electroporated with 200-500 ng of DNA. Restriction enzyme digestion of plasmid DNA isolated from these transformants was consistent with the pattern expected for pDL278. While the shuttle vector, pDL278, was successfully introduced into S. rattus via electroporation, albeit not very efficientl y, double or single cross mutations were not obtained when the adiR and arcB genes were cloned onto suicide vectors, pGEM7 and pJZ26, respectively, and electroporated into S. rattus . An additional strain, S. rattus BHT, was also evaluated but was unable to integrate foreign DNA, in agreement with past studies by other groups (Perry and Kuramitsu, 1981). Several factors might explain why S. rattus is refractile to genetic manipulation. One explanation is that S. rattus does not possess the comCDE genes. Attempts by myself and others to amplify the S. rattus comCDE genes were not successful (Havarstein et al. , 1996), although these experiments were performed using degenerate primers, or
70 primers based on S. mutans comCDE , because the genomic sequence of S. rattus is not available. However, given the close taxonomic relationship between S. rattus and S. mutans , it would be highly surprising if S. rattus did not possess the comCDE genes. It is more likely that genes invol ved in competence exist in S. rattus , but that the conditions required for their optimal expression were not being met in the above experiments. Given the difficulty constr ucting mutant strains of S. rattus , we explored the use of S. mutans as model for studying S. rattus arc regulation. Identification of an ADS-Like Gene Cluster in S. mutans UA159 Although the S. mutans chromosome does not contain the arcABCDT operon, analysis of the genomic seque nce revealed that this bact erium has retained the three regulatory genes that flank this operon in S. rattus (Figure 4-4). These three ORFs are grouped together on the UA159 chromosome and share substantial id entity with their homologs in S. rattus . Reverse transcriptase PCR, using primers specific to flp , adiR1 and adiR2 of S. mutans , was used to verify that the ge nes are transcribe d (Figure 4-5). The %GC content of the intergenic region between flp and adiR1 (26.23%) differs substantially from the average CG content of S. mutans (36.82%), suggesting that the arc operon may have been acquired from a fore ign source during evolution of the mutans streptococci. Furthermore, two i nverted repeats were found in the flp-adiR intergenic region, one of which (GTTCATTTAG-N34-CTAAATGAAC) is located approximately 350 bases downstream of flp and shares substantial homol ogy to an IR found 200 bases downstream of flp in S. rattus FA-1 (GTTCATTAGGCACT-N3AGTGCCTAATGAAC). Together, these obs ervations raise th e possibility that S. mutans either lost or failed to aquire genes encoding the arc enzymes at this locus during the process of diverging from S. rattus .
71 Flp. The amino acid sequence of Flp of S. mutans was used in a BLAST search against other available micr obial genomes using NCBI blastp and homologs were identified in several other bacteria (Figure 4-6). Interestingly, the majority of these homologs were encoded by flp genes linked to arc operons. Highest homology was observed with S. rattus flp (88% ID), and lesser ho mology was observed with a conserved Crp-type regulator in Streptococcus pyogenes (1276; 54% ID), Fnr of B. licheniformis (41% ID), a Crp-like regulator in E. faecalis (36% ID), ArcR of L. sakei (32% ID), and Flp of S. gordonii DL-1 (29% ID). Sequence analysis of S. mutans Flp revealed three conserved glycine residues at positions 47, 59 and 70. However, this protein terminates prematurely and does not contain the conserved HTH domain or RE motif, suggesting that perhaps th is protein performs an alte rnate function re lated to redox sensing in S. mutans . Alternatively, Flp may be capab le of modulating gene expression by interacting with an additional DNA-bindi ng regulatory protein, such as AdiR. AdiR1. In contrast to Flp of S. mutans , which shares highest similarity with other arc -associated regulatory protei ns, homologs of UA159 AdiR1 and AdiR2 are typically linked to DNA modification proteins in other bacteria (Fi gure 4-6). AdiR1 shares highest homology with S. rattus AdiR1 (80% ID) and S. pyogenes ArgR2 (57% ID). Interestingly, the regions immediatel y downstream of these homologs in S. rattus and S. pyogenes contain homologs of AdiR2 (50% a nd 43% identity, respectively). In S. rattus , the AdiR1 and AdiR2 proteins are associated with the arc operon. However, in S. pyogenes , AdiR12 are linked to the DNA mismatch repair protein, MutS, and the arginyltRNA synthetase, ArgS, as is the case in S. mutans (Figure 4-6). Other homologs of UA159 AdiR1 were also associated with DNA modification protei ns: UA159 AdiR1 is
72 45% and 33% identical to AhrC in L. lactis lactis and B. licheniformis , respectively, which are located immediately upstream of the DNA repair protein RecN. The conserved SR--RE motif thought to be involved in DNA contact by the Escherichia coli ArgR (Tian and Maas, 1994) was f ound at amino acid residues 44-49 of the S. mutans AdiR1, consistent with the theory that the N-terminus of ArgR is involved in DNA binding (Figure 4-1). Conserved ami no acid residues thought to be important for arginine binding in the E. coli ArgR (Burke et al. , 1994; Van Duyne et al. , 1996) were identified near the C-terminus at positions 101 (alanine) and 124 (aspartic acid). In addition, a conserved glycine residue involve d in oligomerization was identified at position 122 (Van Duyne et al. , 1996). AdiR2. When the predicted amino acid seque nce of AdiR2 was blasted against other bacterial genomes using NCBI blastp, the majority of the hits were conserved hypothetical proteins associated with MutS (Figure 4-6). No conserved domains were detected in UA159 AdiR2, or in any of its homologs, so the function of this conserved hypothetical protein is not clear. Howe ver, the association of AdiR12 with DNA modification proteins is not entirely unprecedented. In E. coli , ArgR functions as an accessory protein required for resolution of ColE1 multimers to monomers during sitespecific recombination at the cer locus (Stirling et al. , 1988). Therefore, it is possible that the S. mutans adiR12 genes cooperate with mutS to accomplish a task related to DNA repair. Mutation of mutS results in an increased rate of spontaneous mutations due to ineffective mismatch repair (Schaaper and Dunn, 1987). To determine if the same was true for mutation of adiR12 , S. mutans wild-type and adiR12 mutant strains were independently grown to OD600 = 0.3, serially diluted and pl ated on BHI agar containing
73 500 g mL-1 of streptomycin. No difference was observed in the rate of spontaneous mutations in adiR versus UA159 wild-type. Further e xperiments are necessary to fully characterize the link between AdiR and DNA repair. Regulation of S. rattus parcA by S. mutans AdiR12 and Flp As we were not able to generate knockouts in S. rattus , we explored whether regulated expression from S. rattus parcA could occur in S. mutans , with the goal of using S. mutans to further an understanding of S. rattus ADS regulation. Subsequently, the S. rattus parcA and deletion derivatives were fuse d to a promoterless chloramphenicol acetyltransferase ( cat ) gene and integrated into the wild-type S. mutans chromosome. Consistent with the data obtained from gel shift analyses, efficient expression from parcA was observed when the 80-bp region 5 to the TIS was fused to cat , but not when a shorter fragment consisting of 53-bp 5 to the TIS was used. These results suggest that the binding site of a transc riptional activator, possibly S. mutans AdiR, is located between -53 and -80 of the TIS, most likely at the putative Crp site (-53 to -74). To further examine the roles of AdiR and Flp in ADS regulation, the parcA reporter fusion was introduced into seve ral engineered strains of S. mutans (Figure 4-7). As discussed in Materials and Methods , these strains harbored: (1) an adiR12 mutation ( adiR ::Spec), (2) a flp mutation ( flp ), (3) an adiR12 mutation complemented by the S. rattus adiR12 ( adiR12 ::F adiR1 2Spec) and (4) mutations in both adiR12 and flp ( flpadiR12 ). A fifth strain was constructed in an attempt to complement the double flpadiR mutation with the corresponding genes in S. rattus . However, subsequent comparison of the flp promoter regions in S. rattus, S. mutans and other bacteria reve aled an error in the
74 S. rattus sequence. Because of this e rror, the predicted start codon for S. rattus Flp was located 40-bp downstream of the actual star t codon. After re-se quencing the region, it was determined that the ATG start codon for S. rattus flp was located at nucleotides 2-4 of the available arc operon sequence. Because of this error, the promoter region of S. rattus flp was not included in the fifth engineered strain. Therefore, the fifth strain was actually comprised of a double flpadiR mutation, with only the adiR mutation complemented by S. rattus adiR ( flpadiR12 ::F adiR1 2). This sequencing error revealed a further complication: the S. rattus flp promoter sequence was no longer available, because flp started at the se cond nucleotide on the available sequence. Theref ore, to study the role of S. rattus Flp, the flp structural gene was cloned into the streptococcal protein expr ession vector pNisin, generating pNicinFlp. This vector permitted nicin-inducible expression of S. rattus Flp to complement the flp mutation in S. mutans , forming a sixth strain ( flp ::F flp ). As a control, the empty pNisin vector was introduced into the flp strain, creating a seventh strain ( flp ::pNisin). When parcA reporter activity was measured in th e wild-type and e ngineered strains of S. mutans , slight but significant differences we re observed (Figur es 4-8 and 4-9). CAT specific activity was reduced by 1.3-fold in the adiR12 strain in relation to wild-type S. mutans , consistent with the proposed role of AdiR as a transcriptiona l activator. Addition of arginine to the growth medium did not have a significant effect on parcA expression in the wild-type or adiR12 strains. However, when the adiR12 mutation was complemented with S. rattus adiR12 , CAT activity was 1.6-fold higher than the adiR12 strain and was induced by 1.3-fold in the presence of arginine. Interestingly, CAT activity was 1.3-fold higher in the comp lemented strain than in wild-type S. mutans ,
75 suggesting that the S. rattus AdiR activates the ADS more efficiently than does the S. mutans AdiR. These results suggest that the S. mutans AdiR is capable of regulating the S. rattus arc genes to a limited extent, but that this protein has not retained the ability to respond to arginine in the environment. The S. rattus AdiR appears to function as a transcriptional activat or of the ADS, and is capable of inducing arc gene expression in response to arginine. More over, the fold-induction of parcA observed in the presence of arginine correlates with the fold-induc tion observed in AD assays performed in S. rattus FA-1 (Figures 3-6 and 3-7), further supporti ng a role for AdiR in activation of the arc genes in response to arginine. Several factors may account for the modest decrease in CAT activity in the absence of AdiR. The most likel y explanation is that the arc genes are activated by other regulators in addition to AdiR, as is th e case with many other ADS-positive bacteria (Barcelona-Andres et al. , 2002; Dong et al. , 2004; Larsen et al. , 2004; Lu et al. , 1999; Maghnouj et al. , 1998; Wohlkonig et al. , 2004). In the case of E. faecalis , two ArgRtype regulators are required for optimal arc expression (Barcelona-Andres et al. , 2002). If the same is true for S. rattus , then mutation of one tran scriptional activator would presumably result in only slight repression of parcA, as observed in this experiment. Consistent with previous studies, aer obic growth reduced expression from parcA, suggesting that the binding site for an oxygen -responsive regulator is present within the first 80 bases of parcA, possibly at the conserved CRP site (Figure 4-9). However, inactivation of S. mutans flp did not alter arc expression during growth in aerobic (shaking at 300 rpm) or anaerobic (Gas Pak) environments, as compared to arc expression in wild-type S. mutans . Therefore, it appears that S. mutans flp most likely serves a
76 function unrelated to the ADS. Interestingly, when the flp mutation was complemented with Flp from S. rattus, CAT activity was slightly redu ced during aerobic growth, but was not altered during anaerobic growth (Fi gure 4-9). These results indicate that S. rattus Flp may be capable of repressing the arc promoter during aerobic growth, but does not appear to activate the system during anaerobic growth. This finding contradicts the role of Flp in other bacteria as an activator of the arc genes during anaerobic growth. Although the decrease in parcA expression was statistically significant, further work is needed to determine if the relatively minor decrease in CAT activity truly reflects a role for Flp in regulation of the S. rattus ADS. Summary In conclusion, an AdiR binding site was identified within S. rattus parcA, and we have shown that S. mutans is a suitable model organism for studying arc regulation. Results from this study suggest that the S. mutans AdiR has retained the capacity to activate arc gene expression, albeit not e fficiently or in response to arginine. In contrast, S. mutans Flp does not appear to have retained ADS regulatory capacity. It is likely therefore that S. mutans flp and adiR12 represent orthologous ge nes that have been retained through evolut ion to modulate other cellular functions. As predicted, the S. rattus AdiR and Flp proteins can function as tr anscriptional regulators of the ADS in a heterologous host, cap able of activating arc gene expression in re sponse to arginine or repressing arc expression in response to aerobic growth, respectively.
77 Figure 4-1. Alignment of the S. rattus and S. mutans AdiR1 proteins with homologs in other bacteria, using the MacVector program and ClustalW alignment software.
78 Figure 4-2. Gel shift analysis of purified His6x-AdiR from S. rattus and 32P[ -ATP]labeled probes corresponding to regions within S. rattus parcA. 1 2 3 4 5 6 7 8 9 1, 4, 7: AdiR + Probe 2, 5, 8: AdiR + Probe + Arginine 3, 6, 9: No protein VII* VI* V AdiR + parcA Unbound Probe 35 -1 0 +1 VII* VI*V 53 80 Conserved ARG Box
79 Figure 4-3. Alignment of the S. rattus and S. mutans Flp proteins with Fnr-Crp proteins in other bacteria, using the MacVect or program and ClustalW alignment software.
80 Figure 4-4. Identification of putative ADS regulators in S. mutans UA159 88% ID 80% 50% flp arcA arcB arcC arcD arcT adiR-1 adiR-2 flp S. rattus arc operon S. mutans chromosome 426-bp adiR-1 adiR-2 Site of Evolutionary Divergence?
81 Figure 4-5. Reverse transcriptas e PCR of mRNA isolated from S. mutans UA159. Total mRNA was amplified with random he xamers. Resulting cDNA was amplified using primers specific to adiR1 , adiR2 or flp . adiR1 adiR2 f lp MW 1 2 3 4 5 6 + + + -
82 Figure 4-6. A) Homologies of S. mutans Flp, AdiR1 and AdiR2 to proteins in other bacteria, determined using NCBI Blas tp. B) Organization of the ADS-like cluster on the chromosome of S. mutans UA159. S. mutans UA159 A B
83 Figure 4-7. Integration of the S. rattus parcA cat fusion into wild-type S. mutans and six engineered strains.
84 Figure 4-8. CAT specific activity of S. rattus parcA in S. mutans wild-type and four engineered strains. Cells we re grown at 37 in 5% CO2, in TV medium containing 25 mM galactose, with (+) or without (-) 50 mM arginine. Activity is expressed as nmol/min/mg protein. Results shown are the average and standard deviations (error bars) of a mi nimum of 9 separate cultures for each strain and condition. One-way ANOVAs a nd pair-wise Student t-tests were used to identify significant differenc es (p < 0.05) in CAT activity. Similar letters indicate no signifi cant difference in activity. 50 mM Arg: + + + + + 0 10 20 30 40 50 60 70 80 90 100 UA159 adiR flpadiR adiR flpadiR ::F adiR ::F adiR a ab ac c bd b ab e b f CAT Activity U (mg protein)-1
85 Figure 4-9. CAT specific activity of S. rattus parcA in S. mutans wild-type and three engineered strains. Cells were grown at 37 in an aerated (+; shaking at 300 rpm) or non-aerated (-; anaerobic GasP ak) environment, in TV medium containing 25 mM galactose and 50 mM arginine. Activity is expressed as nmol/min/mg protein. Results shown are the average and standard deviations (error bars) of a minimum of 9 separate cultures for each strain and condition. One-way ANOVAs and pair-wise Student t-tests were used to identify significant differences (p < 0.05) in CA T activity. Similar letters indicate no significant difference in activity. UA159 flp flp :: flp :: pMSP3535 F flp 0 10 20 30 40 50 60 70 80 90 Aeration: + + + + b a b a b a a c CAT Activity U (mg protein)-1
86 CHAPTER 5 ANALYSIS OF AN AGMATINE DEIMINASE GENE CLUSTER IN S. mutans UA159 Introduction Streptococcus mutans is the etiological agent of dental caries and the cariogenicity of this organism is directly rela ted to its ability to survive in a self-induced acidic environment (Kuramitsu, 1993). When dental plaque reaches a pH of around 5.5, significant tooth demineralization can begin and dental caries may result. A primary determinant of acid tolerance by S. mutans is the membrane-bound F1F0-ATPase (Bender et al. , 1986), although other factors contributing to acidurity in clude a reduction in the proton permeability of the cell membrane a nd induction of DNA repair pathways and stress proteins (Quivey et al. , 2001), as described in the Introduction. Although S. mutans uses several complex acid-toler ance responses, it is considered incapable of ammonia generation and is know n to have a strong competitive advantage over acid-sensitive bacteria at low pH. Consequently, it was of interest when a cluster of genes was identified in the S. mutans UA159 genome with similarity to known ADS genes (Ajdic et al. , 2002). The gene cluster was originally annotated otcA , SMU.263, SMU.264 and arcC (Figure 1). In other bacteria, otcA and arcC encode an anabolic ornithine carbamoyltransferase and a catabolic carbamate kinase, respectively, and are involved in catabolism of arginine via the ADS . Because absence of a functional ADS is the major distinguishing factor between S. mutans and its closest relative, S. rattus (Coykendall, 1974), alternate functions of th e gene cluster were suggested (Ajdic et al. , 2002). The most promising theory stated that the genes encoded enzymes of the
87 agmatine deiminase system (AgDS), a pathway highly analogous to the ADS, but involved in catabolism of the primary amine, agmatine (Figure 2) (Simon and Stalon, 1982). Agmatine catabolism via the AgDS. Agmatine is a decarboxylated derivative of arginine that can be catabolized by the AgDS (Simon and Stalon, 1982). The AgDS has been identified in a broad range of organi sms, including maize shoots, rice, soybean, cucumber seedlings, Enterococcus faecalis , Pseudomonas aeruginosa and Lactobacillus hilgardii (Arena and Manca de Nadra, 2001; Nakada et al. , 2001; Sakakibara and Yanagisawa, 2003; Simon and Stalon, 1982; Ya nagisawa, 2001). NCBI Blast searches have revealed potential AgDS genes in othe r organisms, although their functions have not been established. The physiological role of agmatine cata bolism in bacteria appears to vary, depending on the source of the agmatine. In organisms with an intracellular arginine decarboxylase (ADC), such as P. aeruginosa , agmatine is produced directly from arginine and is converted to Ncarbamoylputrescine by AgD (Nakada et al. , 2001). N-carbamoylputrescine is then hydrolyzed to putrescine, CO2 and NH3 by N-carbamoylputrescine amidohydrolase (Nakada et al. , 2001). Putrescine is converted to spermidine by spermidine synthase or broken down to succinate, which enters the TCA cycle. The AgDS in these bacteria may be an important source of polyamines, in addition to providing carbon and nitrogen. In ADC-deficient bacteria, such as E. faecalis , agmatine is derived from exogenous sources via an agmatine-putre scine antiporter (Driessen et al. , 1988; Griswold et al. , 2004b). In E. faecalis , the AgDS closely resembles the ADS, although it is important to
88 note that the ADS is not present in the genome of S. mutans . Biochemical analyses in E. faecalis (Roon and Barker, 1972) showed that the AgDS is highly analogous to the ADS, suggesting a role in acid tolerance, but the ge nes encoding this pathway or their mode of regulation have not been characterize d. Agmatine enters the cell via an agmatine:putrescine antiporter, where it is hydrolyzed to N-carbamoylputrescine and ammonia by agmatine deiminase (AgD; EC 3.5.3. 12). Putrescine carbamoyltransferase (PTC; EC 18.104.22.168) mediates the phosphorolysis of N-carbamoylputrescine, yielding putrescine and carbamoylphosphate. Finally, a phosphate group is transferred from carbamoylphosphate to ADP by carbamate ki nase (CK; EC 22.214.171.124), generating ATP, CO2 and NH3. Putrescine is then exchanged for agmatine via the antiporter. Polyamines, such as putrescine, are known to modulate tr anscription and protein synthesis in addition to conferring protection from acid (Dur and and Bjork, 2003; Tabor and Tabor, 1985; Yoshida et al. , 2002), so agmatine catabolism might pl ay multiple roles in the physiology of AgDS-positive bacteria. To begin to investigate the role of the AgDS in plaque ecology and S. mutans pathogenicity, we have initiated an analysis of the putative AgDS gene cluster. Results Analysis of the Sequence of the Agmatine Deiminase Gene Cluster The S. mutans AgD enzyme, encoded by SMU.264, shares significant homology with both predicted and established agmatine deiminases in at least 10 other bacteria, including the well-characterized AgD of P. aeruginosa (Nakada et al. , 2001). However, the remaining genes in the S. mutans AgDS most closely resemble those of E. faecalis, L. sakei, L. monocytogenes and L. lactis (Figure 5-1) with respect to operonic organization and predicted amino acid sequences . Of these bacteria, only E. faecalis has been fully
89 established as possessing a functional agmatine deiminase system (Simon and Stalon, 1982). The first gene in the operon, annotated as otcA , encodes a putative PTC and has been redesignated aguB . AguB is 80% identical to Ar gF-2, which is annotated as an OTC in E. faecalis V583 (Paulsen et al. , 2003). A lower level of similarity was evident between the S. mutans AguB and L. lactis subsp. lactis OtcA (78% identity), L. monocytogenes LMO0036 (65%), and Lactobacillus sakei ArgF (62%). ArgF-2 is annotated as one of two putative OTCases in the E. faecalis genome (Paulsen et al. , 2003), although our examination of the sequenc e revealed that the gene encoding ArgF2 is located in an apparent operon that en codes agmatine deiminase, carbamate kinase and an antiporter. The other OTC is in the ADS operon (Barcelona-Andres et al. , 2002), and therefore it is reasonable to suggest that ArgF-2 is actually the PTC described by Roon and Barker (Roon and Barker, 1972). Th e amino acid sequence of the PTC has not been published, but the enzyme has been well ch aracterized and it is distinctly different from OTC (Wargnies et al. , 1979). According to Prosite (http://www.expas y.org/prosite), a consensus pattern for carbamoyltransferases is F-x-[E/K]-x-S-[G /T]-R-T, with the third residue allowing differentiation between aspartat e carbamoyltransferase (E) or OTC (K) enzymes. This consensus sequence is present in th e predicted amino acid sequence of S. mutans aguB , as well as E. faecalis ArgF-2, L. lactis subsp. lactis OtcA , L. monocytogenes LMO0036, and L. sakei ArgF (Fig. 5-1), but a highly conserved Q is present in the third position, perhaps reflecting the preference of the enzyme for putre scine, rather than aspartate or ornithine. The HPTQ residues at positions 143-146 known to be involved in carbamoylphosphate
90 binding in OTCases are present in AguB, as are the HCLP residues at positions 281-284, which are conserved in OTCases and facilitate ornithine binding (Houghton et al. , 1984). Since the chemical structures of ornithine a nd putrescine are identical at the position of cleavage, this sequence cons ervation is not surprising. The second gene in the operon, anno tated SMU.263, encodes a putative amino acid antiporter and has been redesignated aguD . AguD is similar to proteins found in the putative AgDS gene clusters of other bacteria. AguD is 68% identical to E. faecalis EF0733, a predicted amino acid permease, 58% identical to the amino acid antiporter YrfD in L. lactis subsp. lactis , 46% identical to the ami no acid transporter LMO0037 in L. monocytogenes , and 25% identical to the probable amino acid permease PA4804 in P. aeruginosa . Similar to other amino acid antipor ters, 11 transmembrane helices were predicted for the amino acid sequence of AguD using the "DAS"-Transmembrane Prediction server at http://www. sbc.su.se/~miklos/DAS (Cserzo et al. , 1997). The third gene, annotated as SMU.264, encodes AgD and has been redesignated as aguA . S. mutans AguA shares significant homology with the known and putative AgD enzymes of at least 10 other bacteria. As demonstrated by CLUSTAL W (Thompson et al. , 1994) amino acid sequence alignments (Fi gure 5-2), several regions are highly conserved in AgD enzymes belonging to S. mutans, E. faecalis, L. lactis subsp. lactis , and L. monocytogenes , while other regions are conserved in AgD from all 10 bacteria. In particular, two conserved regions ar e obvious, the first occurring at residues 120137 in S. mutans AguA [FNAWGGLVDGLYFPWDQD] a nd the second at residues 157172 [DFVLEGGS(I/F)HVDG(E/Q)GT]. Th e [GGGNIHCITQQ] sequence noted elsewhere (Nakada et al. , 2001) was identified at the C-terminus of all 10 AgD enzymes.
91 S. mutans AguA shares the highest leve l of identity with YrfC in L. lactis subsp. lactis (66%), followed by conserved hypothetical protein EF0734 in E. faecalis (64%), which is most likely the AgD identified and char acterized by Simon and Stalon (Paulsen et al. , 2003; Simon and Stalon, 1982). Lower levels of homology were observed with LabD in L. sakei (56%) and AguA in P. aeruginosa (54%). SMU.264 is 56% and 52% identical to conserved hypothetical proteins LMO0038 and LMO0040 in L. monocytogenes , respectively. It is interesting to note that L. monocytogenes has two copies of putative AgD genes that share only 66% identity to each other. The first copy is present in an operon encoding a PTC, antiporter and CK, while the second copy is located approximately 100 base pairs downstream fr om these genes, next to a putative transcriptional regulator. The final gene in the S. mutans AgD gene cluster, annotated arcC , codes for a carbamate kinase that shar es homology with carbamate kinases of Streptococcus agalactiae (61%), Streptococcus suis (61%), Streptococcus pyogenes (60%), L. monocytogenes (56%), E. faecalis (54%), and L. sakei (50%). This gene should be redesignated as aguC to reflect involvement of th e protein in the AgDS, rather than the ADS. Thus, the S. mutans AgDS genes, redesignated as aguB, aguD, aguA and aguC , most likely encode a putative putrescine ca rbamoyltransferase, amino acid antiporter, agmatine deiminase and carbamate kinase, respec tively. In addition to these genes, there is an ORF encoding a transcrip tional regulator of the LuxR family located 239 base pairs upstream of aguB and transcribed in the opposite directi on. It is possible that this protein is involved in regulation of the AgDS in S. mutans . LuxR-type proteins belong to the FixJ-NarL superfamily, which is mainly comprised of two-component response
92 regulators involved in quorum sensing (Fuqua et al. , 2001). A putative nitroreductase and an ABC transporter are located upstream of this regulatory protein, however no sensor kinase is present. Potential homol ogs of this regulatory protein were also identified 399 and 234 base pairs upstr eam of the AgDS gene clusters in E. faecalis and L. lactis subsp. lactis , respectively. Much of the current information regarding AgDS enzymes was derived from biochemical experiments and very little is known of the molecular regulation of this pathway in other bacteria. Analysis of AgDS regulation has primarily focused on the aguBA operon of P. aeruginosa , which encodes N-carbam oylputrescine amidohydrolase and AgD, respectively. Immediately upstream of the aguBA operon is aguR , encoding a negative regulator of the AgDS pathway. In P. aeruginosa , the presence of 1 mM agmatine significantly inhibits AguRDNA interaction, facilitating induction of aguBA in the presence of agmatine (Nakada et al. , 2001). Thus, the ORF located immediately upstream of S. mutans aguB may play a similar role as the P. aeruginosa aguR in the regulation of AgDS. Involvement of this protein in the regulation of AgDS in S. mutans is described in Chapter 6. Agmatine Deiminase Expression in S. mutans Using RT-PCR analysis of mRNA from cells grown in TV containing 0.5% galactose and 10 mM agmatine, it was demons trated that cDNA can be amplified from aguB (Figure 5-3), providing evidence that th e AgDS gene cluster is transcribed in S. mutans UA159. No terminator-like seque nces were identified in the agu intergenic regions. To demonstrat e co-transcription of aguBDAC , a polar mutation in aguB was constructed by insertion of an Km cassette harboring strong transcription/translation
93 termination signals (Perez-Casal et al. , 1991). A Northern blot was performed (Ausubel et al. , 1989) using 10 g total RNA extracted (Chen and Burne, 1996) from wild-type and aguBcells grown in a low-carbohydrate tryp tone-vitamin (TV) based broth (Burne et al. , 1999) supplemented with 0.5% galact ose and 10 mM agmatine. The RNA was hybridized to an aguB probe, labeled using the Ambion Br ight Star labeling kit (Ambion Inc, Austin, TX). The probe hybridized to a 4.6 kb wild-type mRNA, consistent with the size of the operon (Figure 5-4). To quantify mRNA under different growth condi tions, slot blot analysis was used. Total RNA was extracted from cells grown in TV broth supplemented with 0.5% glucose or 0.5% galactose, with or without 10 mM agmatine. RNA was transferred to a nitrocellulose membrane and probed with an aguB probe. AgDS-specific mRNAs were detected under all growth conditions, although expression wa s several-fold higher when cells were grown in medium cont aining 10 mM agmatine (Figure 5-5). AgD activity in different growth cond itions was measured by colorimetric determination of N-carbamoylputrescine produc tion from agmatine (Figure 5-6, top). In wild-type S. mutans , peak AgD activity was observed in cells grown in galactose and agmatine, while activity decreased around 65% in cells grown in glucose and agmatine. Galactose has been shown to be a non-repressi ng sugar in oral streptococci, suggesting that the AgDS may be under th e control of carbohydr ate catabolite repr ession (CCR). No activity was observed when cells were gr own without agmatine. The pattern of expression of enzyme activity is consistent with the mRNA anal ysis, suggesting that agmatine is necessary for induction of the system. A polar mutation in aguB , the first gene in the apparent operon, was cons tructed and assayed for production of N-
94 carbamoylputrescine from agmatine. The aguB mutation abolished production of Ncarbamoylputrescine from arginine, proving that the genes are involved in agmatine utilization and that they are probably tr anscribed from a single promoter, because aguA is the last gene in the cluster. Agmatine Deiminase Specificity To determine if AgD was specific for agmatine, wild-type S. mutans was grown under inducing conditions and assayed for the ab ility to produce citrulline from arginine. Citrulline was not detected ev en after the reaction time wa s extended to several hours, suggesting that AgD is specific for agmatine. Additional experiments were conducted to determine if the S. mutans AgDS could produce N-carbam oylputrescine from primary amines sharing structural similarity with ag matine (Figure 5-7). For these experiments, S. mutans was grown in TV media containing 25 mM galactose and 10 mM of agmatine, -alanine, -amino butyric acid (GABA), histamine, cadaverine or ornithine. To determine if these primary amines could induce the AgDS, bacteria grown in each condition were assayed for the ability to produce N-carbamoylputrescine from agmatine. Significant induction of the AgDS was only observed when cells were grown in the presence of 10 mM agmatine. However, low le vels of activity were detected when cells were grown in the presence of ot her amines, suggesting that the cis or trans -acting factors that mediate activation of AgD may be capable of responding, to a much lesser degree, to other primary amines as well as agmatine. It is noteworthy that agmatine is requir ed for induction of the AgDS from the perspective of plaque ecology. There apparen tly is no arginine decarboxylase activity in
95 S. mutans , and thus, a close association with ar ginine decarboxylase-producing organisms in dental plaque could benefit S. mutans . Ammonia Production from Agmatine Consistent with AgD enzyme activity, ammonia production from agmatine was only observed in cells grown in media cont aining agmatine (Figure 5-6, bottom). In addition, ammonia producti on increased about two-thirds in wild-type S. mutans grown in galactose and agmatine, as compared to those grown in glucose and agmatine. Ammonia production by the AgDS was appr oximately two-fold higher than Ncarbamoylputrescine production, cons istent with the fact that one mole of agmatine yields two moles of ammonia. The polar mutation in aguB eliminated ammonia production from agmatine. The optimum pH for ammonia production fr om agmatine appears to be around pH 4, although the system is capable of producing ammonia at a broad range of pH values and activity drops off sharply below 4.0 (Figur e 5-8). The ability to catabolize agmatine at pH 4 implies that the AgDS has the potential to provide S. mutans with a competitive advantage over other oral bacteria during environmental acidifica tion. Specifically, the production of ammonia from agmatine by S. mutans at low pH would increase pH and provide ATP, which could be used for growth or to move protons out of the cell. Since the minimum pH observed in dental plaque after carbohydrate cons umption is usually around pH 4 (Stephan, 1940), the Ag DS may be a significant fact or in acid tolerance of S. mutans in vivo. Given that S. mutans is not capable of producing ammonia from agmatine at pH values below 3.5, measuring the capacity of the AgDS to protect the organisms from acid killing was not practical since efficient killing of strain UA159 does
96 not occur at values above pH 2.8. Attemp ts to perform acid killing assays on UA159 wild-type and the polar aguB mutant were further hindered because agmatine itself protected both strains agains t killing (Figure 5-9). As the presence of agmatine is required for AgD expression, these problems made it difficult to rely on acid killing assays to explore the role of the AgDS in acid tolerance. Instead, the effects of agmatine on the acid:base physiology of S. mutans is explored in Chapter 6 using continuous culture experiments. It has been established that the structurally related am ino acid, arginine, is capable of alkalinizing dental pla que via the arginine deimin ase pathway (Wijeyeweera and Kleinberg, 1989a). It is therefore noteworthy that th e highest levels of AgDS activity detected in S. mutans are some 200-fold lower than AD activity in S. rattus (Griswold et al. , 2004a) or S. gordonii (Dong et al. , 2002) when these systems are fully induced. While it is possible that the conditions used to induce AgDS activity in this study were not optimal, it is more likely that the lower le vel of expression may be related to either substrate availability in vivo or to the role of the system in S. mutans. In particular, if agmatine is present in much lower concentr ations in the mouth than arginine, which seems likely, then a highly active AgDS sy stem may not be needed to catabolize the amount of substrate that is typically available in vivo . Another compatible explanation is that low levels of agmatine catabolism woul d produce sufficient intracellular ammonia to neutralize the cytoplasm, but would not have a major impact on the surrounding pH. By catabolizing agmatine at a relatively slow rate, S. mutans could reap the benefits of intracellular ammonia production without concu rrently increasing the extracellular pH
97 and providing its less acid-tolerant, AgDS-deficient competitors with more favorable growth conditions. Identification of the AgDS in Related Viridans Streptococci To determine if the AgDS is conserved among other oral streptococci, production of N-carbamoylputrescine from agmatine was assayed in S. mutans UA159 and eight additional strains of oral stre ptococci (Figure 5-10). The highest levels of AgD enzyme activity were observed in S. mutans and S. rattus strains FA-1 and BHT. AgD activity was significantly lower in Streptococcus sobrinus , a third member of the mutans group, as well as in Streptococcus sanguis , Streptococcus oralis and Streptococcus salivarius . AgD activity was not detected in S. gordonii . S. mutans aguA was used in blastx searches of the available streptococcal genomic sequences in the NCBI, TIGR and Sanger databases. Homologs of S. mutans aguA were identified in the genomes of S. sobrinus (72% ID), Streptococcus uberis (46% ID), Streptococcus pneumoniae (45% ID), and Streptococcus mitis (44% ID). The genomic sequences of S. sanguis, S. oralis , and S. salivarius are not available. Homologs of aguA were not found in the genomes of S. gordonii , Streptococcus suis , Streptococcus thermophilis , Streptococcus pyogenes , Streptococcus agalactiae , Streptococcus equi , or Streptococcus zooepidemicus . Because the genome of S. rattus is not available, the agu operon of this bacterium was sequenced to determine relative homology to the agu genes in S. mutans (Figure 511). Not surprisingly, the S. rattus agu genes are organized in an aguRBDAC cluster and the predicted amino acid sequences of thes e ORFs are nearly identical to their counterparts in S. mutans , sharing 100%, 98%, 97%, 79% and 94% identity with AguR, AguB, AguD, AguA, and AguC of S. mutans . Northern blot anal ysis of RNA isolated
98 from S. rattus grown in the presence and absence of 10 mM agmatine confirmed that the agu operon is transcribed on a fragment of 4.4kbp, and is transcriptionally induced in the presence of agmatine. AgD assays in the presence and absence of agmatine confirmed induction by agmatine (Figure 5-12). However, consistent with previous observations regarding CCR regulation of catabolic systems in S. rattus , CCR did not appear to play a major role in AgDS regulation in this bacterium when cells were grown in 25 mM glucose or galactose (Burne et al. , 1991; Griswold et al. , 2004a). However, when S. rattus was grown in 2% carbohydrate, the effect s of catabolite repression were more apparent and the effects of agmatine induction were not statistically significant (Figure 513). Interestingly, AgD expres sion was observed even in the absence of agmatine when cells were grown in 2% carbohydrate. The final pH of the spent growth media was considerably lower, suggesting that the AgDS may be acid-inducible, as well as agmatine-inducible in S. rattus . A pH effect was also observed for the S. mutans AgDS and is discussed in detail in Chapter 6. The physiological function of the AgDS in or al streptococci is unclear. However, it is interesting to note that several AgDS-positive bacteria, such as S. rattus , S. sanguis , S. oralis and S. salivarius , also utilize other, highly-activ e, ammonia generating systems, such as the ADS (Burne et al. , 1989; Griswold et al. , 2004a) and urease (Chen et al. , 2002). In the dental plaque biofilm, organi sms undoubtedly benefit from the ability to metabolize a wide variety of substrates, de pending on nutrient availability. The AgDS may be especially advantageous because it al lows generation of ammonia and ATP. It is possible that the AgDS either enhances am monia and energy production in these bacteria, or serves an additional purpos e unrelated to acid tolerance. It is also noteworthy that
99 AgD enzyme activity was significantly higher in S. mutans and S. rattus than in the other oral streptococci examined. In vivo , levels of AgD expression could be related to the maturity and complexity of the dental plaque biofilm. Unlike S. mutans (Nyvad and Kilian, 1990), S. sanguis , S. oralis and S. gordonii are early colonizers of dental plaque (Kolenbrander and London, 1993; Kolenbrander et al. , 2002; Li et al. , 2004). If S. mutans resides in close proximity to bacteria that produce agmatine from arginine via arginine decarboxylase (ADC), then elevated AgD expression would be beneficial. However, if fewer ADC-utilizing bacteria are pr esent in the portion of the plaque biofilm colonized by S. sanguis , S. oralis and S. gordonii , then expression of AgD would not be particularly be neficial. To investigate this theory further, blas t searches were performed to find oral bacteria that carry ADC genes. The results demonstrated that Wolinella , Prevotella , Bacteroides , Desulfovibrio , and Neisseria species possess the ADC enzyme, while no evidence was found of the ADC enzyme in Porphyromonas gingivalis , oral streptococci, Actinobacillus acti nomycetemcomitans , Actinomyces naeslundii , Fusobacterium nucleatum , Treponema denticola , and Lactobacillus species , albeit not all of these genomes are complete. Although the specifi c relationships that exist between the hundreds of bacterial species in the oral cavity are largel y unknown, it has been established that S. gordonii co-aggregates with P. gingivalis (Brooks et al. , 1997; Lamont et al. , 1992), an ADC-negative bacteria, and that S. oralis , S. mitis and S. sanguis are known to co-aggregate with Actinomyces species (Cisar et al., 1979; Ganeshkumar et al. , 1991; McIntire et al. , 1978), which also appear to be ADC-deficient. Co-aggregation of S. mutans has been observed with various or al bacteria, some of which are ADC-
100 positive, including Veillonella , Nocardia , Neisseria , Actinomyces and Candida albicans (Cisar et al. , 1979; Egland et al. , 2004; Ellen and Balcerza k-Raczkowski, 1977; Gibbons and Nygaard, 1970; Kara et al. , 2006; McBride and van der Hoeven, 1981; Mikx and Van der Hoeven, 1975; Miller and Kleinman, 1 974). Therefore, it is possible that the AgDS is a convenient means for S. mutans to generate ATP and ammonia when associated with ADC-positive bacteria in th e dental plaque biofilm. The physiological role of agmatine catabolism in S. mutans is discussed in more detail in Chapter 6, while future studies are necessary in order to dissect the relationshi ps between AgDSand ADC-utilizing bacteria in various niches of the oral cavity. Summary We have characterized a cluster of genes ( aguRBDAC ) in S. mutans UA159 that are involved in agmatine utilization and appear to be most closely related to the E. faecalis AgDS. These genes, originally annotated as otcA, SMU.263, SMU.264 and arcC , appear to encode a putative putresc ine carbamoyltransferase, am ino acid antiporter, agmatine deiminase and carbamate kinase, respectively. These enzymes are components of the agmatine deiminase system (AgDS), which ge nerates putrescine, ATP, carbon dioxide and ammonia from agmatine, a decarboxylated derivative of arginine. The AgDS appears to have been conserved in many of the viridans streptococci, particularly S. rattus , S. sanguis , S. sobrinus , S. oralis and S. salivarius . Expression of the system requires the presence of agmatine and the genes are regulated by carbohydrate catabolit e repression. A polar inserti on in the first gene in the operon abolished agmatine deiminase activity and ammonia production from agmatine. The ADS of S. mutans was shown to be capable of pr oducing ammonia from agmatine at pH values as low as 4, suggesting that th is pathway may provide an ancillary acid-
101 tolerance mechanism. The identifica tion of an ADS-like gene cluster in S. mutans is significant, given the contribu tion of acid tolerance to viru lence. Production of base by S. mutans may have critical implications for the pa thogenicity of this organism and for the ecology of dental plaque.
102 Figure 5-1. The putative AgDS gene cluster in S. mutans UA159 and percent identity to similarly organized clusters in other bacteria, based on NCBI Blast. PTC, putrescine carbamoyltransferase; Port, agmatine:putrescine antiporter; AgD, agmatine deiminase; CK, carbamate kinase; N/A, xasA sequence is not available. Source: GenBank, National Center fo r Biotechnology Information. L. sakei argF xasA labD epkA PTC Port AgD CK E. faecalis argF-2 EF0733 EF0734 EF0735 L. monocytogenes LMO36 LMO37 LMO38 LMO39 80% 68% 64% 54% 62% N/A 56% 50% 65% 46% 56% 56% S. mutans aguB aguD aguA arcC 1.5-kbp 1.36-kbp 1.11-kbp 0.948-kbp
103 S. mutans 120FNAWGGLVDGLYFPW -134 L. lactis 116FNAWGGLVDGLYFPW -130 E. faecalis 88FNAWGGLVDGLYFPW -102 L. mono 114FN S WGGLVDGLYFPW -128 L. sakei 114FNAWGGLVDGLYFPW -128 P. aeruginosa 115FNAWGG FEG GLYFPW -129 Y. pestis 123FNAWGGL NG GLY AD W -137 S. pneumoniae 111FNAWGG TY DGLY QDY-125 S. avermitilis 112FN G WG---AQSWAR W -123 S. coelicolor 106FN G WG---AQDWAR W -117 157DFVLEGGSIHVDGEGT -172 153DFVLEGGS F HVDG Q GT -168 125DFVLEGGS F HVDG Q GT -140 151DFVLEGGSIHVDGEGT -166 151D L VLEGGS T HVDGEGT -166 152DFVLEGGSIHVDGEGT -167 159-PLI LEGGSIH T DGEGT -174 148-P FVLEGG A IH S DG Q GT -163 145-KL V N EGG A IHVDGEGT -160 139-PL V N EGG A IHVDGEGT -154 Figure 5-2. Conserved regions of actual and putative agmatine deiminase enzymes based on CLUSTAL W formatted alignments of predicted amino acid sequences. The GGGNIHCITQQQP motif that was prev iously identified by Nakada et al. (2000) was identified at bases 355-367 in S. mutans UA159 (not shown). Comparisons were made between the S. mutans AgD enzymes and those indicated, where L. mono = Lister ia monocytogenes and the two bottom sequences are derived from Streptomyces avermitilis and coelicolor.
104 Figure 5-3. Reverse transcriptas e PCR analysis of mRNA from S. mutans grown in TV containing 0.5% galactose and 10 mM agmatine. Primers specific to aguB were used to amplify cDNA. Lane 1, molecular weight markers; Lane 2, the product amplified from S. mutans c hromosomal DNA; Lane 3, the product obtained from cDNA; and Lane 4, cont rol in which a reaction containing mRNA with no reverse transcriptase wa s used in the amplification reaction. 1 2 3 4
105 Figure 5-4. Northern blot analysis of total RNA isolated from wild-type and aguBS. mutans grown in TV broth containing 0.5% galactose and 10 mM agmatine. 10 g RNA was hybridized to an aguB DNA probe. WT aguBaguB 4.6 kb
106 Figure 5-5. Slot blot analysis of mRNA isolated from wild-type S. mutans grown in TV broth containing 0.5% glucose (Glu), 0.5% glucose and 10 mM agmatine (GluAg), 0.5% galactose (Gal) or 0.5% galactose and 10 mM agmatine (GalAg). The mRNA was transferred to a nitrocellulose membrane and probed with aguB DNA. RNased indicates a control in which 2 g of RNA was treated with RNase prior to application to the membrane. RNased 2 g GluAg Glu Gal GalAg 1 g 2 g
107 Figure 5-6. A) Agmatine deiminase enzyme activity (nmol N-carbamoylputrescine produced/min/mg protein) of S. mutans (WT) grown in TV containing glucose (Glu), glucose and agmatine (GluAg), galactose (Gal) or galactose and agmatine (GalAg). Activity of a polar mutant (AguB ) was measured in cells grown in galactose and agmatine. B) Production of ammonia from agmatine (nmol ammonia/min/mg protein) by WT grown in TV containing Glu, GluAg, Gal or GalAg. Activity of AguB was measured in TV containing galactose and agmatine. Results shown are the av erage and standard deviations (error bars) of a minimum of 9 separate cult ures for each strain and condition. Oneway ANOVAs and pair-wise Student t-test s were used to identify significant differences (p < 0.05) in AgD activity. Si milar letters indicate no significant difference in activity. 0 10 20 30 40 50 60 70 80 90 100 A. WT Gal WT GluA g AguB-GalAg WT Glu WT GalAg 0 20 40 60 80 100 120 140 160 180 200 220 B. WT Glu WT GluAg AguB-GalAg WT GalAg WT Gal b c a a a b c a a a AgD Activity U (mg protein)-1 Ammonia Production nmol/min/mg protein
108 Figure 5-7. AgD activity of wild-type S. mutans UA159 grown under inducing conditions and assayed for production of N-carbam oylputrescine from various primary amines. Activity is expressed as nmol N-carbamoylputrescine produced/min/mg protein. Results show n are the average and standard deviations (error bars) of a minimum of 9 separate cultures for each strain and condition. One-way ANOVAs and pair-wise Student t-tests were used to identify significant differences (p < 0.05) in AgD activity. Similar letters indicate no significant difference in activity. 0 20 40 60 80 100 Gal Agmatine -Alanine GABA Histamine Cadaverine Ornithine 120 a ac cd b cd c c AgD Activity U (mg protein)-1
109 Figure 5-8. Ammonia production from agmatine at various pH values by wild-type S. mutans that was grown in TV broth c ontaining 0.5% galactose and 10 mM agmatine. Ammonia production is expr essed as nmol ammonia /min/mg protein. Results shown are representativ e of three independent experiments. 0 50 100 150 200 250 300 350 3.0 4.0 5.06.07.08.0 pHAmmonia (nmol/min/mg protein)
110 -6.00 -5.00 -4.00 -3.00 -2.00 -1.00 0.00 03060 Time (min)Log [N/No] WT Gal + WT GalAg + BGal + BGalAg + WT Gal WT GalAg BGal BGalAg Figure 5-9. Survival of wild-type S. mutans UA159 (WT) and an AgDS-defective strain harboring a polar mutation in aguB (B-) in 0.1 M glycine buffer, pH 2.8. Results shown are representative of three independent experiments.
111 Figure 5-10. A) AgD enzyme activity in select ed oral streptococci, expressed as nmol Ncarbamoylputrescine produced/min/mg prot ein. Results shown are the average and standard deviations (error bars) of a minimum of 9 separate cultures for each strain and condition. ND: not det ected. B) Other ammonia-generating systems found in each organism. One-way ANOVAs and pair-wise Student ttests were used to identify significant differences (p < 0.05) in AgD activity. Similar letters indicate no si gnificant difference in activity. Strain AgD Activity (TV GluAg) Other Known NH3 (nmol/min/mg protein) Systems Present S. mutans UA159 57.3 (7.45) none S. mutans GS-5 76.7 (0.57) none S. rattus FA-1 31.34 (6.0) ADS S. rattus BHT 52.4 (4.9) ADS S. sobrinus SL-1 3.5 (1.20) none S. sanguis 10556 10.0 (1.56) ADS S. oralis SK92 25.7 (3.20) ADS S. salivarius 57.I 10.0 (7.23) Urease S. gordonii DL-1 0.0 (0.00) ADS 0 10 20 30 40 50 60 70 80 90 S. mutans UA159 S. mutans GS-5 S. rattus FA-1 S. rattus BHT S. sobrinus SL-1 S. sanguis 10556 S. oralis SK92 S. salivarius 57.I S. gordonii DL-1 N DB A a a b c c d e de AgD Activity U (mg protein)-1
112 Figure 5-11. A) Northern bl ot of RNA isolated from S. rattus FA-1 grown in the presence and absence of 10 mM agmatine, using an aguA DNA probe derived from S. rattus FA-1. B) The agu operon of S. rattus FA-1 and homology of each ORF to the corresponding gene in S. mutans UA159. The molecular weight and pI of each predic ted AgD protein is shown for S. rattus (black) and S. mutans (parentheses, grey). 10 mM Agmatine: + 4.40 kb aguR aguB aguD aguA aguC pI (UA159) : 9.00 (9.00) 4.74 (4.83) 9.14 (9.14) 4.61 (4.63) 6.26 (5.21) MW, kDa (UA159) : 37.6 (37.6) 39.8 (39.8) 49.6 (49.6) 34.5 (41.8) 33.9 (33.8) % Identity (aa) to S. mutans UA159: 100% 98% 97% 79% 94% B A
113 Figure 5-12. AgD enzyme activity of S. rattus FA-1 grown in TV medium containing 25 mM glucose or galactose, with or without 10 mM agmatine. Activity is expressed as nmol N-carbamoylputresc ine produced/min/mg protein. Results shown are the average and standard deviat ions (error bars) of a minimum of 9 separate cultures for each strain and condition. One-way ANOVAs and pairwise Student t-tests were used to iden tify significant differences (p < 0.05) in AgD activity. Similar letters indicate no significant difference in activity. 0 5 10 15 20 25 30 35 40 45 Glu Glu Ag Gal Ag Gal a b a bAgD Activity U (mg protein)-1
114 Figure 5-13. AgD enzyme activity of S. rattus FA-1 grown in TV medium containing 2% glucose or galactose, with or without 10 mM agmatine. Activity is expressed as nmol N-carbamoylputrescine produ ced/min/mg protein. Results shown are the average and standard deviations (e rror bars) of a minimum of 9 separate cultures for each strain and conditi on. One-way ANOVAs and pair-wise Student t-tests were used to identify significant differences (p < 0.05) in AgD activity. Similar letters indicate no significant difference in activity. 0 10 20 30 40 50 60 70 80 Glu Glu Ag Gal Gal Ag b a a abAgD Activity U (mg protein)-1
115 CHAPTER 6 REGULATION AND PHYSIOLOGIC SI GNIFICANCE OF THE AGMATINE DEIMINASE SYSTEM OF S. mutans UA159 Introduction Recently, we described a pathway for ammonia production in S. mutans that has not yet been identified in oral bacteria (Griswold et al. , 2004b). Alkali production by the agmatine deiminase system (AgDS) ma y increase the competitive fitness of S. mutans , contributing in major ways to the persisten ce and pathogenesis of this organism. We previously demonstrated that Streptococcus mutans expresses a functional agmatine deiminase system (AgDS) encoded by the agmatine-inducible aguBDAC operon (Griswold et al. , 2004b). The AgDS yields ammonia, CO2 and ATP while converting agmatine to putrescine, and is proposed to augment the acid resistance properties and pathogenic potential of S. mutans . The agu operon in S. mutans is induced in the presence of agmatine and is regulated by carbon catabolite repression (CCR) (Griswold et al. , 2004b). Overall, the AgDS is expressed at a rela tively low level compared to other ammonia-generating pathways of oral streptococci, and it is unlikely that agmatine catabolism results in significant environmental alka linization. However, ammonia production by the AgDS under acidic conditions would increase pH and provide ATP, thereby contributing to acid tolerance and growth at low pH, which w ould substantively augm ent the virulence of the organism . In order to better understand the role of the AgDS in the physiology and virulence of S. mutans , we have initiated a study of th e factors regulati ng the AgDS in
116 this organism (Griswold et al. , 2006). We have also described a novel mechanism by which S. mutans copes with the production of an antagonistic compound generated in response to environmental acidification of oral biofilms by competing organisms (Griswold et al. , 2006). Results Role of the S. mutans SMU.261c Protein Previously, we reported that efficient expression of the S. mutans AgDS requires agmatine (Griswold et al. , 2004b). To date, the only reported trans -acting factor involved in AgDS regulation is AguR, a TetR-type transcri ptional repressor encoded by an ORF located immediately upstream of the P. aeruginosa aguBA operon. In P. aeruginosa , the presence of 1 mM agmatin e significantly inhibits AguR-DNA interaction, facil itating induction of aguBA in the presence of agmatine (Nakada et al. , 2001). Although there were no ORFs that showed substantial similarity to P. aeruginosa AguR in the S. mutans genome, a putative LuxR-like tran scriptional regulator, SMU.261c (NCBI database), was identified 239 bases upstream of aguB and transcribed in the opposite direction (Figure 6-1). LuxR-t ype proteins belong to the FixJ-NarL superfamily, which is mainly comprised of two-component response regulators involved in quorum sensing (Fuqua et al. , 2001). Using NCBI Blastp, it was determined that SMU.261c shared the highest levels of simila rity with putative LuxR -like regulators that are encoded approximately 399 and 2 34 bases upstream of the putative agu operons of E. faecalis and L. lactis subsp. lactis (62% and 56% similarity, re spectively). A C-terminal helix-turn-helix (HTH) domain, implicated in DNA binding by members of the LuxR family, was identified in SMU.261c, as well as in the E. faecalis and L. lactis ORFs. However, the conserved acylated homoserine lactone (acyl-HSL) binding region in LuxR
117 could not be found in any of the three prot eins. Interestingly, three transmembrane domains were also predicted to occur in the SMU.261c protein, so it is possible to predict a topology for the protein that could result in exposure of a substa ntial portion of the molecule to the external environment. The SMU.261c gene was mutated by allelic exchange with the insertion of an EmR marker ( aguR ), using the PCR ligation mutagenesi s method described by Lau et al. (Lau et al. , 2002). When increasing concentrations of agmatine were added to the growth medium, AgD activity increased prop ortionately in UA159, whereas the aguR strain displayed a low basal level of AgD activity, regardless of the agmatine concentration in the growth media (Figure 6-2). This data suggests that AguR could be a bifunctional regulator capable of repressing the AgDS in the absence of agmatine and inducing the system in the presence of agmatine. Thus, SMU.261c was named aguR to reflect a role in regulation of the AgDS. Localization of paguB and paguR Primer extension was used to map the aguB and aguR transcription initiation sites (TIS) (Figure 6-3). A single band correspondi ng to a G residue 22 bases upstream of the aguB start codon was observed. Examination of the upstream sequence revealed a putative 70-like promoter. The region ( TAT G AT ) shared 5 out of 6 bases with the consensus sequence (bold), whereas the region ( T A G TA A ) identified 18 bases upstream of the region shared only 3 base s with the consensus (bold). A single band corresponding to an A residue 34 bases upstream of the aguR start codon was observed. Examination of the upstream sequence revealed a putative 70-type promoter. The region ( TATAAT ) was identical to the consensu s sequence (bold), whereas the
118 region ( TT C A AT) identified 18 bases upstream of the region shared only 3 bases with the consensus (bold). Carbon Catabolite Repression We have demonstrated that the S. mutans agu genes are transcri ptionally repressed in the presence of the preferred ca rbohydrate source glucose (Griswold et al. , 2004b). In AT-rich gram-positive bacteria, CCR is mediated by the trans -acting catabolite control protein A (CcpA), which bi nds to highly conserved, cis -acting catabolite response elements ( cre ) during growth on preferred carboh ydrate sources to regulate the expression of catabolic genes and operons (Saier et al. , 1996). Two potential CcpAdependent cre sites were identified at ( TGTAATCGTTTACA ) and ( TGAAAA G GCTTT GT) relative to the aguB transcriptional start site, with bases matching the consensus shown in bold (Figure 6-3) (Hueck et al. , 1994). The identification of putative cre sites upstream of paguB, as well as the AgDS regulation patterns observed previously, prompt ed us to investigate the role of CcpA (RegM) (Simpson and Russell, 1998) and CcpB-like proteins (SMU.105, RegA) (Wen and Burne, 2002) of S. mutans in AgDS regulation. Initially, we attempted to examine the role of the putative cre sites in catabolite repression by cloning deletion derivatives of paguB behind the cat reporter gene, integrating these c onstructs into the chromosome of UA159 wild-type and aguR strains at an unrelated locu s, and comparing CAT activity after growth in the preferred sugar, gluc ose, and the non-repressing sugar, galactose (Figure 6-4). Expression from paguB was extremely low; although this was expected because AgD enzyme activity in this bacteria is relatively low, approximately one log lower than AD activity observed in other streptococci (Dong et al. , 2002; Griswold et al. , 2004a). However, data from this experiment clearly demonstrates that the full length
119 paguB is expressed at a higher level during growth in galactose, in agreement with data obtained from AgD enzyme assays. Deletion of the cre substantially reduced expression from paguB, and deletion of both the and cre sites reduced expression beyond the limit of detection. It is reasonable to conclude that the cre sites are required for optimal expression from paguB, although their role in catabo lite repression is unclear. Several palindromic sequences th at could potentially serve as trans -acting sites for transcriptional regu lators were identif ied within the 250-bp region upstream of aguB. Thus, it is possible that deletion of one or both cre sites inadvertently removed the AguR binding site, which has not yet been identified. To continue evaluating the role of CcpA and CcpB in catabolite repression of the S. mutans AgDS, AgD activity was measured in S. mutans UA159 and in otherwiseisogenic mutants of this strain lacking the ccpA or ccpB genes that were constructed previously in our laboratory (Wen and Burne, 2002). The strains were grown in TV broth containing 25 mM glucose or the non-repressing sugar, ga lactose, with or without 10 mM agmatine, to mid-exponential phase prio r to measurement of enzyme activity. Mutation of ccpA or ccpB did not alleviate catabolite repression of the AgDS (Figure 65), although AgD activity was slightly higher in the ccpA and ccpB strains following growth in either glucose or galactose, supplemented with ag matine. This observation is consistent with studies of other metabolic pathways in S. mutans that showed that even though cre sequences were tightly linked to the regulatory regions, that CcpA (RegM) and CcpB are not primary factors controlli ng CCR in this organism (Wen and Burne, 2002).
120 Regulation of the AgDS by Growth Phase and Environmental Stress Identification of environmental factors that regulate expression of bacterial genes can provide much insight into the role of the gene products. Thus, we began our investigation of the environm ental factors regulating the S. mutans AgDS by examining AgD activity in relation to grow th phase. Batch cultures of S. mutans UA159 were grown in TV medium containing 25 mM galact ose, with or without 10 mM agmatine, and samples were taken during ear ly-exponential phase, mid-expone ntial phase and two hours after the cultures entere d stationary phase (OD600 = 0.35, 0.5 and 0.65, respectively). The pH of each culture medium was measured and AgD assays were performed. AgD activity was 1.5-fold higher in stationary phase compared with mid-exponential phase when agmatine was included in the growth media (Figure 6-6). Enhancement of AgD expression during post-exponential phase growth could be due to nutri ent depletion or to the reduced pH of the grow th medium, which is pH 5.5 during stationary phase, as compared to pH 6.2 during exponential phase. When batch cultures were grown in TV medium buffered at pH 7, stationary phase AgD induction of approximately 1.5-fold was still observed (Figure 6-7), but enzyme activity levels in all cases were lower than in unbuffered medium. Thus, some induction of the AgDS occurs duri ng stationary phase, probably due to partial alle viation of CCR or another pa thway that senses nutrient depletion. However, AgDS induction is clearly enhanced at low pH values (Figures 6-8 and 6-9) and we determined that ammonia production from agmatine by intact cells of S. mutans is optimal at pH 4 (Griswold et al. , 2004b). To examine the effects of low pH on Ag D activity under conditions where growth phase, growth rate and glucos e availability could be tigh tly controlled, steady-state continuous cultures of S. mutans were maintained in a Biostat i chemostat at pH 5 or 7 as
121 detailed in the methods section. AgD activit y was up-regulated 4-fold at pH 5 compared to pH 7 in agmatine-induced cultures (Figure 6-8). Of note, AgD activity was significantly lower in chemostat cultures than in batch cultures. This discrepancy was most likely caused by differences in the me thods of AgD induction between the two experiments. Specifically, in the chemostat studies the vessel was pulsed with a final concentration of 3 mM agmatine for one hour prior to sampling, whereas batch cultures were grown in medium containing 10 mM agmatine. This was done because the presence of an ammonia-yielding compound dur ing the fermentation makes it difficult to maintain acidic pH in the vessel, aside from the fact that it is pr ohibitively expensive to include agmatine in the continuous cultures. Al so of interest is that a low level of AgD activity was observed in continuous cultures in the absence of agmatine, whereas the substrate was required for AgD expression in batc h cultures. It may be possible to detect some AgD activity in batch cultures (grown without agmatine) if the reaction times are extended. In any case, this obs ervation may be attributable to alleviation of residual CCR under glucose-limiting conditions in the chemosta t. Also, the slower growth rate in the chemostat ( tg = 2.3 h) may also contribute to dere pression of the AgDS, since expression of some gene products required for energy ge neration can be impacted by growth rate (Len et al. , 2003). This could also partially expl ain the induction of the AgDS during stationary phase. Consistent with measurements of Ag D enzyme activity, the expression of aguA , as measured by Real-Time RT-PCR, was approxima tely 3.7-fold higher in cells grown at pH 5 versus pH 7 (Figure 6-9) . Thus, transc ription of the aguBDAC operon is activated at acidic pH levels frequently encountered in dental plaque. I nduction of AgDS at low
122 pH supports the notion that the system may be a compone nt of the adaptive acidtolerance response by S. mutans. Also of note, many bact eria induce arginine decarboxylase expression when exposed to acid stress (Hayes and Hyatt, 1974; Lin et al. , 1995). Consequently, agmatine would likel y be in greater a bundance at low pH in vivo , and the combination of low pH and agmatin e could result in optimal AgD expression, similar to what has been observed for lysine decarboxylase and arginine decarboxylase in E. coli (Meng and Bennett, 1992; Richard and Foster, 2004). To determine if the AgDS could be part of a general stress response pathway in S. mutans , the effects of heat, oxidative and salt stresses were examined. AgD activity was 4-fold higher when S. mutans was grown to mid-exponential phase at 42 C, as compared to growth at 37 C (Figure 6-10), implying that this sy stem is responsive to environmental stresses other than low pH. However, oxi dative or salt stress had no effect on AgD activity (Figures 6-11 and 6-12). Consensus binding site s for the heat shock regulators HrcA and CtsR (Lemos et al. , 2001; Lemos and Burne, 2002) were not identified in the aguB promoter region. Whethe r heat stress acts through aguR specific regulators or through other pathways remains to be determined. Agmatine Deiminase and Biofilm Ecology Oral biofilms are complex ecosystems with hundreds of metabolically and physiologically diverse species. The ability of S. mutans to catabolize agmatine at low pH could impart a selective advantage to this organism through generation of ATP for growth and alkalinization of the cytopl asm by ammonia, which would reduce the investment of ATP in proton extrusion. Under such conditions, S. mutans would gain a competitive advantage over less acid-tolerant sp ecies in oral biofilms, particularly since the system is a low activity system that would not profoundly effect environmental
123 alkalinization. To test th is hypothesis, a competition experiment was performed using S. mutans UA159 and its AgDS-deficient derivative, aguB . Individual cultures were grown separately to OD600 = 0.4 and combined in equal volumes in TV media containing galactose and 0 mM or 20 mM agmatine. The st rains grew equally well in the absence of agmatine. Surpisingly, in the presence of 20 mM agmatine, the doubling time of the wildtype strain was significantly slower and the aguB strain was unable to grow. To further investigate the mechanism of growth inhibition by agmatine, the gene for carbamate kinase was mutated ( aguC ), which would allow the organisms to degrade agmatine, while they would be unable to produce ATP, carbon dioxide and the second mole of ammonia from agmatine. The aguC strain was able to gr ow in the presence of agmatine, albeit not as rapidly as the wild-t ype, suggesting that in hibition of growth by agmatine is the primary explanation for the phenotype displayed by the aguB strain (Figure 6-13). Agmatine inhibition was dose-dependent in the wild-type, aguB and aguC strains (Figure 6-14). However, while mutation of aguB eliminated AgD activity (Figures 5-4 and 5-6) and conveyed an ag matine-sensitive growth phenotype (Figure 614), mutation of aguR resulted in low AgD expression re gardless of the concentration of agmatine in the surrounding media (Figure 6-2), and conveyed an agmatine-resistant growth phenotype, even in the presence of 20 mM agmatine (Figure 6-14). To continue to explore the role of aguR in growth inhibition, an aguRB double mutant strain was generated. When grown in TV medium containing galactose and 20 mM agmatine, the aguRB strain displayed an intermedia te growth phenotype, restoring the ability of aguB to grow in agmatine, although not to the agmatine-re sistant phenotype exhibited by aguR (Figure 6-15). Further experime nts must be performed before
124 conclusions can be drawn about the involve ment of AguR in growth inhibition by agmatine. Of note, growth inhibition by ag matine was also observed in other AgDSutilizing oral streptococci (Figures 6-16, 6-17), suggesting that the AgDS has been conserved in oral streptococci as a mechan ism to cope with the presence of a toxic compound. The basis for inhibition of growth by ag matine is yet to be disclosed, although it may involve competitive inhibition of amino aci d transport or perhaps interference with translation. Inhibition by agmatine was less evident when the organisms were grown in peptideand carbohydrate-rich media that favor ed overall higher growth rates, lending credence to the idea that agmatine may comp ete with the structurally-related amino acid, arginine, either for uptake or charging of tR NAs. Specifically, inhibition was alleviated when the wild-type and aguB strains were grown in a trypt one yeast extract medium or BHI, both of which are abundant in peptides and carbohydrates (Figur es 6-18 to 6-21). Interestingly, in some cases, low AgD activit y seems to correlate with alleviation of agmatine inhibition: AgD is constitutiv ely expressed at low levels in the aguR strain, and AgD activity is significantly repressed when S. mutans is grown in rich media like TY and BHI. However, it is important to note that the aguB strain possesses no AgD activity and yet is the most sensitive strain to growth in agmatine. The most likely explanation is that, while some AgD activity is required for degradation of a toxic compound, too much AgD activity might result in over-production of ammonia. Alternatively, increased activity of the agmatine:putresc ine antiporter could cause agma tine to be brought into the cell at higher rates than it can be me tabolized, resulting in toxic accumulation of agmatine inside the cell. A se ries of checks and balances may be required for expressing
125 enough AgD activity to detoxify agmatine and maintain pH, without alkalinizing the extracellular environment and compro mising the competitive advantage that S. mutans has over other oral bacteria at low pH. If agmatine inhibits growth by competi ng with amino acid tr ansporters, then it would make sense that provision of an abunda nt amino acid source in peptide-form, via growth in TY or BHI media, may allow th e organisms to bypass competitive inhibition for transport. It is also noteworthy that agmatine inhibition was most severe when the strains were grown in TV medium supplem ented with agmatine and the nonrepressing sugar, galactose (incidentally, conditions that also result in the highest levels of AgD activity) (Figure 6-13). Howeve r, the doubling time of the aguB strain was similar to that of UA159 when the cells we re grown in TV medium containing agmatine and the repressing sugar, glucose, (Figure 6-18). In B. subtilis and P. aeruginosa , certain amino acid and peptide transporters are negatively regulated by CCR (Moreno et al. , 2001; Nishijyo et al. , 1998). Repression of the transporte rs during growth in glucose could reduce agmatine uptake and allow growth of the aguB strain. Components of a putative oligopeptide ABC (A TP-b inding c assette) transport system are located approximately1.5-kb upstream of aguR on the S. mutans chromosome (Figure 6-22). The system is composed of a substrate binding protein, encoded by oppA , two permease proteins, encoded by oppB and oppC , and two ATP-binding proteins, encoded by oppD and oppF . A conserved hypothetical ORF encoding a putative nitroreductase is transcribed immediately dow nstream of the ABC tr ansporter, separating it from the agu operon. In other bacteria, ABC trans port systems move a wide variety of substrates across cellular membranes, ranging from import of metabolites such as sugars,
126 peptides, amino acids, polyamines, and meta l ions, and export of waste products, cell membrane components and toxins involved in virulence (Dassa and Bouige, 2001; Fichant et al. , 2006; Higgins, 1995). To determine if the ABC transporter located upstream of the agu operon was involved in transport of agmatine, and to begin to explore the relationship between agmatine transport and growth inhibition, a polar mutation was constructed in oppA and a non-polar mutation was constructed in the agmatine:putrescine antiporter ( aguD ). The strains were grown in the presence and absence of 20 mM agmatine to assess growth inhibition (F igure 6-23). Mutation of the transporters did not alleviate growth inhibition, especially during growth in TV medium, suggesting that (i) either agmatine inhibition occurs independently of transport into the cell, or (ii) other transporters are respons ible for bringing agmatine into the cell. Furthermore, Real Time RT-PCR analysis of oppA , oppB , nitroreductase and aguA suggests that AguR does not regulate expres sion of this ABC transport system or the putative nitroreductase gene located between the ABC transporter and the agu operon (Figure 6-24). Importantly, the finding that agma tine inhibits the growth of S. mutans reveals a novel type of antagonistic interaction that ma y be displayed by competing oral biofilm organisms. Specifically, many bacteria re spond to acidification by inducing arginine decarboxylase (ADC) to produce agmatine, which is pumped out of the cell and alkalinizes the environment, enhancing the growth of acid-sensitive organisms (Hayes and Hyatt, 1974; Kanapka and Kleinberg, 1983; Lin et al. , 1995). In secreting agmatine, the organisms are releasing a com pound that inhibits the growth of S. mutans , a primary
127 generator of acids in oral biofilms. In response, S. mutans induces the AgDS, deriving ATP and ammonia from an inhibito ry compound (Figure 6-25). Sources of agmatine in the oral cavity. The exact source of agmatine in the oral cavity is unclear; however, agma tine has been found in low levels in virtually all human, animal and plant cells (Raasch et al. , 1995). As a result, agmatine is found in a wide variety of foods, including fish, grains, plants , fruits, meats, and fermented foods such as sauerkraut, cheese and beer. As described in Chapter 5, Wolinella , Prevotella , Bacteroides , Desulfovibrio , and Neisseria species possess the ADC enzyme and may be capable of decarboxylating arginine into agma tine, which could then be taken up via the S. mutans agmatine::putrescine antiporter. We we re unable to generate evidence of the ADC enzyme in Porphyromonas gingivalis , oral stre ptococci, Actinobacillus actinomycetemcomitans , Actinomyces naeslundii , Fusobacterium nucleatum , Treponema denticola , and Lactobacillus species. To estimate the amount of agmatine in human plaque and saliva, HPLC was used to analyze derivatized pooled human oral sa mples obtained from healthy laboratory volunteers. Approximately 750 nmol of agmatine (mg of protein)-1 was detected in pooled human plaque, and approximately 200 nmol of agmatine (mg of protein)-1 was detected in pooled human saliva. Collectively, these results demonstrate that agmatine is present in physiologically si gnificant amounts in dental plaq ue, and tentatively identify species that could contribute to the total agmatine pools. These observations support our hypothesis that the AgDS may be invol ved in agmatine detoxification by S. mutans , while concomitantly generating ATP and ammo nia. Additional studies to explore
128 agmatine production and utilization by oral micro-organisms and biofilm communities are underway. Summary To initiate a study of agu gene regulation, the aguB transcription initiation site was identified by primer extension and a putative 70-like promoter was mapped 5 to aguB . Analysis of the genome database revealed an open reading frame (SMU.261c) encoding a putative transcriptional regulator located 239 bases upstream of aguB . Inactivation of SMU.261c decreased AgD activity by 7-fold and eliminated agmatine induction. AgD was also found to be induced by certain environmental stresses, including low pH and heat, implying that the AgDS may also be a part of a general stress response pathway of this organism. Interestingly, an AgDS-defic ient strain was unable to grow in the presence of 20 mM agmatine, suggesting that the AgDS converts a growth-inhibitory substance into products that can enhance acid tolerance and contribute to the competitive fitness of the organism at low pH. The results presented in this chap ter demonstrate that the AgDS of S. mutans is functional and under tight genetic control. The physiological role of the AgDS in S. mutans is complex, conveying bioenergetic advantages through enhancement of pH and generation of ATP, as well as detoxifica tion of agmatine produ ced by acid-sensitive organisms in response to acidification by S. mutans . Agmatine catabolism may thereby increase the competitive fitness of S. mutans , contributing in major ways to the persistence and virulenc e of this organism.
129 Figure 6-1. Predicted structure of the S. mutans AguR protein. COOH Cell Membrane 318 aa HTH_LuxR Cleavable N-terminal signal sequence Trans memb. 1 HTH aguB D A C 249 b p
130 Figure 6-2. AgD enzyme activity of S. mutans UA159 and aguR , in response to increasing concentrations (mM) of agma tine. Results shown are the average and standard deviations (error bars) of a minimum of 9 separate cultures for each strain and condition. Activ ity is expressed as nmol Ncarbamoylputrescine produced/min/mg protein. One-way ANOVAs and pairwise Student t-tests were used to iden tify significant differences (p < 0.05) in AgD activity. Similar letters indicate no significant difference in activity. 0 50 100 150 200 250 [Agmatine]: 0 3 10 20 UA159 aguR a ac a a b bd d AgD Activity U (mg protein)-1
131 Figure 6-3. The aguRB intergenic sequence and primer extension analysis of S. mutans aguB and aguR. Arrow-heads indicate initiation site of aguB and aguR transcription at G and A residues, respec tively. The transcriptional start sites are marked with round arrowheads in the sequence, and the ribosome binding sites are shown in bold. The cre consensus sequences identified at and relative to the aguB transcriptional start site are underlined, with bases matching the consensus shown in bold. TCA ATT TTC ATG GAA GAA CCC TCC TTT AAT TTC TCT TTA ATA TCT ATA GTA ATT ATA TCA TTC AAA AGT TAA AA G TA C CTT CTT G GG AGG A AA TTA AAG AGA AAT TAT AG A TAT CAT TAA TAT AGT AAG TTT TAA ACT TTT AT T GAA AA G GCT TT G T AA AAA AGG CCT AAA CCT TGG GTT TAT TTT TTA AAA ACG GTT ATT TGA AAA TAA CTT TTC CGA AAC ATT TTT TCC GGA TTT GGA ACC CAA ATA AAA AAT TTT TGC CAA TAT TTC ATA AGG TTT AGT TAG CTA AAC CTT TGT T TG TAA TCG TTT ACA GCT CAA AGT TTA TAG TAA ATA AAG TCC AAA TCA ATC GAT TTG GAA ACA AAC ATT AGC AAA TGT CGA CGA GTT TCA AAT ATC ATT ACT GCC CTT TAA AAA AA T ATG AT G GAA AT G AAA AAA ATA G AG GAG G CT TCT ATG ATG AAA TGA CGG GAA ATT TTT TTA TAC TAC CTT TAC TTT TTT TAT CTC CTC CGA AGA TAC TAC TTT A C G T aguR A C G TaguB -10 -35 -35 -10 aguR aguB -137 cre -48 cre TIS TIS
132 Figure 6-4. Construction of paguB reporter gene fusions and CA T specific activity in wildtype UA159 and aguR . A) CAT activity of the full length aguB promoter. B) CAT activity of the aguB promoter derivatives. Activity is expressed as nmol/min/mg protein. Results shown are the average and standard deviations (error bars) of a minimum of 9 separate cultures for each strain and condition. One-way ANOVAs and pair-wise Student t-tests were used to identify significant differences (p < 0.05) in CA T activity. Similar letters indicate no significant difference in activity. cat RBS cre -48 cat RBS UA159: 1.74 0.2 aguR : 0.00 0.0 UA159: 0.00 0.0 aguR : 0.00 0.0 cat RBS cre cre 137 0 2 4 6 8 10 12 Glu WT GluAg WT Gal WT GalAg WT GalAg aguR ND ND ND A B a b CAT Activity U (mg protein)-1
133 Figure 6-5. AgD enzyme activity of S. mutans UA159, and of otherwise-isogenic ccpA and ccpB strains, in response to growth in the presence of the catabolite repressing sugar, glucose, or the nonre pressing sugar, galactose. Results shown are the average and standard deviat ions (error bars) of a minimum of 9 separate cultures for each strain and condition. Activit y is expressed as nmol N-carbamoylputrescine produced/min/mg protein; ND = no activity detected. One-way ANOVAs and pair-wise Student t-tests were used to identify significant differences (p < 0.05) in Ag D activity. Similar letters indicate no significant difference in activity. 0 50 100 150 200 250 UA159 ccpA ccpB Glu Glu Ag Gal Gal Ag ND a b b c c c d e de AgD Activity U (mg protein)-1
134 Figure 6-6. AgD enzyme activity in relation to growth domain. Results are the average and standard deviations (error bars) of 9 separate cultures for each condition. Activity is expressed as nmol Ncarbamoylputrescine produced/min/mg protein. 7.0 6.6 6.2 5.7 5.6 5.5 TV Gal TV Gal Ag Growth in TV Gal Ag 400 Time (hours) AgD Activity OD 600 0 50 100 150 200 250 300 350 0 3 6912 15 18 21 24 0.0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 pH
135 Figure 6-7. AgD activity in mid-exponential phase or stationary phase cultures grown in pH 7-buffered medium. Results shown are the average and standard deviations (error bars) of 9 separate cultures for each st rain and condition. Activity is expressed as nmol Ncarbamoylputrescine produced/min/mg protein. One-way ANOVAs and pair-wis e Student t-tests were used to identify significant differences (p < 0.05) in AgD activity. Similar letters indicate no significant difference in activity. 0 50 100 150 200 250 300 0.5 1.2 â€“ â€“ + + OD600: 0.5 1.2 Agmatine: ND a b AgD Activity U (mg protein)-1
136 Figure 6-8. AgD activity in continuous cultures of S. mutans UA159 maintained at pH 5 or pH 7. Results shown are the average a nd standard deviations (error bars) of a minimum of 9 separate cultures for each strain and condition. Activity is expressed as nmol N-carbamoylputre scine produced/min/mg protein. Oneway ANOVAs and pair-wise Student t-test s were used to identify significant differences (p < 0.05) in AgD activity. Similar letters indicate no significant difference in activity. 0 2 4 6 8 10 12 14 16 18 20 22 24 pH: 7 7 5 5 Agmatine: â€“ + â€“ + a b b c AgD Activity U (mg protein)-1
137 Figure 6-9. Real-Time RT-PCR of aguA expression (copies/ l) in continuous cultures of S. mutans UA159 maintained at pH 5 or pH 7. Results shown are the average and standard deviations (error bars) of a minimum of 9 separate cultures for each strain and condition. One-way ANO VAs and pair-wise Student t-tests were used to identify significant diff erences (p < 0.05) in gene expression. Similar letters indicate no signi ficant difference in expression. 0 50000 100000 150000 200000 250000 300000 350000 pH: 7 7 5 5 Agmatine: â€“ + â€“ + a b b cCopy Number
138 Figure 6-10. Ag D activity in response to growth (to OD600 = 0.6) at 37 C or 42 C in TV medium supplemented with 25 mM glucose and 10 mM agmatine. Results shown are the average and standard deviat ions (error bars) of a minimum of 9 separate cultures for each strain and condition. Activity is expressed as nmol N-carbamoylputrescine produced/min /mg protein. One-way ANOVAs and pair-wise Student t-tests were used to identify significant differences (p < 0.05) in AgD activity. Similar letters indicate no significant difference in activity. 0 50 100 150 200 250 300 37 42 a bAgD Activity U (mg protein)-1
139 Figure 6-11. AgD enzyme activity of S. mutans UA159 grown in TV medium containing 25 mM galactose and 10 mM agmatine. Aerobic: 10 mL bacteria were grown in 250 mL flasks at 37 C, with shaking. Anaerobic: 10 mL cultures were grown in 14 mL screw-cap tubes at 37 C in with a GasPak. Activity is expressed as nmol N-carbamoylputresc ine produced/min/mg protein. Results shown are the average and standard deviat ions (error bars) of a minimum of 9 separate cultures for eac h strain and condition. Aerobic 0 50 100 150 200 250 Anaerobic AgD Activity U (mg protein)-1
140 Figure 6-12. AgD enzyme activity of S. mutans UA159 after incubation with 400 mM NaCl for 30 and 60 minutes. Non-salt shocked cultures are indicated by â€œCtrlâ€. Activity is expressed as nmol N-carbamoylputrescine produced/min/mg protein. Results show n are the average and standard deviations (error bars) of a minimum of 9 separate cultures for each strain and condition. One-way ANOVAs and pair-wise Student t-tests were used to identify significant differences (p < 0.05) in AgD activity. Similar letters indicate no significant difference in activity. 0 100 200 300 400 30 minCtrl 30 60 minCtrl 60 a b ab a AgD Activity U (mg protein)-1
141 Figure 6-13. Growth of S. mutans UA159, aguB , and aguC strains in TV medium containing 25 mM galactose and either 0 mM or 20 mM agmatine. Optical density at 600 nm was determined every 30 minutes for 50 hours using a Bioscreen C. Each point represents th e average of three separate cultures. Standard deviations for each point we re < 0.02 for cultures grown in 0 mM agmatine, and < 0.06 for cultures grown in 20 mM agmatine. 0.000 0.100 0.200 0.300 0.400 0.500 0.600 0 5 101520253035404550 Time (hours) OD 600 WT 0 mM Ag aguB 0 mM Ag aguC 0 mM Ag WT 20 mM Ag aguB 20 mM Ag aguC 20 mM Ag Detoxification ATP & NH3 No Detox, ATP or NH3 Detoxification No ATP; Less NH3
142 Figure 6-14. Growth of S. mutans wild-type, aguB , aguC and aguR strains in 0, 3, 10 and 20 mM agmatine. The y-axis refers to cell density measured at OD 600. Each point represents the average of three se parate cultures. St andard deviations for each point were < 0.05. UA159 0.000 0.100 0.200 0.300 0.400 0.500 0.600 0.700 0.800 0 4 6 8 10 12 14 16202530 aguB 0.000 0.100 0.200 0.300 0.400 0.500 0.600 0.700 0.800 046810 12 14 16 20 2530 Gal + 0 mM Ag Gal + 3mM Ag Gal + 10mM Ag Gal + 20mM Ag Time (hours) aguC 0.000 0.100 0.200 0.300 0.400 0.500 0.600 0.700 0.800 0 4 6 8 10 12 14 16202530 0.000 0.100 0.200 0.300 0.400 0.500 0.600 0.700 046810 12 14 16 202530 aguR Time (hours) OD 600 OD 600
143 Figure 6-15. Growth of an aguRB double mutant strain in TV medium containing 25 mM galactose, supplemented with 0 mM (top) or 20 mM (bottom) agmatine. Each point represents the average of three se parate cultures. St andard deviations for each point were < 0.05. WT aguB aguR aguRBOD 600 0.000 0.100 0.200 0.300 0.400 0.500 0.600 0.700 0.800 0.900 046810121416202530Time (hours) 046810121416202530OD 600 0.000 0.100 0.200 0.300 0.400 0.500 0.600 0.700 0.800 0.900
144 Figure 6-16. Growth of S. mutans and S. rattus strains in TV medium containing 25 mM galactose and 0 mM (open symbols) or 20 mM (closed symbols) agmatine. Standard deviations are as reported in Figure 6-13. 0.000 0.200 0.400 0.600 0.800 1.000 1.200 1.400 0 2 4 6 8 10 12 14 16 18 20 S. mutans UA159 (-) S. mutans GS-5 (-) S. rattus FA-1 (-) S. rattus BHT (-) S. mutans UA159 (+) S. mutans GS-5 (+) S. rattus FA-1 (+) S. rattus BHT (+) OD 600
145 Figure 6-17. Growth of select ed oral streptococ ci in TV medium containing 25 mM galactose and 0 mM (open symbols) or 20 mM (closed symbols) agmatine. Standard deviations are as reported in Figure 6-13. 0.000 0.050 0.100 0.150 0.200 0.250 0.300 0.350 0 2 4 6 8 101214161820 S. salivarius 57.I (-) S. gordonii DL-1 (-) S. oralis SK92 (-) S. sanguis 10556 (-) S. salivarius 57.I (+) S. gordonii DL-1 (+) S. oralis SK92 (+) S. sanguis 10556 (+) OD 600
146 Figure 6-18. Growth of S. mutans UA159 and aguB in the presence and absence of agmatine in TV medium containing 25 mM glucose. Standard deviations are as reported in Figure 6-13. UA159 aguB UA159 + Ag aguB + Ag Time (hours) OD 600 TV Glucose 0.000 0.200 0.400 0.600 0.800 1.000 1.200 0 51015 20
147 Figure 6-19. Growth of S. mutans UA159 and aguB in the presence and absence of agmatine in TY medium containing 25 mM galactose. Standard deviations are as reported in Figure 6-13. UA159 aguB UA159 + Ag aguB + Ag Time (hours) OD 600 TY Galactose 0.000 0.200 0.400 0.600 0.800 1.000 1.200 0 51015 20
148 Figure 6-20. Growth of S. mutans UA159 and aguB in the presence and absence of agmatine in TY medium containing 25 mM glucose. Standard deviations are as reported in Figure 6-13. UA159 aguB UA159 + Ag aguB + Ag Time (hours) OD 600 TY Glucose 0.000 0.200 0.400 0.600 0.800 1.000 1.200 0 51015 20
149 Figure 6-21. Growth of S. mutans UA159 and aguB in the presence and absence of agmatine in BHI medium. Standard devi ations are as reported in Figure 6-13. UA159 aguB UA159 + Ag aguB + Ag Time (hours) OD 600 BHI 0.000 0.200 0.400 0.600 0.800 1.000 1.200 0 51015 20
150 Figure 6-22. Map of the putativ e oligopeptide ABC transport system located upstream of the agu operon in S. mutans UA159. SMU.260 encodes a conserved hypothetical protein with homology to n itroreductases in other bacteria.
151 Figure 6-23. Growth of the oppA and aguD mutants in 0 mM (open symbols) or 20 mM (closed symbols) agmatine. A) Growth in TV containing 25 mM glucose. B) Growth in TY containing 25 mM glucos e. C) Growth in TV containing 25 mM galactose. D) Growth in TY c ontaining 25 mM galactose. Standard deviations are as reported in Figure 613. The x-axis refers to cell density measured at OD 600. Time (hours) A 0 0.1 0.2 0.3 0.4 0.5 0.6 0 5 10 15 20 B 0 0.2 0.4 0.6 0.8 1.0 1.2 0 5 10 15 20 C D 0 0.2 0.4 0.6 0.8 1.0 oppA -Ag aguD -Ag oppA +Ag aguD +Ag 0.1 0.2 0.3 0.4 0.5 0.6 0 5 10 15 20 0 5 10 15 20 OD 600 OD 600
152 Figure 6-24. Real Time RT-PCR (copies/ l) of the oppA , oppB , nitroreductase and aguA genes in A) the S. mutans wild-type and B) aguR background. C) Fold change in gene expression in the absence of aguR . RNA was isolated from cultures grown to mid-exponential phase in TV medium containing 25 mM glucose or galactose, with or without 10 mM ag matine. Results shown are the average and standard deviations (error bars) of a minimum of 5 replicates. One-way ANOVAs and pair-wise Student t-tests were used to identify significant differences (p < 0.05) in expression of individual genes. Similar letters indicate no significant difference in activity. 1 10 100 1000 10000 100000 1000000 oppA oppB nitro aguA oppA oppB nitro aguA -40 -20 0 20 40 60 80 100 120 oppA oppB nitro aguA TV Glu TV Glu Ag TV Gal TV Gal Ag a a a a b c d a b a a a b c A. B. C. Co py Number
153 Figure 6-25. Proposed role of the AgDS in virulence. As S. mutans lowers the pH of dental plaque, acid-sensitive bacteria in the oral biofilm induce arginine decarboxylase to produce agmatine, which inhibits the growth of S. mutans . At low pH and in the presence of agmatine, S. mutans induces the AgDS and converts the inhibitory substance into putrescine, ATP and ammonia, thereby enhancing acid tolerance and contributing to the fitness of this organism at low pH. The ATP generated from agmatine catabolism can be used to power the proton-pumping F1F0-ATPase, while ammonia alkalinizes the cytoplasm, increasing pH. Maintenance of a relatively alkaline cytoplasm compared to the extracellular environment allows S. mutans to continue growth and acid production at low pH, eventually emergi ng as a sufficiently large proportion of the biofilm to effect subs tantial tooth demineralization. S . mutans H+ H+ H+ Arginine + H+ Agmatine H+ Agmatine Putrescine H+ NH 3 NH 3 ATP ADP H+ H+ H+ H+ H+ H+ H+ H+ ADC+ Bacteria
154 CHAPTER 7 SUMMARY AND FUTURE DIRECTIONS Arginine Deiminase System The oral cavity is a complex ecosystem that undergoes frequent changes in environmental factors known to profoundly a ffect bacterial gene expression, such as oxygen tension, pH and nutrient source (Quivey et al. , 2001). In order to persist in this habitat, oral bacteria must possess sophisticated strategies for coping with environmental stress. The role of acid tolerance in oral microbial ecology has been extensively studied, as it is well known that organisms capable of growth and acidogenesis at low pH play an integral role in the development of dental caries (Loesche, 1986). Conversely, organisms capable of alkalinizing dental plaque are thought to play important roles in caries prevention (Burne and Marquis, 2000; Clancy et al. , 2000; Margolis et al. , 1988; Van Wuyckhuyse et al. , 1995; Wijeyeweera and Kleinbe rg, 1989b). Thus, a detailed understanding of the ammonia-generating mechan isms used by oral bacteria is required for the development of effective anti-caries treatments. The ADS has long been recognized for its role in ammonia and energy generation by acid-sensitive oral bacteria, particularly when the environment is too acidic for efficient glycolysis of av ailable carbohydrates (Curran et al. , 1995; Curran et al. , 1998; Marquis et al. , 1987). Much of the previous research on the ADS in oral streptococci has focused on the biochemical properties of ADS enzymes (Cunin et al. , 1986; Hiraoka et al. , 1986; Poolman et al. , 1987), and the roles of this sy stem in protection against acid stress and nutrient depletion (Cas iano-Colon and Marquis, 1988; Curran et al. , 1995;
155 Marquis et al. , 1992). Collectively, these studies indi cate that the ADS helps stabilize healthy dental plaque biofilms (Marsh, 1991; Marsh and Bradshaw, 1997). Furthermore, it has been widely speculated that certain oral b acteria, such as S. mutans , might be made less cariogenic if they are e ngineered to express ADS genes. An arginolytic strain of S. mutans might retain the ability to compete with other oral ba cteria, while contributing to a healthier plaque biofilm by neutralizing the acidic products of glycolysis via alkali production. However, substantial gaps in our understanding of the molecular genetics and regulation of the ADS in oral streptoco cci have undermined our ability to engineer such arginolytic strains. The close taxonomic relationship between S. mutans and the ADS-utilizing bacterium, S. rattus , has attracted the atten tion of research seeki ng to establish the ADS in S. mutans . In an attempt to increase our unders tanding of the molecular genetics of the ADS in oral streptococci, a primary goal of this dissertation was to identify the genes encoding the ADS in S. rattus FA-1 and examine the factor s involved in regulation of arc expression. As demonstrated in Chapter 3, the S. rattus ADS has several attractive properties that would ma ke it suitable for use in construction of a S. mutans replacement therapy strain. Notably, the S. rattus arc operon is markedly less sensitive to regulation by oxygen tension, arginine induction a nd catabolite repression than the arc operons of other oral streptococci, such as S. gordonii and S. sanguis (Burne et al. , 1991; Dong et al. , 2004; Griswold et al. , 2004a). These properties would be particularly advantageous during growth in vivo because enhanced arginolysis w ould likely occur regardless of environmental fluctuations in oxygen, arginine and carbohydrate source.
156 Interestingly, the regulators responsible for induction of the S. rattus ADS by arginine and oxygen were identified in the S. mutans UA159 genome. Experiments described in Chapter 4 showed that at least one of these regulators, AdiR, has retained the ability to activate parcA expression in response to arginine in a manner consistent with the AdiR protein of S. rattus . This observation is enc ouraging, as it underlines the practicality of r econstituting the S. rattus arc operon in S. mutans and increases the liklihood that the arc genes would be properly regulated. In the future, an arginolytic strain of S. mutans could be implanted into the dental plaque of an animal model or added to an in vitro multi-species biofilm model to evalua te the effect of alkali production on cariogenicity and biofilm communi ty stability, respectively. Previously, the Burne laboratory conduc ted a similar experiment in which the urease operon of S. salivarius 57.I was integrated into the chromosome of S. mutans GS5 (Clancy et al. , 2000). In vitro pH drop analyses of the wild-type and urease-positive S. mutans demonstrated that production of ammonia from urease greatly offsets the pH drop normally observed after a carbohydrate cha llenge. Thus, enhancing the ammoniagenerating capability of cario genic plaque bacteria will likely reduce the induction and progression of dental caries. Agmatine Deiminase System Examination of the S. mutans genomic sequence uncovered a cluster of ORFs with significant homology to the arc genes of S. rattus FA-1. As it has been well-established that S. rattus is the only ADS-utilizing member of the mutans streptococci, the discovery of an arc -like gene cluster in S. mutans was especially intriguing. Further characterization of the ORFs revealed a novel pathway for ammonia production that has not been identified in other oral bacteria. In this pathway, ammonia is generated from
157 agmatine in a manner highly analogous to the pathway for ammonia production from arginine by the ADS. The identification of an ammonia-generating system in S. mutans is significant, given the contri bution of acid tolerance to virulence. Therefore, the second major goal of this work involved character izing the genes encoding enzymes of the AgDS in S. mutans and identifying factors involved in regulation of this system. As demonstrated in Chapter 5, the opt imum pH for ammonia production from agmatine is similar to the minimum pH obs erved in dental plaque after carbohydrate consumption (Stephan, 1940). It is possible th at agmatine catabolism imparts a selective advantage to S. mutans during growth at low pH through generation of ATP, which could be used for growth or to move protons out of the cell, and alkalinization of the cytoplasm by ammonia, which would reduce the investme nt of ATP in proton extrusion. Under such conditions, S. mutans would gain a competitive advantage over less acid-tolerant species in oral biofilms, particularly since the system is expressed at a low level and would not profoundly effect environmental alkalinization. Further studies will be required to conclusively demonstrate that the AgDS conveys a competitive advantage to S. mutans at low pH. Specificall y, the wild-type or the aguB strain could be integrated into a 10-species in vitro biofilm model, with ADC-produc ing bacteria. Physiologically significant amounts of agmatine could be adde d to the growth media to determine if agmatine enhances the ability of the wild-typ e strain to dominate the biofilm community at low pH. The AgDS of S. mutans is tightly regulated by carbon catabolite repression, as well as induction by agmatine, low pH and heat, as described in Chapter 6. Given the potential role of the AgDS in acid tolerance, future studies should focus on the effects of
158 low pH and attempt to unravel the cis and trans -acting factors responsible for induction of the agu operon by acid. For example, th e lysine decarboxylase system of E. coli , encoded by the cadBA genes, is similar to the AgDS in that it is activated by substrate (lysine) and by low pH. Both facets are medi ated by the regulatory protein CadC. The mechanisms used by CadC to activate the genes in response to pH is still unclear, but the general consensus is that the protein undergoe s a conformational change at low pH that allows it to bind to and activate the cad prom oter (Watson et al., 1992). Therefore, future work could attempt to determine if AguR me diates induction by low pH and agmatine in a similar manner as CadC. One of the most interesting conclusions to emerge from Chapter 6 is the potential antagonistic interaction revealed among S. mutans and competing oral biofilm organisms. Many oral bacteria respond to acidificati on by decarboxylating arginine into agmatine, which is pumped out of the cell and serves two purposes: (1) alka linization of the environment, and (2) growth inhibition of S. mutans , the primary producer of acid (Hayes and Hyatt, 1974; Kanapka and Kleinberg, 1983; Lin et al., 1995). In response, S. mutans induces the AgDS, deriving ATP and ammoni a from an inhibitory compound (Figure 624). To further explore the interaction between S. mutans and arginine decarboxylase (ADC)-utilizing bacteria, it would be interesting to determine if S. mutans AgD enzyme activity is up-regulated in response to co-cultiv ation with oral bacter ia known to utilize ADC, with arginine included or excluded from the growth media. In addition, it would be interesting to implant S. mutans wild-type and aguB strains in a rat model to evaluate the contribution of the Ag DS to cariogenicity.
159 Role of Ammonia Generation in Mutans Streptococci A comparison of the ammonia-generating pa thways used by mutans streptococci reveals that the S. mutans AgDS is expressed at a much lower rate than the S. rattus ADS. If agmatine is present in much lower concen trations in the mouth than arginine, which seems likely, then a highly active AgDS sy stem may not be needed to catabolize the amount of substrate that is typically available in vivo . Furthermore, if the AgD enzymes possess a high affinity for agmatine, then rapid turnover (e.g. high enzyme activity) would not necessarily be required for efficien t agmatine catabolism. Another compatible explanation is that low levels of ag matine catabolism would produce sufficient intracellular ammonia to neutra lize the cytoplasm, but would not have a major impact on the surrounding pH. By catabolising agmatine at a relatively slow rate, S. mutans could reap the benefits of intracellular ammonia production without concur rently increasing the extracellular pH and providing acid-sensitive competitors with more favorable growth conditions. In contrast, ammonia generation from ar ginine by the ADS of S. rattus has been shown to promote alkalinization of dental plaque following a carbohydrate challenge, and alkali generation via the ADS is generally thought to contribute to maintenance of healthy biofilm commun ities (Wijeyeweera and Kleinberg, 1989b). The presence of analogous ammonia-gene rating mechanisms in two very closelyrelated bacteria â€“ S. rattus , a non-cariogenic bacterium, and S. mutans , the etiological agent of dental caries â€“ has presented a unique opportunity to examine the role of ammonia-generating mechanisms in the viru lence and acid:base physiology of mutans streptococci. The results desc ribed in this dissertation have revealed many similarities and differences in the regulation of ammoniagenerating systems in mu tans streptococci. Notably, expression of the S. rattus ADS and the S. mutans AgDS are both modulated by
160 environmental fluctuations in substrate le vels and carbohydrate source. However, these variables play a more significan t role in regulation of the S. mutans AgDS, while S. rattus is capable of efficient arginolysis during gr owth in a wide vari ety of environmental conditions. A thorough examination of ADS and AgDS genetic organization and regulation has suggested diverse roles for ammonia production in the physiology of or al streptococci. Ammonia production appears to enhance the fitness of both cariogenic and noncariogenic mutans streptococci . However, while the ADS appears to function primarily as an acid-tolerance mechanism for healthy or al flora, we have demonstrated that the AgDS converts agmatine into products that ma y enhance acid tolerance and contribute to the pathogenicity of S. mutans at low pH. Thus, it can be argued that S. rattus produces ammonia via the ADS to promote health, while S. mutans produces ammonia via the AgDS to promote disease.
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178 BIOGRAPHICAL SKETCH The author was born in Charlotte, Nort h Carolina, on February 3, 1979, to Patrick V. and Darlene A. McCarthy. She was married on June 23, 2001, to Marcus W. Griswold. Ann attended the University of Maryland at College Park from 1997 to 2001. As an undergraduate, she was awarded a paid internship to conduct food microbiology research at the U.S. Food and Drug Ad ministration in Washington, DC, from 1998 to 2001. She graduated from the University of Maryland in May 2001 with a Bachelor of Science degree in microbiology. In August 2001 she was awarded an Alumni Fellowship from the University of Florida and began gra duate studies in the Co llege of Medicineâ€™s Interdisciplinary Program in Biomedical Scie nces. From March 2002 to the present, her graduate research in the Immunology and Microbiology advanced concentration was supervised by Robert A. Burne, Ph.D. and was supported by Public Health Service grant DE10362 from the National Institute of Dental and Craniofacial Research. In 2005, she was awarded first place in the Immunology and Microbiology di vision of the UF Medical Guild Graduate Student Research Competition, and third place overall. Also in 2005, she was awarded second place for an oral presenta tion in the Graduate Student/Post-Doctoral division of the UF College of Den tistry Research Day Competition. The author is a member of the Am erican Society for Microbiology, the International Association for Dental Res earch, the American Association for the Advancement of Science and the National A ssociation of Science Writers, Inc..