Analysis of the Transcriptional Regulation of the Tpr Protease Gene of Porphyromonas Gingivalis

ANALYSIS OF THE TRANSCRIPTIONAL REGULATION OF THE TPR PROTEASE GENE OF PORPHYROMONAS GINGIVALIS

BIQING LU

B. Med/B.Sc, Hubei Medical University, 1988 M.Sc, The University of British Columbia, 1993

A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENT FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

THE FACULTY OF GRADUATE STUDIES (Department of Microbiology & Immunology)

We accept this thesis as conforming to the required standard

THE UNIVERSITY OF EfRITISH COLUMBIA

November, 1997

© Biqing Lu, 1997 In presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department or by his or her representatives. It is understood that copying or publication of this thesis for -financial gain shall not be allowed without my written permission.

Department of MiUT&n&ter^ (^MUMQI^JJ

The University of British Columbia Vancouver, Canada

Date dec £ , mi

DE-6 (2/88) ABSTRACT

The expression of Porphyromonas gingivalis W83 tpr protease gene is regulated at the transcription level by the nutritional environment. To investigate the regulation of tpr expression in P. gingivalis, we determined the sequence of the upstream region of tpr and by primer extension studies we determined that the transcription start sites were at two A residues 215 nucleotides upstream of the coding region. Computer analysis of the immediate upstream region did not find any consensus sequence to the

-35 and -10 region of E. coli promoters. A tprr.lacZ reporter gene construct was introduced into P. gingivalis, the reporter gene expression is under the control of tpr promoter. Deletion mutations in the tpr upstream regions confirmed the promoter region to be the same as that determined by primer extension studies. Three identical direct repeats of 17 bp were identified in the upstream region of tpr. Deletion analysis of these repeats suggested they were involved in tpr regulation. Tpr expression was repressed by the addition of nutrients, such as brain heart infusion (BHI), trypticase peptone, bovine serum albumin (BSA) and gelatin. Single amino acids had little effect. BHI fractions of low molecular weight rich in proline, alanine and phenylalanine repressed tpr expression the most. The di-peptide phenylalanyl-phenylalanine repressed tpr expression. Heat shock, pH change and hemin starvation had little effect on tpr expression. The tpr::lacZ reporter constructs cloned in E. coli also expressed (3-galactosidase activity under the control of the tpr upstream region.

However, this expression was not regulated in the same way as in P. gingivalis W83.

Primer extension analysis revealed that E. coli initiated transcription after an AT rich region of DNA but did not recognize P. gingivalis native tpr promoter. This was confirmed by deletion mutagenesis analysis. TABLE OF CONTENTS

ABSTRACT ii LIST OF TABLES vi LIST OF FIGURES vii ABBREVIATIONS, NOMENCLATURE AND SYMBOLS ix ACKNOWLEDGMENTS xii I. INTRODUCTION 1 1. Microbiology of periodontal disease 1

2. Porphyromonas gingivalis 3

2.1. Nutrition requirements of P. gingivalis 3

2.2. Pathogenecity of P. gingivalis 4

3. Virulence factors of P. gingivalis 5 3.1. Adherence 5 3.1.1. Fimbriae 5 3.1.2. Hemagglutinin 6 3.1.3. Other surface binding molecules 7 3.2. Capsule 7 3.3. Lipopolysaccharide (LPS) 8 3.4. Vesicles 8 3.5. Proteases 9 3.5.1. Trypsin-like activity 9 3.5.2. Collagenase activity 13 3.5.3. Glycylprolyl proteolytic activity 13 3.5.4. The Tpr protease and Pz-peptidase activity 13

3.5.5. Potential roles of P. gingivalis proteases in pathogenesis 17

iii 3.6. Other enzymes and metabolic end products 18

4. Overview of molecular biology of P. gingivalis 19 4.1. Cloning of P. gingivalis genes 19 4.2. Genetic manipulation of P. gingivalis 22 5. Gene regulation 24 5.1. Transcription regulation in prokaryotes 24

5.2. Gene regulation in P. gingivalis 30 5.2.1. Effect of nutrition 30 5.2.2. Effect of hemin 31 5.2.3. Effect of environmental pH 33 5.2.4. Effect of temperature 33 6. The purpose of this study 33 II. MATERIALS AND METHODS 34 1. Bacterial strains and culture conditions 34 2. DNA manipulation methods 34 3. Construction of P. gingivalis W50 DNA library 38 4. Screening for proteolytic clones 39 5. PCR and primers 39 6. DNA sequencing 41 7. Primer extension analysis 43 8. RNA isolation 44 9. Northern blot 45

10. Construction of tprr.lacZ reporter vectors and selection of P. gingivalis transconjugants 45 11. Southern blot 48 12. Electrophoresis and gelatin zymography 49 13. Western immunoblot 50

iv 14. p-galactosidase assay 50 15. Stress conditions and hemin limitation 52 III. RESULTS 53 1. Identification of proteolytic clones 53 2. Analysis of proteolytic clone TZ18-1 53 3. The effect of nutrient limitation on tpr transcription 61

4. Analysis of tpr expression by a tpr::lacZ reporter construct 66 5. Primer extension and sequence analysis of the promoter region

of tpr. 70 6. Molecular analysis of tpr 5' region by deletion mutations 76 7. Growth factors that regulate tpr expression 81 8. Effect of other environmental conditions on tpr expression 85 IV. DISCUSSION 87 V. BIBLIOGRAPHY 97

V LIST OF TABLES

TABLE PAGE

1. Comparison of reported P. gingivalis genes encoding homologous Arg-X specific proteases 11 2. Comparison of active-site amino acid residues of various cysteine proteinases 16

3. Sequenced P. gingivalis genes and some features 20 4. Consensus nucleotide sequences for promoters recognized by RNA polymerase containing various sigma factors 27 5. Strains and plasmids 35 6. Primers for PCR and primer extension analysis 40 7. Southern blot analysis of SamHI digested chromosomal DNA

from P. gingivalis W83 transconjugants 73

vi LIST OF FIGURES

FIGURE PAGE 1. Periodontium in healthy (A) and diseased (B) state 2 2. Nucleotide sequence of the cloned tpr gene from

P. gingivalis W83 15 3. Schematic representation of the overall transcription cycle 29 4. PCR products and their cloning vectors 42

5. Construction of tpr.UacZ reporter shuttle vector pNTX-400 46 6. Construction of suicide shuttle vector pBYZ 47 7. Proteolytic activity of clone TZ18-1 and TZ18-2 on skim milk agar plate 54 8. Restriction analysis of plasmids pTZ18-1 and pTZ18-2 55 9. Partial restriction map of pTZ18-1 and pYS307 56 10. Southern blot analysis of pTZ18-1 with a tpr probe 57 11. Western immunoblot of TZ18-1 with antisera against

P. gingivalis W50 and W80 59 12. SDS-PAGE and gelatin zymogram of clone TZ18-1 60 13A. Northern blot analysis of fprmRNA from W83 grown in BHI, TYE, 0.5TYE and W83/PM grown in TYE 62 13B. Densitometry scan of the blot 62 14. Northern blot analysis of tpr mRNA in various nutritional conditions 63

15. Northern blot analysis of tpr mRNA expression in P. gingivalis W83 and W83/PM 64

16. Northern blot analysis of tpr mRNA expression in P. gingivalis ATCC33277 65

vii 17. tpr mRNA expression at different growth stage 67 18. |3-galactosidase activity in P. gingivalis W83/pNTX-400 lysates 68

19. Effect of growth nutrient on lacZ expression in P. gingivalis W83/pNTX-400 69 20. Possible homologous recombinations between pBYZ and P. gingivalis W83 chromosome 71

21. Southern blot analysis of chromosomal DNA from P. gingivalis W83 strains 72 22. f3-galactosidase expression in P. gingivalis transconjugants 74 23. Nucleotide sequence of the 5' region of fprgene from

P. gingivalis W83 75 24. Primer extension analysis of tpr transcription in

P. gingivalis W83 77

25. p-galactosidase activity in P. gingivalis and E. coli with various

tpr.-.lacL constructs 78 26. Primer extension analysis of tpr transcription in E. coliXL-1 /pTXZ19-400 80 27. Growth curve (A) and (3-galactosidase activity (B) of

P. gingivalis W83//acZ 82 28. Effect of individual amino acids on (3-galactosidase expression

i n P. gingivalis W83//acZ 83

29. Effect of peptides and some other chemicals on lacZ

expression in P. gingivalis W83//acZ 84 30. Effect of hemin limitation, heat shock and pH on p-galactosidase

expression in P. gingivalis\N83/lacl 86

viii ABBREVIATIONS, NOMENCLATURE AND SYMBOLS

2D-PAGE two-dimensional polyacrylamide gel electrophoresis Amp ampicillin APMA 4-aminophenylmercuric acetate ATCC American Type Culture Collection BApNA a-N-benzoyl-L-arginine p-nitroanilide BCIP 5-bromo-4-chloro-3-indolyl phosphate BHI brain heart infusion (3ME P-mercaptoethanol BPB black-pigmented Bacteroides species BSA bovine serum albumin DTT dithiothreitol EDTA ethylenediamine tetraacetate Eh reduction potential Em erythromycin G+C guanine + cytosine GApNA glycyl-L-arginine p-nitroanilide GCF gingival crevicular fluid Gm gentamycin h hours IL interleukine IPTG isopropyl-p\D-thiogalactoside IS insertional sequence Km kanamycin LB Luria-Bertani medium LPS lipopolysaccharide

ix min minutes NBT nitroblue tetrazolium NEM N-ethylmaleimide OD optical density ONPG o-nitrophenyl-p-D-galactopyranoside PAGE polyacrylamide gel electrophoresis PBS phosphate-buffered saline pCMB p-chloromercuribenzoate PCR polymerase chain reaction PMNL polymorphonuclear leukocyte PMSF phenylmethysulfonyl fluoride

Pz-peptide p-phenylazobenzyloxycarbonyl-L-prolyl-L-leucyl-glycyl-L-prolyl-D- arginine r resistant SAAPpNA N-succinyl-L-alanyl-L-alanyl-L-prolyl-L-phenylalanine p-nitroanilide SDS sodium dodecyl sulfate SOD superoxide dismutase SSC NaCI-Sodium citrate buffer (3 M NaCI, 0.3 M Sodium citrate, pH 7.0) TBS Tris-buffered saline Tc tetracycline TLCK N-a-p-tosyl-L-lysine chloromethyl ketone Tp trimethoprim TPCK N-tosyl-L-phenylalanine chloromethyl ketone tpr::lacZ tpr-lacZ transcriptional and translational fusion Tris Tris [hydroxymethyl] aminomethane TYE trypticase peptone-yeast Extract medium v/v volume/volume w/v weight/volume X-gal 5-bromo-4-chloro-3-indolyl (3-D-galactopyranoside

xi ACKNOWLEDGMENTS

I am deeply grateful to Dr. B.C. McBride for his understanding and guidance, and for his continued encouragement and support of this research project, and for the financial support of the University of British Columbia and Medical Research Council of Canada. My sincere thanks to Drs. R.E.W. Hancock, G. Weeks, and V.V.J. Uitto for serving on my committee and for their guidance and support. I wish to give my thanks to Dr. P. Hannam, C. Fenno, Y. Park, W.K Leung and A. Joe for sharing their knowledge and experience and their constant encouragement. I am thankful to S. Wan, G. Wong, and M. Tamura for sharing their experience and knowledge. My special thanks to my dear wife Wen Luo who provided great help for the completion of this thesis. I take this opportunity to express my gratitude to my family and friends for their constant encouragement and support.

Xll I. INTRODUCTION

1. Microbiology of periodontal disease The periodontium consists of gingiva, periodontal ligament, root cementum, and alveolar bone (Figure 1). Diseases that affect the periodontium are collectively called periodontal diseases. Various microorganisms can cause virtually all forms of inflammatory periodontal diseases which can be broadly grouped into gingivitis and periodontitis, and each can be further divided into subgroups according to disease activity and severity, age of onset, related systemic disorders and other factors. Gingivitis is inflammation of the gingiva, but it does not affect the attachment apparatus of the teeth. Periodontitis affects connective tissue attachment and adjacent alveolar bone. The classic progression of inflammatory periodontitis is characterized by a highly reproducible microbiological transition of the subgingival microflora from a mainly facultative Gram-positive microbiota to highly pathogenic Gram-negative rods and motile organisms. Although it was estimated that the subgingival microbiota consists of more than 300 different species (207), only very few of these species are actually involved in the initiation and progression of the periodontal disease process. Among these putative periodontal pathogens are members of the genera

Porphyromonas, Fusobacterium, Wolinella, Actinobacillus, Capnocytophaga, and

Eikenella. For example, Porphyromonas gingivalis has been implicated in chronic and advanced adult periodontitis (206). Actinobacillus actinomycetemcomitans in localized juvenile periodontitis (211, 257). Prevotella intermedius and Treponema denticola are involved in acute necrotizing ulcerative gingivitis (28, 113).

Capnocytophaga spp. appear to play an important role in advanced periodontitis in juvenile diabetes (128). Research studies suggest that Wolinella recta, Eikenella corrodens, and Bacteroides forsythus may play a role in the progression of periodontal disease (36, 124, 125, 187, 230).

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2 The subgingival plaque microbiota is of such complexity that, at present, it is not clear whether these periodontal diseases are the result of a pathogenic synergy, or are the result of a monoinfection by an invading or opportunistic member of the resident oral microbiota. It is more likely that the development of periodontal disease involves a consortium of the plaque microbiota that interacts in a cooperative or synergistic manner (62, 129, 232).

2. Porphyromonas gingivalis

2.1. Nutrition requirements of P. gingivalis

P. gingivalis is strictly anaerobic, asaccharolytic (201) and requires hemin and menadione (vitamin K) for growth (59). It was suggested that the dark pigment of P. gingivalis colonies on blood agar plates resulted from storage of hemin (59). Rizza et al. (183) showed that P. gingivalis cells grown in hemin replete medium were able to divide 8 to 10 times in hemin depleted medium, suggesting the accumulation and storage of this compound. The ability of P. gingivalis to scavenge hemin from hemoglobin after initial hemagglutination followed by lysis of the erythrocytes by the membrane-bound cysteine proteinase, gingivain, was suggested (196,198). Shah et al. (193) demonstrated that the hemoglobin derivative produced by P. gingivalis was protohemin with traces of protoporphyrin. While P. gingivalis may use hemin or protohemin in its electron transport system, it is not clear how adenosine 5'- triphosphate is generated via electron transport. Rizza et al. (183) demonstrated a membrane-bound respiratory system involving flavoprotein, cytochrome C, and a carbon monoxide-binding protein. Gibbons and MacDonald (59) proposed that vitamin K functioned as an electron carrier in electron transport. Vitamin K was also found to stimulate synthesis of phosphosphingolipids in the cell envelop, suggesting it may play a role in membrane permeability (110). Since P. gingivalis can not catabolize carbohydrates, alternative sources of energy was investigated. It was shown that protease peptone and trypticase were highly stimulatory to P. gingivalis

3 growth, bactocasitone and acidase peptone were moderately stimulatory, while casamino acids was a poor substrate for the growth of this microorganism (194). P. gingivalis preferred peptides over amino acids for growth (199). Studies showed that

P. gingivalis preferentially took up short peptides (10 to 14 residues) than long peptides (55). Although peptides appeared to be the favorite substrate for growth, P. gingivalis can also use amino acids under peptide limitation (55). Among them, aspartate, and to a lesser extent glutamate (and their corresponding amides, asparagine and glutamine), were significant catabolic amino acids (55, 200). In the gingiva, peptides are probably derived from the hydrolytic activities of both host and bacterial cells against host proteins such as immunoglobulins, hemoglobin and related complexes, and other host tissue proteins and secretions (21, 22, 173, 222). P. gingivalis possesses a large arsenal of proteases which can degrade host proteins to peptides (see P. gingivalis proteases).

2.2. Pathogenecity of P. gingivalis

P. gingivalis has been repeatedly implicated in the establishment and progression of periodontal diseases (206, 210, 242). White and Mayrand (247) demonstrated that

P. gingivalis was present in the gingival sulcus of patients with severe inflammation, but absent from healthy sites. P. gingivalis has been recovered from subgingival cultures of patients with generalized advanced periodontitis (247) and from adult patients with actively progressing periodontitis (208, 231). Di Murro and co-workers

(31) found P. gingivalisto be consistently associated with the subgingival microflora in patients with rapidly progressive periodontitis. P. gingivalis has also been suggested to be involved in severe, recurrent adult periodontitis (238) and both generalized and localized juvenile periodontitis (97, 114, 142, 249). Immunological studies provide additional evidence of the involvement of P gingivalis in periodontal diseases. Patients diagnosed with adult periodontitis and generalized juvenile periodontitis have higher serum antibody levels against P. gingivalis Vnan other groups of individuals (38,

4 43). Studies also showed elevated antibody levels to P. gingivalis in gingival crevicular fluid of patients with periodontitis (37, 233). Genco et al (50) showed that P. gingivalis strains in monoinfected murine tissue cages caused cachexia, ruffling, general erythema, phlegmonous ulcerated necrotic lesions, and death. The pathogenicity of P. gingivalis has also been demonstrated in other animal model infection studies (50, 51, 63, 66, 224, 241).

3. Virulence factors of P. gingivalis

A variety of putative pathogenic factors of P. gingivalis have been identified which may contribute to the colonization and virulence of this oral pathogen. These factors have been shown to have wide-ranging effects on tissue comprising the periodontium as well as on host immune mechanisms. These determinants include the ability to adhere to epithelial surfaces or other bacterial species, the ability to invade host tissues, the production of toxins or enzymes which are destructive to the host tissue, as well as the bacterial surface components such as lipopolysaccharides or capsular material which can induce infection or protect the bacteria from host immune responses. These studies have usually been conducted in vitro and their virulence remains to be determined in vivo.

3.1. Adherence The ability of periodontopathic bacteria to adhere to surfaces in the oral cavity are prerequisites for the production of infectious diseases. Surface structures are associated with adhesins which are responsible for attachment to specific host receptors. P. gingivalis cells have several adhesion factors for tooth surface, periodontal tissues, and other oral bacteria. Surface components including fimbriae, hemagglutinating factors, and some other surface binding properties, may play a significant role in subgingival colonization by P. gingivalis.

3.1.1. Fimbriae

5 P. gingivalis fimbriae have been isolated and purified, and their morphological, immunological, and chemical properties characterized (254, 255). The fimbrial subunit (FimA) has an apparent molecular weight of 43 kDa and is located on the cell surface (216). The size and antigenic heterogeneity of the 43 kDa protein among strains of P. gingivalis was compared by Lee et al. (107). P. gingivalis fimbriae was shown to be involved in interactions of this organism with host cells (70, 156, 256).

Fimbriae are specific antigens of P. gingivalis (70), and immunization of germ-free rats with fimbriae prevented P. gingivalis induced periodontitis (42). Fimbriae likely promoted adherence and colonization of P. gingivalis in periodontal pocket (83, 106).

A fimA-inactivatedP . gingivalis mutant had significantly reduced ability to bind saliva- coated hydroxyapatite, and in a gnotobiotic rat model of periodontal disease, the mutant had diminished ability to cause periodontal bone loss (121). These results suggest the fimbriae of P. gingivalis may play a significant role in bacterial pathogenicity.

3.1.2. Hemagglutinin Hemagglutinating activity is a marker of the ability of bacterial cells to attach to the host cells. P. gingivalis cell surface proteins possessed strong hemagglutinating activities. In contrast, capsular polysaccharide extracted with alkali, acid, EDTA, or phenol had no hemagglutinating activity (229). Several research groups have extracted hemagglutinating factors from P. gingivalis strains (82, 161). The extracts consisted of several proteins but no detectable lipopolysaccharide. They found the extracted hemagglutinating factor could be inhibited by a low concentration of L- arginine. The hemagglutinin is distinct from the fimbriae (143, 144). There was evidence that some of the hemagglutinating activity and trypsin-like enzyme activity were linked (154). However, other studies found that hemagglutinin was not associated with enzymatic activity and was heat stable (161).

6 Several studies have reported the characterization of hemagglutinin isolated form

P. gingivalis (17, 24, 34, 82, 143, 153, 161). Other research groups studied the hemagglutinins at the genetic level (108, 132, 179, 180). 4 hemagglutinin genes, hagA (179), hagB (179, 180), hagC (108), and hagD (109) have been cloned and sequenced. They all conferred hemagglutinating properties to E. coli transformants. HagA and HagD have 73.8% identity, while HagB and HagC shared 98.6% homology. hagA shared no homology to hagB by restriction mapping, Southern analysis as well as sequencing (180, 248). On the other hand, although the upstream and down stream regions of hagB and hagC had less than 40% homology, their open reading frames shared 99% homology. The high degree of homology between hagB and hagC suggests that they likely evolved from the duplication of a common ancestral gene. These multiple copies provide a way to solve the problem of synthesizing enormous amounts of a required gene product. The duplication of the hemagglutinin genes may indicate the importance of these proteins in the establishment of P. gingivalis colonization in the oral cavity.

3.1.3. Other surface binding molecules Boyd and McBride (17) isolated a bacterial aggregating component from the outer membrane of P. gingivalis and found it was composed of protein, carbohydrate, and a high-molecular-weight LPS fraction. Slots and Gibbons (209) and Okuda et al. (160) showed that P. gingivalis could attach to human epithelial cells. Lantz et al. (103) found P. gingivalis could bind to fibrinogen. P. gingivalis is able to adhere to Gram- positive plaque bacteria (17, 209). Ellen et al. (40) showed Actinomyces viscosus enhanced subsequent tooth colonization by P. gingivalis. P. gingivalis was able to coaggregate with strains of Treponema denticola (192) and Fusobacterium nuleatum (92). However, these possible surface binding molecules have not been well defined.

3.2. Capsule

7 Bacterial capsules have various functions: they can serve as physicochemical barriers for the cell, they provide protection against desiccation by binding water molecules, and they are antiphagocytic in that they function to avoid engulfment by polymorphonuclear leukocytes (PMNLs). Capsule may also promote attachment of bacteria to other bacteria (19).

P. gingivalis has an electronic-dense layer external to and associated with the outer membrane. It consists of exopolysaccharides and the thickness varies from strain to strain (239). It was suggested that the degree of encapsulation and resistance to phagocytosis were factors which might influence the invasiveness of P. gingivalis strains in induced infections (68, 239). However, in a mouse pathogenicity model, the correlation between the presence of capsule on P. gingivalis strains and virulence of the bacteria was not established (223). The exact role of capsule in the pathogenicity of P. gingivalis is not clear. Grenier and McBride (66) found P. gingivalis cells recovered from an experimental infection had a thicker and much denser extracellular material than cells grown in culture medium. This result suggests that surface structures are of importance for the pathogenicity of P. gingivalis.

3.3. Lipopolysaccharide (LPS)

The LPS from P. gingivalis showed very little endotoxic activity in classical endotoxin assay such as Limulus lysate assay or Schwartzman test (78, 225, 226). P. gingivalis whole cells or its purified LPS stimulated bone resorption (81, 137, 145,

227). Miller et al. (137) also showed that P. gingivalis LPS could inhibit bone collagen formation. The P. gingivalis LPS could induce interleukin-1 production which might play a role in the pathogenesis of adult periodontitis (72). They could also inhibit gingival fibroblast proliferation (104).

3.4. Vesicles

P. gingivalis cells can produce vesicles whose composition appears to be similar to the outer membrane (131). The vesicles exhibited proteolytic and collagenolytic

8 activities and were able to hemagglutinate erythrocytes (61). They also promoted bacterial adherence between homologous P. gingivalis strains and mediated attachment between non-congregating bacterial species (61). Since P. gingivalis vesicles carry virulence factors from the outer membrane, they may act as damaging weaponry delivered by the bacteria. Naito et al. (146) showed that P. gingivalis vesicles adhered to collagen-coated hydroxyapatite beads which mimicked the gingival ligament. Pretreatment of collagen-coated hydroxyapatite beads with vesicles inhibited the subsequent adherence of P. gingivalis cells.

3.5. Proteases There are four types of proteinases according to their catalytic mechanisms: serine proteinase, metallo-proteinase, aspartic (or acid) proteinase, and thiol (or cysteine) proteinase. Proteinases can be distinguished on the basis of their sensitivity to various inhibitors.

P. gingivalis produces a number of extracellular and/or cell surface associated proteolytic enzymes which can be divided into 4 types: trypsin-like activity, collagenase activity, glycylprolyl peptidase activity, and Pz-peptidase activity.

3.5.1. Trypsin-like activity

Most of P. gingivalis proteases are referred to as "trypsin-like", on the basis of their substrate specificities. However, it is now clear all of them appear to be cysteine proteinases (14, 26, 48, 49, 87, 93, 101, 153, 174, 191, 197). Trypsin-like activity is used here referring to proteinases that have arginine-specific and/or lysine-specific proteolytic activities. Numerous studies reported the purification of trypsin-like proteinases from P. gingivalis (47, 48, 162, 164, 215, 218, 253). These purified preparations were able to hydrolyze benzoyl-DL-arginine-p-nitroanilide (BApNA) and the proteolytic activities could be enhanced by reducing agents such as dithiothreitol, cysteine, and (3-mercaptoethanol. This suggests that these enzymes belong to the thiol proteinase group. These enzymes can hydrolyze a variety of native proteins

9 including BSA, gelatin, and casein. The molecular mass of the enzymes are in the range of 35 to 300 kDa. From these data, it is hard to determine how many proteases exist in P. gingivalis, due to the different strains studied, different culturing conditions, different fractionation and purification procedures. Moreover, a variety of substrates, activators, and inhibitors have been used in various assays to characterize these enzymes. The majority of trypsin-like proteases have Arginine-X hydrolytic ability. Recently, several studies showed that P. gingivalis produced distinct cysteine proteinases that have peptide cleavage activity for lysine residues (29, 49, 174, 191). The lysine- specific cysteine proteinase and arginine-specific cysteine proteinase are considered to be major etiologic enzymes of P. gingivalis because of their abilities to degrade various physiologically important proteins (49, 191). However, much of the data on these proteinases with regard to molecular mass, association with hemagglutinin activity, substrate specificity, and sensitivity to various protease inhibitors are ambiguous. Potempa et al. (177) compared the arginine-X and lysine-X specific activity distribution among different fractions of 6 commonly used P. gingivalis strains and analyzed each strain for the presence of gingipain (gingipain is a name proposed for Arg-X or Lys-X proteinase in P. gingivalis) by Western blot analysis, zymography, and enzyme active-site labeling. They concluded that the multiple forms of trypsin-like activity in P. gingivalis were due to the presence of either Arg-gingipain (RGP) or Lys- gingipain (KGP). This hypothesis was partially supported by recent studies in molecular cloning and characterization of cloned P. gingivalis protease genes. A number of P. gingivalis protease genes have been cloned and sequenced, including rgp-\ (172) and agp (159), prpRI (3), prtR (93), prtH (44), cpgR (54) , prfT (120, 163), tpr (16), and prtC (89). Comparison of the genes associated with Arg-X activities-rgp-

1, prpRI, prtR, agp, prtH and cpgR, showed they were highly homologous to one another. As shown in Table 1, these 6 genes may represent identical loci on P.

10 Table 1. Comparison of P. gingivalis genes encoding homologous Arg-X specific proteases3.

5' DNA regionb DNA coding region 3' DNA region Gene/strain Ref. Length % Iden.d Length c % Iden. Length % Iden. rgp-1 948 100 5,115/5,115 100 264 100 (172) prfR/W50 84 100 5,115/5,121 99.1 75 96.0 (93) prpR1/W50 627 99.8 4,480/4,578 99.0 NAe NA (3) agp/381 535 100 2,976/2,976 99.9 264 98.8 (159) prflH/W83 NA NA 2,369/2,369 98.1 264 96.9 (44) cpgR/ NA NA 1,452/1,464 85.2 NA NA (54) ATCC33277 a. Adopted from Barkocy-Gallagher et al. (12). b. Regions are defined by the rgp-\ sequence. For example, the 3" DNA region for the prtH gene comparison included DNA within the predicted coding region of prtH but beyond the rgp-1 coding region. c. Length of compared region in bases. Ratios are length of the compared region to total length of available sequence. Regions of low homology due to frameshifts, gaps, or deletions were excluded. d. % Iden., percent identity of the compared region with rgp-^. e. NA, not available.

11 gingivalis chromosome. They also shared high homology to the C-terminal portion of

P. gingivalis protease gene prt? (12) and a hemagglutinin gene hagA (71). Pike et al. (174) suggested that besides its proteolytic activity, the C-terminal region of Rgp-1 could act as a hemagglutinin. The highly homologous region found in the C-terminal halves of HagA, Rgp-1 and PrtP could represent regions functionally important in hemagglutination. Further studies are needed to clarify the relationship between these proteases and their association with hemagglutinating activity. Recently, Okamoto et al. (157) and Pavloff et al. (171) cloned and sequenced a lysine-specific gingipain {kgp) gene in P. gingivalis. They found the initial translation product was a large precursor, which was composed of four functional regions including a signal peptide, the N-terminal prosequence, the mature proteinase domain, and the C-terminal hemagglutinin domain which were very similar to the structure of the rgp gene product (158). Comparison of the nucleotide sequences of kgp and rgp revealed that there was great homology between them. The reason of why so many protease and hemagglutinin genes share high homology to each other is not clear. One hypothesis is that they resulted from recombinational rearrangement by transposition. Maley and Roberts (122) characterized an insertion element, IS1126, from P. gingivalis. Recently, Wang et al. (244) reported a second endogenous P. gingivalis IS element, PGIS2, and provided evidence that this element can transpose within P. gingivalis . These IS elements may be responsible for the gene rearrangements and duplications of the trypsin-like proteases and the hemagglutinins. Another explanation for the different organization of some of the cloned protease genes is that they may have originated during cloning procedures.

However, one cloned "trypsin-like" P. gingivalis protease, PrtT (120, 163), can not be classified as RGP or KGP since it has no homology to them either by DNA sequence or by amino acid sequence analysis. There is also evidence that some P. gingivalis proteases possessed both Arg-specific and Lys-specific activities (29, 174).

12 Therefore, the suggestion to classify P. gingivalis "trypsin-like" proteases to either Arg- gingipain or Lys-gingipain simplified the complexity of protease system in P. gingivalis.

3.5.2. Collagenase activity Gibbons and MacDonald (58) first demonstrated the collagenolytic activity of P. gingivalis. This was confirmed by later reports that P. gingivalis had collagenase activity (15, 130, 184, 213, 214, 221, 234, 237). Differing from vertebrate collagenases, which produce a 3/4 to 1/4 cleavage pattern, P. gingivalis collagenolytic activity cleaves collagen to multiple fragments of low molecular weight (15, 184, 234). It is not clear if this collagenolytic activity resulted form the action of nonspecific proteinases or collagenases or both since these enzyme preparations were not pure. Kato (89) reported the cloning of a collagenase gene prtC. The recombinant collagenase degraded soluble and reconstituted fibrillar type I collagen and heat denatured type I collagen.

3.5.3. Glycylprolyl peptidase activity

There has been several reports of glycylprolyl peptidase activity in P. gingivalis (1, 13, 65, 90, 141, 219). Four enzymes able to cleave the glycylprolyl dipeptide were purified and characterized (1, 13, Grenier, 1987 #24, 65, 141). These enzymes could be inhibited by phenylmethylsulfonyl fluoride (PMSF) and diisopropylfmorophosphate, suggesting they belonged to serine protease. Reducing agents did not enhance their activities. Since collagen has high contents of glycine and proline, these glycylprolyl peptidases may participate in the degradation of short peptides following the initial cleavage of collagen by specific collagenases.

3.5.4. The Tpr protease and Pz-peptidase activity

Park and McBride (169) reported the cloning of a P. gingivalis protease gene. Subsequently Bourgeau et al. (16) cloned and sequenced a tpr protease gene. Further analysis of the gene restriction map and characterization of the recombinant protease suggested they had cloned the same tpr gene. Analysis of the DNA

13 sequence (16) found putative -10 and -35 promoter region and a potential Shine- Dalgarno site located 10 nucleotides upstream from the start codon. There is a sequence resembling a rho-independent transcription termination sequence that contains an inverted repeated sequence with the potential to form a hair pin structure with a 6-nucleotide loop and a 14-nucleotide stem including 5 pairs of CGs. A stretch rich in T-residues much like typical E. coli terminators and another stop codon (TAA) immediately follow this structure (Figure 2). The molecular weight of the cloned protease was approximately 90 kDa for the active protease on gelatin zymography and 50 kDa for the denatured protein in Western Blot. In vitro translation study and high level expression study proved that the gene encoded the 50 kDa protein. An isogenic mutant of the tpr gene was made by insertional mutagenesis in P. gingivalis W83 (168). On 2D-PAGE, a 55 kDa protein was missing in the mutant. This is consistent with the molecular weight of Tpr. A comparison of the amino acid sequences listed in the gene bank revealed the presence of a 23-amino-acid consensus sequence (amino acid 406-430) near the C- terminus that had a 60-65% homology to thiol or cysteine proteases from a wide range of species and also some P. gingivalis thio-proteases (Table 2). This region, which begins with a histidine residue, corresponds to the active site of papain (135) and may represent part of the active site of the thiol proteases in P. gingivalis. The protease was inactive at or below 20°C against azocoll as the substrate. Activity increased rapidly above 30°C and reached a maximum at approximately 50°C. Maximum activity was maintained up to 70°C and was lost completely at 90°C. This temperature stability may be surprising but there are examples of other proteases, such as protease K, that are active at 80°C. The effect of pH on the protease activity was also analyzed. The protease was inactive below pH 6.0, it reached maximum activity at pH 8.5 and retained its activity up to pH 12. Since P. gingivalis is an asaccharolytic organism that ferments peptides and amino acids, it is expected to

14 90 GTTAACGGTTTTTCATGCGTGATTTGTCCAAATTGCACCTTAAT^

-35 -10 180 CGTGATTTGTCCAAGTCTTTTCCTTAATATATCCCTTCATTTGTGTGACTCTTAAAGTSTAA

SD 270 TAAAATTTTTCAATTATGGAAAAGAAATTAGTACCGCAATCCAT^ MEKKLVPQS ISKERLQKLEAQAT''LT 360 CCTCAACAAGAGGAAGCGAAAGCCCGTAAAATCGAAAGAGAGAAAGCCAGACTAAAAGAACTGAACATTCCTACCGAATCTAAAGAATCC PQQEEAKARKI EREKARLKELN1 PTES KES 450 AAAGATTGCAGCCCTGCAGGGATGATCAATCCATATGCACTTACTGAAG KDCSPAGMINPYALTEVILERPLDWSNPRT 540 ACCGATATAGTAGAACGCGTGTTGGGTTCAAGCATGCAAGATCTATCCAAAGGCGACTCTGTATTAAGAGCCGGAAGAGACCAAAATGCT TDIVERVLGSSMQDLS-KGDSVLRAGRDQNA / . 630 GAGGTCAAAATCGTGGATTCAGTTTTGACTAAGACCCAAAGAGGGCAGGACGGTCT^ EVKIVDSVLTKTQRGQDGLERILESFNDTD 720 ATGCCTCCCGAAGAAAAAGAAGAAGCTGCTCCAAAGGCAAAAAAAGCAGCCCAAAAACTCGACATCGACGACCTCAGAGAGCAAGCACTG MPPEEKEEAAPKAKKAAQKLDIDDLREQAL 810 TCATCCACTACTATCACAAAGGAGATCAGCAAGATCATCCTTCCGACAAAAAATTTAAGAGATGATAATAATACAGTACATCAGTACAGA SSTTITKEI . S KIILPRKNLRDDNNTVHQYR 900 GAAGTCGGCTTCCAAAGCAATGGAGCACACAACTTGTGGGACACAGTAGTTCAAGGAATC^ EVGFQ.SNGAHNLWDTVVQGIAGDCYMLAAL 990 TCGGCTATAGCTTGGGTATGGCCCGCTCTATTGAATATGGATGTGGACATCAT3TCTAATCAAGATGAATC SAIAWVWPALLNMDVDIMSNQDEWRLYRYF 1080 ATAGGGCGCTCTAAACAGACATATGCCAACCGGCCGTCGGGGTCAGGTACCTCCACCAATGAAATTCTTCAGGAAGGATATTACAAAGTT IG'RSKQTYANRPSGSGTSTNEILQEGYYKV 1170 CCGATCTTTGCAAGGAGTCGCTATTGGTTCAACGGAGAATAC'K^CCGGCTC P I FA RSR YWFNGEYWPALFEQAYANWKF PN 1260 GATTCCAAATACAACGCAATCCTACAGATTGGAGGTGGCTGGCCTGAGGAAGCACTTTGCGAGCTGAGTG^ DSKYNAILQIGGGWP. EEALCELSGDSWFTS 1350 TCGGGAAAACTCATGCTTTCTTCTTTCACAGATCTGTCATTGC^ SGKLMLSSFTDLSLLNFMKSMCYSWKTIKP 1440 ATGGTGATTGTAACCCCATGCTGGGACCCTCTACCTCCTATGATCCCCGGA^ MVIVTPCWEPLPPMMPGIAAT HAYTVLGYT 1530 GTTTCCAATCGAGCCTATTACCTGATTATCCGCAATCCATCGGGAGTGACTGAGCCAACAGGAGATGGAGTGCTAAGCAAAAGAGATTGG VSNGAYYL.IIRNPWG VTEPTGDGVLS'KRDW 1620 GTTATCCACTTCGATAATATGAAGTGGTTCAATCTATCCAAAGACGATCGCATTTTC • VIHFDNMKWFNLSKDDG IFALRLDKVRENF 1710 TGGTACATCGCATATATGTATISATATCAGATTATATTATACGGTTTGAGAACG^TGAGTCGGCCCGGTAGAGCGGGCA WYIAYMY* 1800 TTTCTGTACACGTCAGGCAACAGGCTCT^^CCATCGATCTATCTGCCCAATTCCGAATCGACAAAGCTCCATATCGGAGCATACACT

Figure 2. Nucleotide sequence of the cloned tpr gene from P. gingivalis W83 (16). The deduced amino acid sequence of the open reading frame is indicated below the nucleotide sequence. Two stop codons are indicated by asterisks. Putative -35 and -10 promoter regions, as well as a possible Shine-Dalgarno (SD) sequence are indicated. The possible rho-independent dual terminator is indicated by divergent arrows. A 23-amino-acid consensus sequence which is highly homologous with thiol or cysteine proteases from a wide range of species is underlined.

15 Table 2. Comparison of active-site amino acid residues of various cysteine proteinases.

Active-site amino acid residues3 Proteinases

HAYTVLGYTVSNGAYYLIIRNPWGVT E P P. gingivalis Tpr

HTLTVVGKTITVSWQGEAMIYDMN P. gingivalis PrtH

LTLTWGYNKETVIKTINTNQEPN P. gingivalis PrtR

HAFVCDGYEPDGTFHFNWGWGGMSNGNF P. gingivalis PrtT

HAVTAVGYGKSGGKGYILIKNSWGPGWG Papaya cysteine proteinase IV

HAVLAVGYGEQNGLLYWIVKNSWGSNWG cathepsin H

HAVNIVGYSNAQGVDYWIVRNSWDTNVG Dermatophagoides pteronyssinus

cysteine protease

HGVLLVGYNDNSNPPYWIVKNSW Trypanosome brucei cysteine

protease

HAIVIVGYGTEGGVDYWIVKNSWDTTWG kiwi actinidin

HAIRILGWGIENGVPYWLVANSWNVDWG cathepsin B

HAVAAVGYNPG YILIKNSWGTGWG E papain

a. Sequences are compared to Tpr, conserved amino acid residues are in bold.

16 produce protease that is functional over this range of pH. Tpr activity was sensitive to TLCK, TPCK, and EDTA, which inhibit both serine and cysteine proteases. It was also inhibited by PCMB and divalent cations, including Hg2+, which suggests that thiol groups may be involved in a configurational or catalytic role. Furthermore, reducing agents such as cysteine, dithiothreitol, and (3-mercaptoethanol stimulated activity. Other protease inhibitors such as amastatin, bestatin, pepstatin A, and aprotonin had no effect on protease activity up to 50 |ig/ml. The protease activity against other natural and synthetic protease substrates was tested. This enzyme was particularly active against general protease substrates such as azocoll, casein, BSA, and gelatin, but had no effect on native collagen or synthetic peptide such as BApNA, SAAPpNA an glycyl-L-arginine p-nitroanilide. It did not cleave [14C]-labeled type I chick embryo collagen, suggesting that it is not a collagenase, nor a trypsin-like protease. In the tpr isogenic mutant, Pz-peptidase activity was greatly reduced suggesting Tpr protease is a Pz-peptidase (167). Pz-peptidase activity in P. gingivalis has been reported by several research groups (85, 152). Ng and Fung (152) suggested that the Pz- peptidase activity of P. gingivalis is associated with the cell membrane, which is consistent with the result of the tpr analysis (170). Hino et al. (77) showed high levels of Pz-peptidase activity in inflamed gingiva. Sojar et al. (217) reported purification of a membrane-associated protease from P. gingivalis ATCC33277 capable of hydrolyzing Pz-peptide as well as salt-solubilized collagen. This enzyme is believed to be another

P. gingivalis protease since its N-terminal sequence is different from the deduced N- terminal sequence of Tpr.

3.5.5. Potential roles of P. gingivalis proteases in pathogenesis

P. gingivalis can produce various kinds of proteases that may play numerous roles in pathogenesis, such as host tissue destruction, disruption of host defense mechanisms, bacterial adherence and bacterial nutrition. For example, P. gingivalis can degrade collagen (14, 15, 105, 213, 214, 221, 228, 234) and fibrinogen (102)

17 which are important matrix proteins in maintaining tissue integrity. P. gingivalis proteases can hydrolyze immunoglobulin A (IgA), IgG (46, 64, 91, 186, 222), several complement proteins, including C3, C4, C5, C5a, and factor B (188, 220, 222, 224). Degradation of these key proteins of the immune system may enhance its pathogenic potential. P. gingivalis proteases may also promote bacterial adherence to subgingival sites (57, 60, 153, 154) and to other oral microorganisms (41, 112).

Kontani et al. (96) found RGP produced by P. gingivalis enhanced the adherence of purified fimbriae to fibroblasts or matrix proteins, such as collagen, fibronectin, and laminin. They recently found P. gingivalis RGP protease could expose a cryptitope in matrix protein molecules, i.e. the C-terminal Arg residue of the host matrix proteins, so that P. gingivalis could adhere through fimbriae-Arg interaction (95).

P. gingivalis is an asaccharolytic bacterium whose metabolism is dependent on the uptake of small peptides and amino acids (195, 199, 243). P. gingivalis proteases can degrade host proteins to provide nutrients for the growth of the bacterium (25). A key factor for the growth of P. gingivalis, hemin, is found in complexes with host proteins such as albumin, hemopexin, haptoglobulin, and transferrin. P. gingivalis proteases can degrade these proteins (22) and release small peptides, amino acids, and hemin- related compounds to promote bacterial growth.

3.6. Other enzymes and metabolic end products

Superoxide dismutase (SOD) is an enzyme of P. gingivalis presumed to be involved in protecting the cells against neutrophils generated bactericidal superoxide anion (O2") (4). It may also protect P. gingivalis cells when occasionally exposed to oxygen in the air (6). A sod gene-inactivated P. gingivalis mutant died immediately after exposure to the air, whileas the wild-type strain showed no decrease in viability after 5 hours of exposure to the air (147). Therefore, SOD was considered a virulence factor that contributes to the pathogenicity of P. gingivalis (131).

18 End products of P. gingivalis metabolism are toxic to mammalian cell lines. Butyrate and propionate are potent inhibitors of various cultured human or animal cell lines (205, 236, 240). Indole and ammonia (119) and volatile sulfur compounds including hydrogen sulfide, dimethyl disulfide, and methylmercaptan (235) are potential toxic factors.

4. Overview of molecular biology of P. gingivalis

4.1. Cloning of P. gingivalis genes

Molecular biology techniques were applied to P. gingivalis only recently. It was not until 1988 that the first P. gingivalis gene was cloned (32). Since then, numerous P. gingivalis genes have been cloned and sequenced (Table 3). There is little knowledge of P. gingivalis promoter structure. Klimple et al. (94) used antisera to the

E. coli RNA polymerase core enzyme and antisera to the sigma-70 factor to investigate if P. gingivalis RNA polymerase was related to E. coli RNA polymerase. Their studies showed no cross-reacting proteins between E. coli and P. gingivalis. P. gingivalis

RNA polymerase was much less processive on E. coli template than on its own and vice versa. They proposed that P. gingivalis RNA polymerase and E. coli RNA polymerase were so different that the genes from one organism may not be expressed in the other.

However, some of the cloned P. gingivalis genes appeared to be transcribed from promoters located on the cloned DNA sequence, while others were expressed under the control of E. coli promoters. The putative P. gingivalis promoter regions in some of these cloned genes were identified (Table 3), but there was no genetic analysis. Therefore, it is not clear if these putative promoter regions identified in some cloned P. gingivalis genes and expressed in E. coli are actually functioning in P. gingivalis. Qn the other hand, it is not clear whether a P. gingivalis promoter could be recognized by the RNA polymerase of E. coli. Expression of some recombinant P. gingivalis proteins

19 Table 3. Sequenced P. gingivalis genes and some features.

Strain Gene ORF Promoter mRNA Transcription Ref. start site

381, 2651 fimbrillin {fimA) 1.04 kb -35 TTGGGAC 1.3 kb NDa (5, 32) -10 TGTAAC

381 hemagglutinin 1.05 kb -35 TTGACC 1.15 kb ND (179) (hagB) -10 TATAAA or -35 TTGACA -10 AATAAT

W50 prtR 1.0 kb NFD ND ND (93)

ATCC53977 prfT 2.7 kb -35 TTCAGA 3.3 kb Start 510 bps (120) -10 TACCAT upstream of ATG

ATCC53977 prtC 1.0 kb NCC 4.4 kb ND (89)

W83 tpr 1.45 kb -35 TTCAGG 1.7 kb Start at an A This -10 GCTCTT residue 215 bp study upstream of ATG

W83 tpr 1.45 kb -35 TTTGTG ND ND (16) -10 AAGTC

ATCC53977, sod 0.58 Kb -35 TTTCACT 1.1 kb ND (5, 27)

2651 (superoxide -10 TAAAAG dismutase)

ATCC33277 sod 0.58 kb -35 TTACACT ND ND (148) (superoxide -10 TAAAAG dismutase)

ATCC33277 pgagMgdh 1.34 kb -35 TTGAAA ND ND (86) (glutamate -10 TAGCTA dehydrogenase)

381 Argingipain 3.0 kb NF ND ND (159)

W83 prtH 2.9 kb NF ND ND (44)

W50 prpRI 4.6 kb NF ND ND (3) (incompl (>300bp

ete) up)

20 ATCC53977 hemR 1.27 kb NF 3.0 kb At a C residue (88) 240 bp upstream of ATG

W50 mcmk 1.85 kb -35 TAAACA ND ND (84) (methylmalonyl- -10 TAAAGT

CoA mutase)

W12 prtP 5.2 kb NC ND ND (12) (porphypain) a. ND, not done b. NF, not found. c. NC, not cloned in E. coli.

21 may be due to the fortuitous presence of DNA sequences homologous to that of a E. coli promoter but are not the promoters in P. gingivalis as indicated in this study.

4.2. Genetic manipulation of P. gingivalis

P. gingivalis produces a large number of putative virulence factors that are implicated in the initiation of periodontal diseases and could cause tissue destruction. One of the most common strategies to determine the exact role of a virulence factor in disease process is to generate an isogenic mutant deficient in the suspected virulence factor. However, molecular manipulation of P. gingivalis was hindered by the lack of a well-defined genetic system in this organism. Genetic manipulation of some Gram-negative bacteria with plasmid DNA or linearized DNA was achieved by treating the cells with divalent cations, freeze/thaw methods, or electroporation to induce competence for DNA uptake. However, these methods had not worked successfully in P. gingivalis. Yoshimoto et al. (252) tried to transform P. gingivalis by electroporation and found if the DNA was isolated from a heterologous strain or other bacteria, no transformant could be obtained. The transformation frequency with homologous DNA was only 5x103 per u.g DNA, suggesting that P. gingivalis possesses a restriction-modification system that modifies its DNA. A gene, pg/IM, that encodes a DNA methylase has been identified and cloned from P. gingivalis (10).

One means of genetically altering P. gingivalis is by transposon mutagenesis. The successful introduction of R751 ::*Q4 into P. gingivalis and transposition of Tn4351 into the chromosome made it possible to select shot gun mutations (35, 52, 53, 79, 202). However, due to the difficulty of assigning a specific phenotype change to the specific change in the chromosome and to the possibility that the transposon might cause polarity effects, activation of nearby genes, DNA rearrangements or could move around on the chromosome randomly, it was necessary to have a better defined mutagenesis strategy.

22 Most of the genetic studies of extrachromosomal elements and gene transfer in anaerobic bacteria were done with the Bacteroides species and Clostridia species

(155, 185). P. gingivalis does not have any naturally occurring plasmids. Due to the close relationship between Bacteroides species and P. gingivalis, several investigators have tried to use shuttle vectors developed for gene transfer between E. coli and colonic Bacteroides species to introduce foreign or modified genes into P. gingivalis. One advantage of a conjugation system for the transfer of genetic material is that P. gingivalis restriction systems can be circumvented. Plasmids pE5-2 (178), pVAL-1 (35), pNJR5 and pNJR12 (123) were transferred successfully from E. colito P. gingivalis by conjugal mating. The frequency of transferring pE5-2 and pVAL-1 into P. gingivalis was lower than transfers into colonic Bacteroides species. Whereas transfer frequencies for pNJR5 and pNJR12 into P. gingivalis were similar to that of Bacteroides recipients. However, the stability of the transferred plasmids in P. gingivalis was not consistent. Maley et al. (123) reported these transconjugants were difficult to maintain on selective media. Park and McBride (170) reported the shuttle plasmid pNJR12 was not stably maintained in P. gingivalis. They found after several transfers in growth medium with antibiotic selection the transconjugants stopped growing. Dyer et al. (35) found that plasmid pVAL-1 was lost from P. gingivalis transconjugants when antibiotic selection was not maintained. Targeted homologous recombination mutagenesis is an excellent tool to determine the specific roles of genes in bacterial growth and pathogenicity. The development of suicide shuttle vectors pJRD215 (30), RSF1010 (9, 190), and pGP704 (139) have made this possible in P. gingivalis. Several research groups (45, 69, 86, 121, 149,

189) have successfully generated isogenic mutants in P. gingivalis by insertional mutagenesis.

Hamada et al. (69) and Malek et al. (121) found the fimA deficient mutant had a diminished adhesive capacity to tissue-cultured human gingival fibroblasts and

23 epithelial cells and was less pathogenic in animal infection models, suggesting that the FimA protein of P. gingivalis plays an important role in the pathogenesis of periodontal disease. Fletcher et al. (45) reported the mutation of p/tH protease gene in

P. gingivalis. Compared to the wild-type strain, they found the mutant was not able to cleave the C3 component of the complement system and in a mouse infection model, the prtH isogenic mutant was not virulent. Nakayama et al. (149) constructed RGP- deficient mutants and found two separate RGP-ehcoding genes {rgpA and rgpB) located on P. gingivalis chromosome. By analyzing the rgpA rgpB double mutant, they showed RGP might play important role in the virulence of P. gingivalis. Park and

McBride (166, 167) generated a tpr protease deficient mutant, P. gingivalis W83/PM. This mutant had greatly reduced ability to hydrolyze Pz-peptide, suggesting tpr is a Pz- peptidase. Kuramitsu et al. (98) isolated a cysteine protease mutant which was defective in the rgp-\ gene and evaluated the role of rgpA in the virulence properties of P. gingivalis. Compared to the parent strain, they found the mutant to be defective in interacting with Gram-positive bacteria as well as cultured epithelial cells. This suggested a potential role for cysteine proteases in the colonization process. They also observed reduced expression of fimbriae in the mutant. By analyzing rgpA rgpB double mutant, Nakayama et al. (150) also found the proteases were required for fimbrial protein processing. Decreased expression of fimbriae in P. gingivalis can result in loss of interaction with other oral bacteria and host epithelial cells (69).

Compared to the wild-type P. gingivalis strain, fimA mutant had no altered hemagglutinating ability, suggesting that fimbriae and hemagglutinin are two different identities (69, 121).

5. Gene regulation

5.1. Transcription regulation in prokaryotes Gene regulation can occur at any stage between the transcription of the genes to the formation of mature protein products. Numerous mechanisms exist for organisms

24' to regulate gene expression. Multiple layers of regulation may exist within a single gene. In prokaryotes, gene regulation is achieved mainly at the transcriptional level. Transcription in the eubacteria is catalyzed by a multisubunit RNA polymerase. The catalytic machinery resides in the core enzyme, designated p(3'a2 or E. The core enzyme can synthesize long, complementary RNA molecules from single-stranded templates, nicked DNA, and some copolymers such as poly d (AT). However, the core enzyme does not faithfully initiate transcription at promoter sites in the absence of a sigma (a) factor. Sigma factors are a family of relatively small, dissociable subunits that bind to the core subunits to form an initiation-specific enzyme, the RNA polymerase holoenzyme (Eg). Each holoenzyme specifically recognizes a set of related promoter elements with a consensus sequence determined by the bound a subunit.

Studies of the E. coli RNA polymerase showed the o subunit catalytically increased the number of RNA chains initiated without affecting the overall elongation rate of each chain (20), suggesting a factor may determine the specificity of transcription initiation.

Studies of transcription in Bacillus subtilis and its phage led to the first evidence for alternative a factors that play a role in altering the program of transcription during developmental processes such as phage growth and sporulation (116). It is now clear that most, if not all, eubacteria have a set of alternative a factors that control genes for specialized functions. These functions include endospore formation in gram-positive organisms (117), response to environmental change (76), expression of flagella and chemotaxis genes (75), and control of nitrogen metabolism (99). Based on the DNA sequences, and by inference the amino acid sequences, a factors can be divided into two apparently unrelated groups: the a70 family and the o54 family. The a70 family is named for the prototypic o factor: the 70-kDa primary a factor from E. coli. All eubacteria appear to have a single primary o factor that is sufficient for transcription of

25 all essential genes. However, some essential genes have multiple promoter elements and are transcribed by more than one type of holoenzyme. The a54 family of alternative a factor is named for the 54-kDa nitrogen regulation a factor of E. coli encoded by the ntrA (rpoN) gene. Members of this family participate in the regulation of diverse sets of genes often encoding proteins involved in nitrogen metabolism. In addition to their structural distinction, the process of transcription initiation by holoenzymes containing these two types of sigma factors differs. All promoters recognized by holoenzymes containing o70-type sigmas consist of two blocks of conserved sequences that are located at about 10 bp and 35 bp upstream of the transcription start at +1 and are separated by 15-20 bp of nonconserved sequence (Table 4) (33, 182). Both the DNA sequences of the two conserved regions and the optimal spacing between them are unique to the particular sigma factor. In contrast, all holoenzymes containing o54-type sigmas recognize promoters with conserved blocks of sequence that are separated by only 5 bp and are located at 24 bp and 12 bp upstream of the start point of transcription (Table 4) (67). Whereas holoenzymes containing a70-type sigmas are competent to form "open complexes" in the absence of auxiliary factors, those with o54-type sigmas require an auxiliary protein to carry out this process (245). Transcription represents only one component of a regulatory network whose overall purpose is to control the relative amounts of specific proteins produced at various stages of the cell cycle and in response to environmental and developmental signals. This process is also controlled at the level of protein synthesis by the competition between various mRNAs for ribosomes and by the relative stabilities of the various mRNAs. Regulation can also occur at the DNA level by selective gene duplication processes.

26 Table 4. Consensus nucleotide sequences for promoters recognized by RNA polymerase containing various sigma factors.

Sigma factor Organism "-35" region "-10" region

O-70 E. coli TTGACA TATAAT a32 E. coli TCTC-CCCTTGAA CCCCAT-TA o-A B. subtilis TTGACA TATAAT rjB B. subtilis AGGI I I AA GGGTAT

O-D B. subtilis CTAAA CCGATAT rjE B. subtilis ATATT ATACA

B. subtilis TGAATA CATACTA a* B. subtilis AC CATA--T

B. subtilis CAGGA GAATT--T a9P28 phage SP01 AGGAGA 111-111*

Ggp55 phage T4 none TATAAATA

Sigma factor Organism "-24" region "-12" region rj54 E. coli, Phizobium CTGG-A TTGCA

S. typhimurium

K. pneumoniae a. Tin phage SP01 DNA represents hydroxymethyluracil, which replaces thymine.

27 In E. coli, control over gene expression is exerted primarily at the level of transcription. The transcription cycle is presented in schematic form in Figure 3. This cycle is conventionally divided into the phases of initiation, elongation, and termination. Each of these phases is subject to modulation by several regulatory mechanisms. For example, there is regulation of complex gene sets by alternative sigma factors, these alternative sigma factors may play either a passive or an active role in directing the transcription of these diverse genes. In the passive mode of regulation, the sigma factor does not itself control change in the level of transcription of cognate genes, rather, an auxiliary transcription factor is the regulator of gene expression. Changes in the level of expression of the gene are mediated by altering the activity or amount of a regulatory protein in response to an environmental stimulus (e.g., the catabolite regulatory protein (CRP), AraC). In active control, the sigma factor itself is the regulatory protein in the induction of a cognate gene set. A remarkable diversity of control mechanisms regulates these sigmas: transcriptional activation, translational activation, degradation, proteolytic processing, and chromosome rearrangement.

Activators of initiation can stimulate promoter efficiency in E. coli in various ways. They can increase the affinity of the holoenzyme for the promoter at the level of closed promoter complex formation, they can increase the rate of formation of the open promoter complex, or they can speed the rate of promoter clearance by affecting one of the several steps of transcript initiation. In general , positive regulatory factors respond to the need of the cells by sensing chemical signals, including the concentrations of metabolic substrates, cofactors, and secondary messengers (151).

A classic example in E. coli\s the synthesis of cAMP in response to low glucose levels. The cAMP binds to the CRP, which in turn binds with increased affinity to a specific DNA element of its target operons.

28 Figure 3. Schematic representation of the overall transcription cycle. Adopted from Piatt and Bear" (175).

29 Repressors are subject to the same type of metabolic and developmental control and can also operate at various levels of promoter function. A repressor can block a promoter entirely thus prevent binding of the holoenzyme to the promoter. Other possibilities include interference with the closed-to-open promoter transition and the promoter clearance process. The end result of these and other mechanisms for controlling transcript initiation is an integrated network that ensures the appropriate expression of a given protein in a cell at a particular stage of life cycle or environment. NusA influences elongation and termination process in RNA transcription and also serves as an "anchor" protein that allows other factors to bind and exert their specific effects. As indicated in Figure 3, NusA protein interacts with core RNA polymerase at the time of sigma factor dissociation and remains bound until termination, at which point it dissociates. The bacterial transcription cycle is regulated by systems that are extremely complicated, activator and repressor proteins regulate initiation by RNA polymerase containing a70 or another a factor, and various termination and antitermination factors regulate RNA chain elongation by elongation complexes containing NusA. In each case, there are multiple protein-protein and protein-nucleic acid interactions that alter the functional properties of the RNA polymerase.

5.2. Gene regulation in P. gingivalis Bacterial growth in nature is affected by environmental conditions such as essential nutrient or cofactor availability, the accumulation of toxic products of metabolism, or the change in pH, Eh, or temperature. To cope with the changing environment, microbes have evolved the ability to change structurally and functionally to adapt to new environments. P. gingivalis is able to respond to environmental changes and modify its physiology and pathogenicity (see below).

5.2.1. Effect of nutrition on P. gingivalis

30 In the chemostat, changes in nutrient availability can affect the growth rate of P. gingivalis, which can in turn affect the enzyme profile (140). When P. gingivalis was grown in the chemostat with mucin, hemoglobin, or collagen as the limiting nutrient, the activity of hydrolytic enzymes was affected. Trypsin-like proteolytic activity was highest when the growth medium was supplemented with hemoglobin. In contrast, glycylprolyl dipeptidase activity was higher in the basal medium, and was repressed during growth with some of the limiting nutrients. Robertson et al. (184) reported that in a peptide-depleted growth medium, P. gingivalis collagenolytic activity was enhanced. In the analysis of the PZ-peptidase activity of Tpr protease, we found the membrane fraction of P. gingivalis W83 cells grown in 0.5 TYE had 2 times of Pz- peptidase activity as the membrane fraction of cells grown in TYE, and 5 times as much activity as cells grown in BHI. Northern blot analysis indicated that the regulation of tpr expression mainly occurred at the transcriptional level (170).

5.2.2. Effect of hemin

P. gingivalis has an absolute growth requirement for hemin. In the host, hemin is usually complexed by the plasma proteins haptoglobin, albumin, and hemopexin. P. gingivalis proteases can degrade these hemin-complex molecules (22) and may release free hemin for uptake. Studies showed that hemin levels in the growth media affected the expression of P. gingivalis proteases. Under hemin restricted conditions, collagenolytic activity was the major enzyme activity whereas trypsin-like enzyme activity dominated under hemin excess conditions (126). P. gingivalis cells grown in hemin limited conditions had large numbers of extracellular vesicles (ECV) both surrounding the cell surface and free in the environment, whereas cells grown in hemin rich media had fewer ECV. Smalley et al. (212) showed that both cells and ECV from hemin-limited cultures had higher hemagglutinating activity and collagenolytic activity and suggested their production was a stress-induced response by the cell to the depletion of an essential nutrient. Growth of P. gingivalis in batch

31 cultures and in a chemostat showed the induction ot novel proteins in outer membranes under hemin limitation (165). Bramanti et al. (18) identified a 26 kDa hemin-repressible surface protein in P. gingivalis.

The pathogenicity of P. gingivalis is affected by hemin in the growth medium.

McKee et al. (134) found P. gingivalis W50 cells grown in hemin replete medium were more virulent than cells grown in hemin depleted or hemin limited medium in mice.

However, Bramanti et al. (18) found P. gingivalis cells were more virulent under hemin limiting contions in an animal infection study.

Although these studies supported the concept that some P. gingivalis virulence factors were regulated in response to hemin, the evidence at the molecular level was scarce. By using the Bacteroides fragilis transposon Tn4351 , Genco et al. (53) isolated a'transpositional insertion mutant in P. gingivalis, which is defective in hemin utilization and transport. The inability of this mutant to transport hemin resulted in a increase in extracellular vesicle production as well as increases in hemolytic, hemagglutination, and trypsin-like protease activities. The exact nature of this mutation is not clear. They proposed that this mutation may result from an insertion in a hemin-responsive regulatory gene that controls the expression of trypsin-like protease, hemagglutinating, and hemolysin activities, as well as the production of extracellular vesicles. Another hypothesis was that Tn4351 might have inserted in a gene required for hemin transport. Mutation of this gene would render the cell in a hemin-limited state. The observed changes were consistent with a state of hemin- limitation.

Recently, Karunakaran et al. (88) reported the cloning of a hemin-regulated protein,

HemR, from P. gingivalis ATCC53977. Based on the N-terminal homology to TonB proteins in other bacteria, it was suggested that this protein may be involved in hemin

(iron) uptake. Northern blot analysis and a hemR-lacZ reporter gene analysis indicated the expression of hemR is negatively regulated at the transcription level by

32 hemin, suggesting this gene product is expressed only under hemin-restricted conditions.

5.2.3. Effect of environmental pH The pH of the healthy gingival crevice is neutral but becomes slightly alkaline during inflammation and periodontal disease (39). pH changes in the growth medium can cause marked alteration in the enzyme profile of P. gingivalis (133). P. gingivalis produced collagenolytic activity optimally at or below neutral pH, whereas trypsin-like activity was maximal at pH 8.0.

5.2.4. Effect of temperature

Several heat shock proteins have been identified in P. gingivalis (118), and GroEL and GroES gene have been cloned (80). They have high homology to E. coli heat shock proteins GroEL and GroES and exhibited standard heat shock response.

Amano et al. (5) reported the biosynthesis of fimbriae and SOD in P. gingivalis is regulated in response to elevated temperature. At 39°C, there was an approximately

50% reduction in the amount of fimbriae and the expression of fimA mRNA also decreased. On the other hand, at the same temperature, there was a more than twofold increase in the amount of SOD activity and Northern blot showed there was higher expression of sod mRNA. Recently, regulation of fimA gene expression by temperature change was confirmed by a lacZ reporter gene analysis (250).

6. The purpose of this study Regulation of gene expression can be important in the expression of virulence factors and for adaptation of pathogens to the host environment. P. gingivalis is able to regulate the expression of various proteins and enzymes to adapt to altered environments. However, there has been no study to analyze P. gingivalis gene regulation at the genetic level and there is little knowledge of regulation mechanisms in P. gingivalis. The finding that the tpr protease is regulated by nutritional conditions provided us with a good candidate to analyze gene regulation in P. gingivalis.

33 II. MATERIAL AND METHODS

1. Bacterial strains and culture conditions

P. gingivalis strains were grown in brain heart infusion (BHI) broth (Difco Laboratories, Detroit, Ml) supplemented with hemin (5 |ig/ml) and vitamin K (1 (j.g/ml), or TYE broth (17 mg/ml trypticase peptone, 3 mg/ml yeast extract, 5 mg/ml NaCI and 2.5 mg/ml KH2PO4) supplemented with hemin (5 u.g/ml) and vitamin K (1 (ig/ml). Nutrient limited medium 0.5 TYE had the same contents as TYE except the trypticase peptone content was reduced to 5 mg/ml. To make BHI blood agar plates, laked human blood and agar were added to BHI broth to a concentration of 5% v/v and 1.5% w/v respectively. The cultures were incubated in a Coy anaerobic chamber (Coy Manufacturing, Ann Arbor, Ml) in a 5% CO2-10% H2-85% N2 atmosphere at 37°C. For selection of P. gingivalis transconjugants, gentamicin (200 u.g/ml), erythromycin (10 u.g/ml) or tetracycline (10 |ig/ml) were added to the media. E. coli strains were grown in Luria-Bertani (LB) broth (10 mg/ml tryptone, 5 mg/ml yeast extract, 10 mg/ml NaCI, pH 7.2) or Minimum A medium (2 mg/ml glycerol, 1 mM MgS04-7H20, 1 mg/ml

(NH4)2S04, 4.5 mg/ml KH2PO4, 10.5 mg/ml K2HPO4, 0.5 mg/ml sodium citrate-2H20). E. coli XL-1 blue was used as host for plasmid DNA. LB agar plates contained 1.5% agar. For selection purposes, ampicillin (50 |ig/ml), kanamycin (50 u.g/ml), tetracycline (10 u.g/ml), and trimethoprim (200 jig/ml) were used unless stated otherwise. Stocks of bacteria were stored at -70°C in 15% glycerol. Bacteria strains and plasmids used in this study are listed in Table 5.

2. DNA manipulation methods Plasmid DNA was isolated by the alkali lysis method and if necessary further purified by cesium chloride-ethidium bromide gradient centrifugation (8). Chromosomal DNA was routinely extracted from bacterial cells grown to early stationary phase by the miniprep method described by Ausubel et al. (8). Restriction enzyme digestion of DNA samples were carried out by following the enzyme

34 Table 5. Strains and plasmids.

Strain/plasmid Descriptions3 Source/reference

E. coli strain XL-1blue F'::Tn10proA+B+/aciqA(/acZ)M15/recA1 Stratagene MC1061 F-araA139(ara-leu)A(/ac)X74/7scfR2mcrAmcAB1 (181, 246)

P. gingival is strain W50 P. gingivalis wild type, Gmr W83 P. gingivalis wild type, Gmr ATCC33277 P. gingivalis type strain, Gmr W83/PM P. gingivalis W83 fpr-isogenic mutant, Emr (168) W83//acZ P. gingivalis W83 with tpr::lacZ fusion, Emr this study

Plasmid pTZ18R Ampr (136,251) pTZ19R Amp' (136,251) pBluescript Sk(-) /\mpr Stratagene pYS307 Ampr, tpr cloned in pTZ18R (169) pXCA601 Promoterless lacZ gene, Tcr (2) pBLU-1 Bam\-\\-Pst\ tpr fragment of pBY307 cloned into pBluscript SK(-) this study pBX-1 BamHI-H/'ndlll 3.5 kb lacZ fragment from pXCA601 ligated to EcoRI-H/ndlll digested pBLU-1 this study pTXZ18R Pst\-Hind\\\ 3.5 kb lacZ of pBX-1 cloned into pTZ18R this study pTXZ19R Psft-HindlU 3.5 kb lacZ of pBX-1 cloned into pTZ19R this study PTXZ18R-400 BamHI-H/'ndlll 4.3 kb tpr::lacZ fragment from pBX-1 cloned into pTZ18R this study pTXZ19R-400 BamHI-H/ncflll 4.3 kb tprr.lacZ fragment from pBX-1 cloned into pTZ19R this study pTXZ18R-100 BamHI-Psfl 500 bp PCR product XZ-100 cloned into pTXZ18R this study

35 pTXZ19R-100 BamH\-Psti 500 bp PCR product XZ-100 cloned into pTXZ19R this study pTXZ18R-30 Kpn\-Pst\ 430 bp PCR product XZ-30 cloned into pTXZ18R this study pTXZ19R-30 Kpn\-Pst\ 430 bp PCR product XZ-30 cloned into pTXZ19R this study pTXZ18R20 BamH\-Pst\ 380 bp PCR product XZ20 cloned into pTXZ18R this study pTXZ19R20 BamH\-Psti 380 bp PCR product XZ20 cloned into pTXZ19R this study pTXZ18R52 Kpn\-Psti 348 bp PCR product XZ52 cloned into pTXZ18R this study pTXZ19R52 Kpn\-Pstt 348 bp PCR product XZ52 cloned into pTXZ19R this study pTXZ18R137 BamH\-Pst\ 263 bp PCR product XZ137 cloned into pTXZ18R this study pTXZ19R137 BamH\-Pst\ 263 bp PCR product XZ137 cloned into pTXZ19R this study pTXZ18R-100A EcoRI-BamHI 120 bp PCR product XZ-100A cloned into pTXZ18R137 this study pTXZ19R-100A EcoRI-BamHI 120 bp PCR product XZ-100A cloned into pTXZ19R137 this study pTXZ18R-30A Kpn\-BamV\\ 50 bp PCR product XZ-30A cloned into pTXZ18R137 this study pTXZ19R-30A Kpn\-BamH\ 50 bp PCR product XZ-30A cloned into pTXZ19R137 this study R751 Tpr Tra+ IncP plasmid used to mobilize vectors from E. coli to Bacteroides recipients (203) pJRD215 Kmr, Smr, Mob+ Broad-host-range RSF1010-based cosmic vector, mobilized at high frequencies by IncP plasmids (30) pNJR12 Kmr, Smr, Mob+, Tcr, replicable in P. gingivalis (123) pBY2-IN Emr- tpr mutation (168) pBYZ BamHI-H/ncflll 4.3 kb tprr.lacZ fragment from pTXZ19-400 ligated to pBY2-IN this study pNTX-400 BamHI-/-//no1ll 4.3 kb tprr.lacZ fragment from

36 pTXZ19R-400 cloned into pNJR12 this study pNTX-100 BamHI-H/ndlll 4.0 kb tpr::lacZ fragment from pTXZ19R-100 cloned into pNJR12 this study pNTX-30 Kpn\-Hind\\\ 3.93 kb tpr::lacZ fragment from pTXZ19R-30 cloned into pNJR12 this study pNTX20 BamH\-Hind\\\ 3.88 kb tprr.lacZ fragment from pTXZ19R20 cloned into pNJR12 this study pNTX52 Kpn\-Hind\\\ 3.85 kb tprr.lacZ fragment from pTXZ19R52 cloned into pNJR12 this study pNTX137 BamHI-H/'ndlll 3.76 kb tprr.lacZ fragment from pTXZ19R137 cloned into pNJR12 this study pNTX-100A EcoRI-H/ndlll 3.9 kb tprr.lacZ fragment from pTXZ19R-100A cloned into pNJR12 this study pNTX-30A Kpn\-Hind\\\ 3.81 kb tprr.lacZ fragment from pTXZ19R-30A cloned into pNJR12 this study pNTX Pst\-Hind\\\ 3.5 kb lacZ fragment from pTXZ19R cloned into pNJR12 this study a. Kmr, Smr, Tcr, Emr, Tpr, Ampr and Gmr resistance to kanamycin, streptomycin, tetracycline, erythromycin, trimethoprim, ampicillin and gentamicin, respectively. Mob+, capable of being mobilized, Tra+, capable of self-transfer.

37 manufacturer's recommendations. After restriction digestion, DNA samples were analyzed by 0.8% agarose gel electrophoresis in a Tris-borate-EDTA buffer (89 mM Tris base, 89 mM boric acid, 2 mM EDTA, pH 8.0) unless stated otherwise. Following electrophoresis, DNA bands were stained by a 0.5 |ig/ml ethidium bromide solution and visualized under UV light. 1 kb DNA ladder (Gibco BRL) was used as DNA molecular weight standard. Subcloning of DNA fragments or PCR products was done by restriction digestion and electrophoresis in agarose gels which were prepared and run in a Tris-acetate-EDTA buffer (40 mM Trisacetate, 2 mM Na2EDTA-2H20, pH 8.5). The desired DNA fragments were excised from the gel and recovered by Glass milk purification as described by the manufacturer (The GENECLEAN® Kit, Bio 101 Inc. La Jolla, CA). DNA ligations were performed as described (8) with T4 DNA ligase.

Transformation of E. coli cells was done by electroporation as described (8). P. gingivalis transformation was done by E. coli-P. gingivalis conjugation.

3. Construction of P. gingivalis W50 DNA library

P. gingivalis W50 chromosomal DNA was isolated as described below. Bacteria cells grown to early stationary phase were harvested and washed once in TES buffer (50 mM Tris-HCI 50 mM EDTA, 50 mM NaCI, pH 7.5). Cells were resuspended in TES buffer with 1.6% SDS, 50 u.g/ml RNaseA, and incubated at 37°C for 1 hour. 25 |ig/ml of proteinase K was added and further incubated at 37°C for 1 hour. Proteins were removed from the cell lysate by extracting with buffer-saturated phenol 3 times, phenol/chloroform/isoamyl alcohol (25:24:1) 2 times, and chloroform/isoamyl alcohol (24:1) once. DNA in the aqueous phase was precipitated by ethanol. Chromosomal DNA was further purified by equilibrium centrifugation in a cesium chloride-ethidium bromide gradient as described (8). Purified chromosomal DNA was partially digested with Sau3A\ and size fractionated by using a sucrose gradient as described (8). DNAs from those fractions were analyzed on an agarose gel to determine their sizes.

38 Fractions containing DNA fragments of 3-12 kb were pooled, and ligated to SamHI digested, alkaline phosphatase treated pTZ18R by T4 DNA ligase.

The ligation mixture was used to transform E. coli XL-1 blue cells by electroporation as described (8) at 2.5 KV using a 0.2 cm gap cuvette. After incubation at 37°C for 1 hour in LB broth, the cells were plated on LB agar containing ampicillin (50 |ig/ml) and X-gal (40 |j.g/ml). After overnight incubation at 37°C, white colonies were isolated for further analysis.

4. Screening for proteolytic clones

E. coli transformants containing foreign DNA were grown on LB plates containing 50 |ig/ml ampicillin and saved as the master plates. They were replica plated on skim milk agar plates (LB agar supplemented with 15 mg/ml skim milk and 50 u,g/ml ampicillin) and incubated at 37°C in the air overnight. The cells were lysed by exposing to chloroform vapor for 30 min, and then incubated anaerobically at 37°C for up to a week. Proteolytic clones were identified as colonies showing clearing zones around them.

5. PCR and primers To analyze the effect of deletion mutations on the expression of tpr, the following tpr 5' fragments were generated by PCR. Primers used for PCR amplification are shown in Table 6. DNA fragment XZ-100 was generated by primers Bm300 and tprPst, fragment XZ-30 used primers Kpn365 and tprpst, fragment XZ20 used primers Bm200 and tprpst, fragment XZ52 used primers Kpn452 and tprpst, fragment XZ137 used primers Bm100 and tprpst. Fragment XZ-100A used primers Eco300 and Bm424, fragment XZ-30A used primers Kpn452 and Bm424. PCR amplification was carried out with Taq I polymerase (Gibco BRL). Twenty-five cycles were carried out at a denaturing temperature of 95°C for 1 minute, an annealing temperature of 55°C for 1 minute, and an extension temperature of 72°C for 1.5 minutes in a GeneAmp PCR system 9600 (Perkin-Elmer Cetus, Norwalk, Conn.) according to the manufacturers.

39 Table 6. Primers for PCR and primer extension analysis.

3 0 Primer Sequence Nucleotide number

Bm300 5'-ATTCGGATCCTCGGGTCTCGTCTG-3 ' 291-314 Fc BaMrW

Kpn365 5'•TTCAGGTACCAATTGTCAATT-3 ' 360-380 F Kpril

Bm200 5'TAATG^GATTiC.TAACGGTTTTTCATGC-3 ' 410-435 F BairH\

Kpn452 5'-AAATGGTACCTTAATTCG-3' 447-464 F Kpn\

Bm100 5'•CCTTAATGGATCCCTTCATTTGTG-3 ' 529-552 F SamHI

Eco300 5'•CACGAATTCGGCTGTTCG-3 ' 286-303 F EcoRl

Bm424 5'-•CGTTAGGATCCATTATTTCAA-3' 424-404 Rd BamH

tprPst 5'-CATCCCTGCAGGGCTGC-3 ' 801-785 R Psfl

tpr293 5'•GCTTTCGCTTCCTCTTGTTGAGGA-3 ' 710-687

tpr170 5'-AACTGTGACTTTABBCTCTTAC-3' 587-566 tpr64 5-•TGTGTACAAAAAAACTAACGAATTA-3' 482-458 a. The restriction sites are underlined and indicated. b. Nucleotide number corresponds to Figure 23. c. F stands for forward strand. d. R stands for reverse strand.

40 recommendations. The amplified DNA XZ-100, XZ-30, XZ20, XZ52, XZ137, XZ-100A and XZ-30A are indicated in Figure 4. The PCR products were digested with restriction enzymes whose sites were present on the primers and then cloned into corresponding sites on the lacZ fusion vector pTXZ19, resulting in deletions in tpr 5' region.

6. DNA sequencing DNA sequencing reactions were conducted by using Sequenase® Version 2.0 DNA polymerase by following the protocols provided by the manufacturer (United States Biochemical Molecular Biology Reagents/Protocols 1992, USB, Cleveland, Ohio) with some modifications: Annealing template and primer. 1.0 pmol primer, 0.5 pmol denatured double-stranded plasmid DNA pYS307, 2 u,l annealing buffer were combined in a 0.5 ml microcentrifuge tube and water was added to bring the total volume to 10 uJ. The tubes were incubated at 55°C for 2 minutes, and allowed to cool slowly to room temperature over a period of about 30 minutes. The tubes were then chilled on ice. Labeling reaction. 1 ul DTT (0.1 M), 2 uJ labeling nucleotide mix, 5 fj.Ci [a-35S] dATP, 0.5 \i\ Mn solution, and 3 units Sequenase enzyme were added to the annealed template- primer, mixed thoroughly and incubated for 2-5 minutes at room temperate. Termination reactions. 4 tubes were labeled "G", "A", "T" and "C". Each was filled with 2.5 uJ of the appropriate dideoxy termination mixture. These tubes were pre-warmed to 42°C. When the labeling reaction was complete, 3.5 u,l was transferred to the tubes containing dideoxy termination mixture, incubated at 37°C for 5 minutes, and then 4 uJ of stop solution was added to each termination reaction. Before loading on the gel, the samples were heated at 95°C for 3 minutes and 2-3 |il was loaded in each lane.

41 kb + + - + - + - + - + - kb

5.0 5.0 4.0 4.0 3.0 3.0 2.0 2.0

0.5 • 0.4 , 0.5 4 0.4 8:1 : 4 0.22 • 8:t 0.22

Figure 4. PCR products and tprr.lacZ reporter vectors. DNA fragments XZ-400, XZ- 100, XZ-30, XZ20, XZ52, XZ137, XZ-100A and XZ-30A were cloned in pTXZ18R (+) or pTXZ19R (-) and restriction digested with SamHI and Pst\. kb, DNA molecular weight in kilo bases.

42 Denaturing gel electrophoresis for DNA sequencing was carried out as described (8). A 6% polyacrylamide-urea gel was used. The gel was preheated on the power supplies to 1700V, 50W constant power for 30 minutes before loading sequencing samples. Electrophoresis was carried out at 1700 V, 50 W until the dye front reached the bottom of the gel. The gel was transferred to the top of a pre-wetted Whatman filter paper and dried at 80°C under vacuum for 1 hour before exposure to Kodak X-Omat film at -70°C.

7. Primer extension analysis

To map the 5' terminus of tpr mRNA in P. gingivalis or E. coli, primer extension analysis was conducted as described (8) with modifications. Three primers, tpr293, tpr170 and tpr64 (Table 6), were used for primer extension analysis. Labeling the primers with 32P was carried out as follows: The following reagents were mixed in a 0.5ml microcentrifuge tube: 2.5 u.l H2O, 1 uJ 10X T4 polynucleotide kinase buffer, 1 |J 0.1 M DTT, 1 (il 1 mM spermidine, 1 |il 50- 100 ng/ul oligonucleotide primer, 3 u1 10 uGi/ul [y-32P] ATP, and 0.5 u.l 20-30 U/uJ T4 polynucleotide Kinase. After incubation for 1 hour at 37°C, the reaction was stopped by adding 2 |il of 0.5 M EDTA and 50 |il TE buffer and incubated 5 min at 65°C. A small ion-exchange column was prepared by inserting a small plug of silanized glass wool into the narrow end of a silanized 1000 u.l pipette tip. After adding 20 u.l of AG 50W-X8 resin and 100 fil of DE-52 resin, the column was washed with 1 ml TEN100 buffer. The labeling reaction mix was loaded onto the column and washed with 1 ml TEN100, then with 0.5 ml TEN300. Radiolabeled oligonucleotides were eluted with 0.4 ml TEN600, collected as a single fraction and stored at -20°C in an appropriately shielded container until needed. Hybridize radiolabeled primer and RNA In separate microcentrifuge tubes 10 |il total cellular RNA (10 to 20 u.g), 1.5 u.1 10X hybridization buffer, and 3.5 |il radiolabeled oligonucleotide were added. The tubes

43 were sealed securely and submerged in a 65°C bath for 90 minutes. The tubes were then removed and allowed to cool slowly to room temperature. 30 uJ of the following reaction mix (per sample: 0.9 ul 1 M Tris-HCI, pH 8.3; 0.9 uJ 0.5 M MgCl2, 0.25 uJ 1 M DTT, 6.75 |il 1 mg/ml actinomycin D, 1.33 uJ 5 mM 4 dNTP mix, 20 uJ H2O, and 0.2 ul 25 U/|il Avian myoblastosis virus (AMV) reverse transcriptase (Gibco BRL)) was added to each tube containing RNA and oligonucleotide and incubated 1 hour at 42°C. 105 |il RNase reaction mix was added to each primer extension reaction tube and incubated 15 minutes at 37°C. After adding 15 u,l of 3 M sodium acetate (pH 7.0), proteins were extracted with 150 \i\ phenol/chloroform/isoamyl alcohol (25:24:1), and the aqueous phase was transferred to a fresh tube. DNA was precipitated by adding 375 u,l of ice-cold 100% ethanol and washed with 500 uJ of 70% ethanol. The pellet was air-dried with the cap open for 5 to 10 minutes and resuspended in 10 uJ stop/loading dye. The tubes were heated for 2 minutes at 95°C and 3 |il loaded on a sequencing gel. The dideoxy sequencing reaction using the same primer was run alongside the primer extension products.

8. RNA isolation

Total RNA was isolated from P. gingivalis and E. coli strains with TRIzol Reagent (Gibco BRL, Gaithersburg, MD) according to the protocol described by the manufacturer with modifications. Briefly, bacterial cells grown in desired growth media to logarithmic phase (unless stated otherwise) were pelleted by centrifugation. 5 ml TRIzol Reagent were added to 100 ml culture and the cells were homogenized by repetitive pipetting. The homogenized samples were incubated for 5 minutes at room temperature to permit the complete dissociation of nucleoprotein complexes. 0.2 ml of chloroform was added per 1 ml of TRIzol Reagent used. The tubes were capped securely and shaken vigorously by hand for 15 seconds and incubated at room temperature for 2 to 3 minutes. The samples were centrifuged at no more than 12,000 x g for 15 minutes at 4°C. The RNA pellet was washed with 75% ethanol, using a

44 minimum of 1 ml of 75% ethanol per 1 ml of TRIzol Reagent used in the initial homogenization. The sample was mixed by vortexing and centrifuged at no more than 7,500 x g for 5 minutes at 4°C. The RNA pellet was briefly air dried and resuspended in deionized distilled H2O.

9. Northern blot Northern blot analysis was performed as described by Ausubel et al. (8) with modifications. Briefly, equal amounts of total RNA (10-20 u.g) were separated by electrophoresis using 1% agarose-formaldehyde gel. A duplicate gel was stained with ethidium bromide to check the intensity of rRNA bands . After rinsing the gel in dH20 treated with diethylpyrocarbonate to remove formaldehyde, RNA was transferred onto a nitrocellulose membrane. Procedures for pre-hybridization, hybridization, washes and development of the blots were similar to that of Southern blot except the hybridization temperature was 10°C higher than in the Southern blot and all the reagents and containers were RNase free. A 0.5 kb Hinc\\-Pst\ 5' fragment or a 0.7 kb

Pst\-Kpn\ internal fragment of the tpr gene were labeled with biotin by following the nick translation protocols described by the manufacturer (BluGENE™, Nonradioactive Nucleic Detection System, Gibco, BRL, Gaithsburg, MD) and used as the probes to detect fprmRNA.

10. Construction of tprr.lacZ reporter vectors and selection of P. gingivalis transconjugants

The construction of tpr::lacZ reporter shuttle vector pNTX-400 is shown in Figure 5. The construction of suicide shuttle vector pBYZ is shown in Figure 6. Shuttle vector pBYZ and pNTX series plasmids were introduced from E. coli into P. gingivalis by conjugation as described below. To mobilize shuttle plasmids from E. coli to P. gingivalis, the helper plasmid R751 was introduced into the E. coli strain containing shuttle vectors by E. coli-E. coli mating.

45 Figure 5. Construction of tprr.lacZ reporter shuttle vector pNTX-400.

46 Ligation

BamH\

Figure 6. Construction of suicide shuttle vector pBYZ.

47 Transconjugants containing both R751 and shuttle vector were isolated by streaking the mating mixture on LB agar plates containing 200 u.g/ml trimethoprim (to select R751) and 50 |ig/ml kanamycin (to select the shuttle vector). Shuttle vectors were then introduced from the E. coli donor into P. gingivalis by conjugal mating as follows. Cultures of E. coli XL-1 strains containing R751 and the shuttle vector and P. gingivalis W83 were grown to OD600 of 0.2. 0.2 ml of the E. coli donor were mixed with 1.0 ml of the P. gingivalis recipient in a sterile Eppendorf tube. After centrifugation, the pellet was resuspended in 0.2 ml TYE broth. The resuspended mixture was placed on Millipore HAWP filters (Millipore Co. Bedford, MA) on BHI-blood agar plates. The plates were incubated aerobically at 37°C for 4 hours and then incubated anaerobically for 48 hours. Cells grown on the membrane were resuspended in 3 ml pre-reduced BHI broth and 0.2 ml of the cell suspension was plated on pre-reduced BHI-blood agar containing 200 u.g/ml gentamicin and 10 u.g/ml erythromycin or 10 u,g/ml tetracycline. Gentamicin was used to inhibit growth of E. coli donor cells and erythromycin or tetracycline were used to select for P. gingivalis transconjugants containing a chromosomal integrated Emr gene or a plasmid born Tcr gene which had been introduced into P. gingivalis. The plates were incubated anaerobically at 37°C for up to 4 weeks. Colonies grown on the plates were passaged twice on BHI-blood agar with erythromycin or tetracycline to ensure that they acquired the antibiotic resistance.

11. Southern blot

Restriction digested DNA fragments were separated by 0.8% agarose gel electrophoresis and transferred to nitrocellulose membranes. The membranes were baked at 80°C for 2 hours. After prehybridization for four hours at 42°C in 6X SSC (0.9M sodium acetate, 0.09M sodium citrate, pH7.0) containing 100 u.g/ml denatured salmon-sperm DNA, 0.5% SDS, 0.2% Denhardt's solution, the bound DNA was hybridized to the biotin labeled probes (25 ng/ml) at 55°C for 18 hours. The

48 membranes were washed twice at room temperature in 2X SSC-0.1% SDS, washed twice in 0.2X SSC-0.1% SDS at room temperature, and followed by washing twice in 0.16X SSC-0.1% SDS at 55°C for 15 minutes each. The membranes were incubated for 1 hour at 55°C in 3% BSA in 0.1 M Tris-HCI, 0.15 M NaCI (pH 7.5) and then reacted with streptavidin-alkaline phosphatase (SA-AP) conjugate for 20 minutes. After washing in 0.1 M Tris-HCI buffer (pH 7.5) for 30 min, the membranes were incubated in nitroblue tetrazolium (NBT) and 5-bromo-4-chloro-3-indolylphosphate (BCIP). The incubations were carried out at room temperature until reactive bands appeared. Hybridization bands were detected by the BluGENE Nonradioactive Nucleic Acid Detection System (BluGENE™, BRL, Gaithersburg, MD). The reactions were stopped by washing the membranes in 20 mM Tris-0.5 mM Na2EDTA, pH 7.5.

12. Electrophoresis and gelatin zymography Discontinuous SDS-PAGE was carried out by the method of Laemmli (100). For denaturing bacterial samples under reducing conditions, bacterial cells were added in sample buffer [62.5 mM Tris-HCI (pH 6.8), 5% v/v (3-mercaptoethanol (pTvIE), 1% w/v SDS, 10% v/v glycerol, and 0.012% w/v bromophenol blue] and heated at 95°C for 10 minutes before loaded on a gel. To avoid self-proteolysis in P. gingivalis samples, cells were incubated on ice with 20 mM TLCK for 10 minutes before sample buffer was added. For non-reducing, non-denaturing polyacrylamide gel electrophoresis, samples were prepared in sample buffer as described above except that (3ME and SDS were omitted, and samples were incubated at room temperature instead of heating for 30 min.

SDS-PAGE was carried out using the Mini-PROTEIN II™ vertical electrophoresis system (Bio-Rad Laboratories, Hercules, CA). Proteins were separated on 12% (wt/vol) polyacrylamide resolving gels with 4% (wt/vol) polyacrylamide stacking gels by the method of Ames (7). Electrophoresis was carried out at room temperature at a

49 constant 200V until the dye front reached the bottom of the gels. The gels were stained with Coomassie brilliant blue R-250. Gelatin zymography For gelatin-substrate zymography, the acrylamide in the gels was copolymerized with non-conjugated gelatin at a concentration of 1 mg/ml as described by Bamda et al. (11). After electrophoresis of the non-reduced, non-denatured samples, the gels were soaked in 100 mM Tris-HCI (pH 7.5) buffer containing 2% (v/v) Triton X-100 for 30 min, rinsed twice in water, and then incubated for a further 30 min in 100 mM Tris- HCI (pH 7.5) . The gels were then transferred into the development buffer consisting of 100 mM Tris-HCI (pH 7.5) and 50 mM cysteine. After incubating at 37°C for 2 hours, the gels were stained with Coomassie brilliant blue R-250. After destaining the gels, proteolytic activity was visualized as a clear band against a dark blue background.

13. Western immunoblot Whole cell lysate proteins separated by SDS-PAGE were transferred to nitrocellulose membranes in 25 mM Tris-HCI, 192 mM glycine and 20% methanol buffer (pH 8.3). Transfer was carried out at 60V for 2 hours in a Bio-Rad Trans-Blot™ Cell (Bio-Rad Laboratories, Hercules, CA). After blocking the unreactive sites with BSA, the sheets were washed twice with TTBS (20mM Tris, 500mM NaCI, 0.05%

Tween-20) and then incubated with rabbit antiP. gingivalis antibodies. The unbound antibodies were removed by washing with TTBS. The sheets were then incubated with goat anti-rabbit IgG coupled to alkaline phosphatase for one hour. Antigen- antibody reaction bands were visualized by following the procedures described in the Bio-Rad technical bulletin supplied with the Bio-Rad Immuno-Blot (GAR-AP) assay kit.

14. p-galactosidase assay

Assay of (3-galactosidase activities in E. coli strains was as described (138).

Briefly, E. coli cells were grown in various media to mid-log phase of growth (OD600 of 0.28-0.7). The cultures were cooled to prevent further growth by immersing in ice for

50 20 minutes. The cell density was determined by measuring the absorbance at 600 nm. 0.5 ml of each culture was put in a microcentrifuge tube and the cells pelleted. After discarding the supernatant, the cells were resuspended in 1ml Z buffer (60 mM

Na2HP04-7H20, 40 mM NaH2P04H20, 10 mM KCI, 1 mM MgS04-7H20, and 50 mM pME, pH 7.0). Two drops of chloroform and 1 drop of 0.1% SDS solution were added and the tubes were vortexed for 10 seconds. After placing the tubes in a water both at 28°C for 5 minutes, the reaction was started by adding 0.2 ml of o-nitrophenyl-p-D- galactopyranoside (ONPG) (4 mg/ml) to each tube and shaken for a few seconds. The time of the reaction was recorded by a stop watch. When yellow color had developed in the tubes, the reaction was stopped by adding 0.5 ml of a 1 M Na2CC>3 solution. Optical density reading at both 420 mji and 550 mji were recorded and p- galactosidase activity was calculated as described (138). p-galactosidase activity was expressed in Miller's units. Samples were assayed in duplicate, and the data represented here are the average of at least 3 independent experiments.

To analyze P. gingivalis (3-galactosidase activity, P. gingivalis strains were grown in various nutritional media to the logarithmic growth phase unless stated otherwise. Cells were harvested, washed twice with PBS and incubated in 20 mM TLCK on ice for 10 min. Cells were then resuspended in the reporter lysis buffer and analyzed for p-galactosidase activity as described in the Promega Technical Bulletin of p- galactosidase enzyme assay system with reporter lysis buffer (Promega Co., Madison, Wl). Assays were done in 96 well microplates and the p-galactosidase activity was measured at 405 nm by a microplate reader (Model 3550 Microplate reader, Bio-Rad Laboratories, Richmond, CA). A standard curve for purified p-galactosidase was prepared for each assay. Protein concentration was measured with the Bradford reagent supplied by Bio-Rad (Gibco BRL, Hercules, CA) with BSA as standard. One Unit of p-galactosidase activity was expressed as 1 nanomole ONPG hydrolyzed per

51 min per mg of cellular protein. Samples were assayed in duplicate. The data represented here are the average of at least 3 independent experiments.

15. Stress conditions and hemin limitation

For heat shock and pH change responses, P. gingivalis cells were first grown in 0.5TYE to mid-log phase and the cells were transferred to 42°C or the cells were incubated in 0.5 TYE at pH 5.5 or pH 8.5 for 4 hours before assay. Previous studies showed that heat shock proteins were optimally expressed 4 hours after exposure to the environmental change (118). For hemin starvation, P. gingivalis cells were passaged twice in 0.5 TYE without hemin.

52 III. RESULTS

1. Identification of proteolytic clones

A P. gingivalis W50 DNA library was generated by cloning Sau3M partially digested DNA into BamH\ digested plasmid pTZ18R. 4,000 white colonies were isolated. Plasmids isolated from several random clones were analyzed with restriction endonucleases on an agarose gel and shown to contain DNA inserts in the range of 1-

12 kb. Recombinant E. coli clones were plated on LB agar plates supplemented with ampicillin (50 u.g/ml) and skim milk (1.5%). After overnight incubation aerobically, they were lysed and transferred to an anaerobic chamber and incubated for up to one week. Two colonies showed clearing zones around the colonies after 4 days of anaerobic incubation. When the cells were re-streaked on LB agar plates with 1.5% skim milk, they consistently showed the ability to hydrolyze skim milk when incubated anaerobically. These two clones were designated TZ18-1 and TZ18-2 (Figure 7).

2. Analysis of proteolytic clone TZ18-1 Plasmids pTZ18-1 and pTZ18-2 from clones TZ18-1 and TZ18-2 respectively were isolated and restriction analyzed (Figure 8). There was some kind of rearrangement in the vector plasmid pTZ18R by restriction analysis since when digested with EcoRI, pTZ18R was approximately 2.9 kb (lane 2), but this band was not detected in EcoRI digested pTZ18-1 (lane 3) or pTZ18-2 (lane 4). The result showed that pTZ18-1 and pTZ18-2 were identical plasmids, so only TZ18-1 was analyzed further. This plasmid provided ampicillin resistance and replicated well in E. coli XL-1 blue cells. The restriction map of plasmid pTZ18-1 is shown in Figure 9. A restriction map of the DNA insert suggested the insert size was approximately 11 kb and a portion of it was very similar to that of the tpr clone pYS307 (169). Southern blot analysis using a 1.0 kb

Hinc\\-Kpn\ internal fragment of tpr from pYS307 as probe proved there was shared high homology on the DNA insert to that of tpr (Figure 10). When pTZ18-1 was restriction digested with EcoRI or Hind\\\, a 4.3 kb or a 3.5 kb reactive band was

53 Figure 7. Proteolytic activity of clone TZ18-1 and TZ18-2 on skim milk agar plate. 1, 2 and 3 represent E. coli XL-1/pTZ18R, clone TZ18-1 and TZ18-2, respectively. The plate was incubated aerobically at 37°C overnight, treated with chloroform, and then incubated anaerobically at 37°C for two days.

54 1 2 3 4 5 6 7

23.7

9.50

6.66

4.26

2.25 1.96

Figure 8. Restriction analysis of plasmids pTZ18-1 and pTZ18-2. Lanes: 1, X DNA digested with Hind\\\; 2, 3 and 4, EcoRI digested plasmid pTZ18R, pTZ18-1 and pTZ18- 2, respectively; 5, 6 and 7, Hind\\\ digested plasmid pTZ18R, pTZ18-1 and pTZ18-2, respectively.

55 Sa He Sm Ec HcPsSpKp Ec Ec

1 P1218-1 — I I lllf» I 1 tpr

Bm Ec HcPsSpKp Ec

pYS307 I 1 Mil) 1 tpr

1 kb

Figure 9. Partial restriction map of pTZ18-1 and pYS307. The lines represent the DNA fragments cloned into pTZ18R. The possible location and direction of tpr gene on the DNA insert is indicated by a solid arrow line. Abbreviations are: He, HincW; Sa, Sad; Sm, Sma\; Ec, EcoR\; Ps, Pst\; Sp, Sph\; Kp, Kpn\; Bm, BamH\. The scale of the DNA is indicated as 1 kb.

56 A B 1 2 3 4 1 2 3 4

Figure 10. Southern blot analysis of pTZ18-1 with a tpr probe. A, agarose gel showing restriction fragments of DNA. B, Southern blot. A 1.0 kb Hinc\\-Kpn\ internal fragment of tpr from pYS307 was used as the probe. Lanes: 1, EcoRI digested pTZ18R; 2, EcoRI digested pTZ18-1; 3, Hind\\\ digested pTZ18-1; 4, Kpn\ and HincW digested pTZ18-1. kb, DNA size marker in kilo bases. The position and size of reacting DNA bands are indicated.

57 detected, respectively. When Both HincW and Kpn\ were used to digest pTZ18-1, a 1.0 kb reactive band was seen. These results were in agreement with the restriction pattern of the tpr clone.

The identity of the cloned P. gingivalis protein was determined by immunoblotting with the antisera raised against P. gingivalis W50 whole cell and P. gingivalis W83 membranes. A single immunoreactive protein band with apparent molecular weight of 90 kDa was detected in TZ18-1 whole cell lysate (Figure 11). No immunoreactive band was detected in the E. coli XL- 1/pTZ18R control. When preimmune serum was used as the primary antibody, no immunoreactive band was seen in TZ18-1 or E. coli

XL-1/pTZ18R. This indicated that the cloned P. gingivalis DNA was the source of the immunoreactive 90 kDa protein. This protein also reacted with anti-recombinant Tpr antibodies, though to a lesser extent as compared to the anti-P. gingivalis antibodies (Data not shown). This can be explained by the weak reactivity of the anti- recombinant Tpr antibodies to E. coli JM83/pYS307 which expressed the recombinant Tpr that was used to generate these antibodies. The proteolytic clone TZ18-1 expressed such strong proteolytic activity that its whole cell lysate was devoid of high molecular weight protein bands as analyzed on a SDS-PAGE (Figure 12A). The cellular proteins were probably degraded by the recombinant protease. Expression of the protease activity was demonstrated by gelatin zymography (Figure 12B). A gelatinolytic band of approximately 90 kDa was detected in TZ18-1 but absent in E. co7/XL-1/pTZ18R. This proteolytic band was of the same molecular mass as seen in the Western blot of TZ18-1 probed with antisera raised against P. gingivalis (Figure 11).

Taken together, the data show that the tpr protease gene from P. gingivalis W50 had been cloned. Since this gene had already been cloned, sequenced and partially characterized, I went on to investigate the regulation of its expression.

58 Figure 11. Western immunoblot of TZ18-1 with antisera against P. gingivalis W50 and W83. A, probed with anti-P. gingivalis W50 whole cell antibodies. B, reacted with anti- P. gingivalis W83 membrane antibodies. Lanes: 1, prestained high molecular weight marker; 2 and 3, non-reduced and non-denatured whole cell lysates of clone TZ18-1 and E. coli XL- 1/pTZ18R, respectively. kDa, molecular weight markers in kilo daltons. The position and size of a prominent reacting band is indicated.

59 A B

12 12

Figure 12. SDS-PAGE and gelatin zymogram of clone TZ18-1. Samples for SDS- PAGE (A) were reduced and denatured. Samples for gelatin zymogram (B) were non- reduced and non-denatured (see material and methods). kDa, molecular weight markers. Lanes: 1, whole cell lysate of E. coli XL-1/pTZ18R; 2, whole cell lysate of clone TZ18-1. The position and molecular weight of a gelatinolytic band is indicated.

60 3. The effect of nutrient limitation on tpr transcription Previously we observed that Tpr proteolytic activity (Pz-peptidase) was influenced by growth conditions (166). P. gingivalis W83 cells grown in 0.5 TYE had as much as five times Pz-peptidase activity compared to cells grown in BHI, and two times the activity of cells grown in TYE. To determine if the differences in activity were reflected in levels of transcription, the expression of tpr was examined by Northern hybridization. The DNA probe was the 0.7-kb Pst\/Kpn\ fragment of the tpr gene which had been replaced by the erythromycin resistant gene (Emr) in the isogenic mutant, W83/PM (168). The results paralleled that of the enzymatic analysis. Cells grown in 0.5TYE had the highest amount of tpr transcript, there was less tpr mRNA in cells grown in TYE broth and no transcript was detected in cells grown in BHI (Figure 13A). As expected, there was no reacting band in RNA from the isogenic mutant W83/PM. The Northern blot was analyzed by densitometry scanning and showed the tpr mRNA band in 0.5TYE grown cells was more than 5 times that of TYE and 25 times that of BHI (Figure 13B). The effect of BSA and gelatin supplementation to 0.5TYE was also investigated (Figure 14). BSA and gelatin suppressed tpr expression. To investigate whether the isogenic mutant expressed the 5' region of tpr mRNA, the 0.5-kb Hinc\\/Pst\ fragment of tpr was used as the DNA probe in Northern analysis (Figure 15). The result showed there was no tpr transcripts in W83/PM. This suggested either the truncated tpr gene was not transcribed or its transcript was very unstable due to the short size and was not detected in our Northern assay.

Northern blot analysis with RNA isolated from P. gingivalis ATCC33277 grown in BHI, TYE or 0.5TYE did not show any tpr mRNA transcripts (Figure 16). This suggests strain ATCC33277 may not express fprorthat tpr\n ATCC33277 does not share high homology to the 0.7 kb Pst\-Kpn\ probe. This is in agreement with a Southern blot analysis of the tpr gene which suggested the tpr gene in P. gingivalis ATCC33277 was

61 Figure 13. A, Northern blot analysis of tpr mRNA from W83 grown in BHI, TYE, 0.5TYE and W83/PM grown in TYE. B, Densitometry scan of the blot. 20 u.g of total RNA was loaded on each lane. A 0.7 kb Kpn\-Pst\ internal fragment of the tpr gene was used as the probe, kb, RNA size markers in kilo bases. The position and size of a single reacting band is indicated.

62 Figure 14. Northern blot analysis of tpr mRNA expression in P. gingivalis grown in different nutritional conditions. 10 uig of total RNA was loaded on each lane. A, formaldehyde agarose gel. The position of 23S and 16S ribosomal RNAs are indicated. B, Northern blot. A 0.7 kb Pst\-Kpn\ internal fragment of the tpr gene was used as the probe. Lanes: 1, 2, and 3, RNA isolated from P. gingivalis W83 cells grown in BHI, TYE, and 0.5TYE, respectively; 4 and 5, RNA isolated from W83 cells grown in 0.5TYE supplemented with 0.25% BSA and 0.25% gelatin, respectively. The position of tpr mRNA is indicated.

63 Figure 15. Northern blot analysis of tpr mRNA expression in P. gingivalis W83 and W83/PM. 10 jo.g total RNA was loaded on each lane. A, formaldehyde agarose gel. The position of 23S and 16S ribosomal RNAs are indicated. B, Northern blot. A 0.5 kb Hinc\\-Pst\ 5' fragment of the tpr gene was used as the probe. Lanes: 1, 2, and 3, RNA isolated from P. gingivalis W83 cells grown in BHI, TYE, and 0.5TYE, respectively; 4, 5, and 6, RNA isolated from W83/PM cells grown in BHI, TYE, and 0.5TYE, respectively. The position of tpr mRNA is indicated.

64 Figure 16. Northern blot analysis of tpr mRNA expression in P. gingivalis ATCC33277. 10 u,g total RNA was loaded on each lane. A, formaldehyde agarose gel. The position of 23S and 16S ribosomal RNAs are indicated. B, Northern blot. A 0.7 kb Pst\-Kpn\ internal fragment of the tpr gene was used as the probe. Lanes: 1, 2, and 3, RNA isolated from P. gingivalis W83 cells grown in BHI, TYE, and 0.5TYE, respectively; 4, 5, and 6, RNA isolated from P. gingivalis ATCC33277 cells grown in BHI, TYE, and 0.5TYE, respectively. The position of tpr mRNA is indicated.

65 different than that in P. gingivalis W50 or W83 strains (169). The transcription product of the tpr gene is approximately 1.7 kb. Since the coding region of tpr is 1.45 kb, it suggests tpr is transcribed monocistronically. The effect of growth stage on tpr expression was analyzed by comparing tpr mRNA levels of P. gingivalis W83 cells grown in TYE or 0.5TYE to mid-log or early stationary phase of growth. The result indicated that tpr was expressed more at stationary phase than log phase (Figure 17).

4. Analysis of tpr expression by a tpr::lacZ reporter construct. To facilitate the analysis of tpr expression, we constructed a tprr.lacZ reporter shuttle plasmid pNTX-400 (Figure 5 and Table 5) which carried the 612-bp upstream

region and 183-bp of the coding sequence of tpr fused with a promoterless lacZ gene.

The lacZ gene lacked the first 8 codons and could not be expressed by itself. The 5'

end of the lacZ gene was fused inframe to the coding sequence of tpr to bring the

expression of lacZ under control by the tpr promoter. The plasmid pNTX-400 was

introduced into P. gingivalis W83 by conjugation. The transconjugant showed (3- galactosidase activity on a TYE agar plate containing 50 |ig/ml X-gal (Figure 18),

whereas the control P. gingivalis W83/pNJR12 had no activity. (3-galactosidase activity expressed in W83/pNTX-400 matched the profile of tpr expression (Figure 19). In

nutrient rich medium, BHI, lacZ expression was repressed compared to nutrient poor

medium 0.5TYE. Hence, the plasmid-borne tprr.lacZ fusion was regulated like the chromosomal tpr gene and its regulation mainly occurred at the transcriptional level. It

also indicated that the 795-bp BamH\-Pst\ fragment carried all necessary information required for regulation of tpr expression.

One concern is that the plasmid pNTX-400 existed in P. gingivalis in multiple

copies, which is different from the situation that tpr exists in P. gingivalis as a single copy gene. So a suicide shuttle vector pBYZ (Figure 6) was constructed and

introduced into P. gingivalis W83 by conjugation. Since this plasmid can not replicate

66 Figure 17. tpr mRNA expression at different growth stages. 15 u.g total RNA was loaded on each lane. A, formaldehyde agarose gel. The position of 23S and 16S ribosomal RNAs are indicated. B, Northern blot. A 0.7 kb Pst\-Kpn\ internal fragment of the tpr gene was used as the probe. Lanes: 1 and 2, RNA isolated from log phase W83 cells grown in TYE and 0.5TYE, respectively; 3 and 4, RNA isolated from early stationary phase W83 cells grown in TYE and 0.5TYE, respectively. The position of tpr mRNA is indicated.

67 Figure 18. p-galactosidase activity in P. gingivalis W83/pNTX-400 lysates. P. gingivalis W83/pNJR12 (A) and W83/pNTX-400 (B) were grown in BHI broth to mid-log phase, the cells were lysed and spotted on a TYE agar plate containing 40 u.g/ml X-gal and incubated at 37°C overnight. Blue colour indicates p-galactosidase activity.

6H 250

200 A

150 A

"S loo A

50 H

Figure 19. Effect of growth nutrient on lacZ expression in P. gingivalis W83/pNTX-400. Cells were grown in BHI, TYE or 0.5TYE. W83*, P. gingivalis W83 grown in TYE and served as the negative control. One Unit of (3-galatosidase activity equals 1 nanomole ONPG hydrolyzed per minute per mg of cellular protein.

69 in P. gingivalis, the tprr.lacZ construct can only exist on P. gingivalis chromosome by homologous recombination between pBYZ and chromosomal DNA. The result can be either duplication of tpr (single crossover event) or allelic exchange of the wild-type tpr

gene with tprr.lacZ reporter gene (double crossover event) (Figure 20). A total of 75 transconjugants grew on BHI blood agar plates supplemented with gentamicin and erythromycin. Preliminary Southern analysis indicated they all had

lacZ genes integrated on their chromosomes. Southern analysis of several clones indicated that both single and double crossover constructs were obtained (Figure 21

and Table 7). When P. gingivalis chromosomal DNA was digested with BamH\ and probed with a 0.8 kb BamHI-Psfl fragment of tpr, a single 9.5 kb reacting band was detected in clone 29, 55 and 64, indicating an allelic exchange event had occurred that resulted in the replacement of the tpr gene with a tprr.lacZ reporter gene. In clone 17 and 65, the tpr probe hybridized to 18.5 kb and 3.5 kb or 12.7 kb and 9.5 kb bands, respectively, indicating duplication of tpr. The ratio of double cross-over events to single cross-over events was approximately 1 to 28. p-galactosidase activity assays done on these mutants showed both single and double cross-over mutants had the same tpr regulation as in

the wild-type strain (Figure 22), indicating that a single copy tprr.lacZ reporter gene construct could be used to analyze tpr expression. This also suggested that the expression of tpr does not need an intact Tpr gene product.

5. Primer extension and sequence analysis of the promoter region of tpr The finding that tpr is regulated at the transcriptional level prompted us to analyze the 5' non-coding region of this gene. The nucleotide sequence of the 5' region of the

tpr locus of P. gingivalis W83 was determined (Figure 23). The complete sequence was deposited in Genbank (accession number AF022499). A set of putative -10 and -35 elements, suggested by deletion and primer extension analysis (see below), were labeled. Computer analysis of the region upstream of the

70 BamHI BamHl P. gingivalis chromosome DNA 3.5kb \ crossover 1 J \ crossover 2 /I W eamrff I V////A ,acZ

probe tpr pBYZ probe lacZ

18.5kb

Double crossover

BamHI BamH\ V///A la~cZ Em' YZZZA 9.5kb

Single crossover 1

BamHI BamHI BamHI lacZ Err/ V///X V.V////////////A 18.5kb 3.5kb

Single crossover 2 BamHI BamHI BamHI V/////////////A // &7777\ lacl 12.7kb 9.5kb

Figure 20. Possible homologous recombinations between pBYZ and P. gingivalis W83 chromosone. The hatched boxes represent the DNA fragment containing the fprgene, the blank boxes represent the lacZ gene and the Emrgene respectively. The number in kb below each construct indicates the size of that fragment digested with BamHI.

71 1 2 3 4 5 6 7

— 18.5 — 9.5

Figure 21. Southern blot analysis of chromosomal DNA from P. gingivalis W83 strains. A, Southern blot probed with a 0.8 kb BamH\-Psfi 5" fragment of tpr gene. B, Southern blot probed with a 3.5 kb lacZ gene. All DNA were digested by BamH\. Lanes: 1, plasmid pBYZ; 2, chromosomal DNA from P. gingivalis W83; 3, 4, 5, 6, and 7, chromosomal DNA from clones 29, 17, 55, 64 and 65, respectively. The position and size of reactive bands are indicated.

72 Table 7. Southern blot analysis of BamHI digested chromosomal DNA from P. gingivalis W83 transconjugants.

Expected bands (kb) Observed bands0 (kb) Probe3 pBYZ W83 DC SC1 SC2 pBYZ W83 DC SC1 SC2 tpr 18.5 3.5 9.5 18.5 12.7 18.5 3.5 9.5 18.5 12.7 +3.5 +9.5 +3.5 +9.5 lacL 18.5 - 9.5 18.5 9.5 18.5 - 9.5 18.5 9.5 a. A 0.8 kb BamHI-Psfl fragment of tpr was used as the tpr probe. A3.5kbBamHI- Hind\\\ lacZ gene was used as the lacL probe. b. DC (double crossover) represents clones 29, 55 and 64. SC1 (single crossover 1) represents clone 17. SC2 (single crossover 2) represents clone 65.

73 50

40 H

30 A

3 20 -I c P 10 J

-10

U u U o 00 Q Q p—i w K oa >< H H H © © ©

Figure 22. p-galactosidase expression in P. gingivalis transconjugants. P. gingivalis W83, single crossover (SC) transconjugant and double crossover (DC) transconjugant were grown in BHI or 0.5TYE. One Unit of p-galactosidase activity equals 1 nanomole of ONPG hydrolyzed per minute per mg of cellular protein.

74 l GGATCCTGCTCCCATAGCATCATCTTGTGATCCCAAT BamHI 38 GATCTTCAAGCCACTGACATAGCAACTCTTTGATAAC 75 AC C GAAATC GAC TACAC GC C C GATC GAATCAAGAGC C 112 GGTGCACTGCAAATAAAGTGGATACGATAGTTGTGTC 149 CATGCAGATGTCTGCACTTGTGATCGTGCCCCACTAC 186 GCGGTGTCCCATGCTAATGTCGTGATAACGTTCGGCT 223 GTAATCATATATTGTCAAAGCT/AAGTTTCTGCTGTTC 260 AAAGGTAATCAAATAATCCCCTGCTTCACGTATTCGG 297 CTGTTCGGGTCTCGTCTGAAAAACATTTTTGTAATTT -35 334 TTTGAAACACTCTTCAGACACACATTTTCAGGTGAAA -10 +1 371 ATTGTCAATTTTTGCTCTTGCACTCGTAACTGATTGA

408 AATAATGGCTGTTAACGGTTTTTCATGCGTGATTTGT

445 CCAAATTGCACCTTAATTCGTTAGTTTTTTTGTACAC > 482 AAATGCGTGATTTGTCCAAGAAAATGCGTGATTTGTC >

519 C7AAGTCTTTTCCTT7^ATATATCCCTTCATTTGTGTGA - - > 556 CTCTTAAAGTGTAAGAGCCTAAAGTCACAGTTTTAAT

593 CAATCTAAAATTTTTCAATTATG Met

Figure 23. Nucleotide sequence of the 5' region of tpr gene from P. gingivalis W83. Putative-35 and-10 promoter regions are indicated based on primer extension analysis. 3 direct repeats of 17 bp are indicated by arrows. +1 indicates the transcriptional start site in P. gingivalis W83. * indicates the transcriptional start site in E. coli. Met indicates the translation start site of Tpr.

75 transcription start sites found no consensus sequences to the -35 and -10 regions of E. coli promoters. Interestingly, the region between the putative promoter elements and the tpr coding region contains 3 identical direct repeats of 17 bp (Figure 23, indicated by arrows). The first repeat is part of a short open reading frame of 19 codons and the other two repeats are parts of one short open reading frame of 16 codons. We mapped the start site of tpr mRNA by primer extension analysis on total RNA isolated from P. gingivalis W83 grown in 0.5 TYE. First we used primer tpr293 which lies within the coding region of tpr based on the promoter region suggested by Bourgeau et al. (16). Our primer extension product was so large that we estimated it to be more than 200 bp upstream of the coding region. By using primer tpr64, we identified the transcriptional start site to be 215 bps upstream of the coding region of tpr at two A residues (Figure 24). The primer extension product could be seen in P. gingivalis cells grown in 0.5 TYE but not from cells grown in BHI. This is in agreement with the results of Northern blot that showed P. gingivalis grown BHI had no detectable tpr mRNA. The putative -35 region is similar to that of o70 promoters but the -10 region does not resemble the Pribnow box of o70 promoters.

6. Molecular analysis of tpr 5' region by deletion mutations

To analyze the regulatory sequence in the tpr 5" region, we generated several deletion fragments of the tpr promoter region by PCR. Plasmids containing these tpr deletion constructs were introduced into P. gingivalis and (3-galactosidase activity analyzed in the transconjugants (Figure 25). Deletion of the putative -35 region of tpr promoter in pNTX-30 resulted in elimination of (3-galactosidase expression. Further deletions downstream in pNTX20, pNTX52 and pNTX137 also resulted in loss of (3- galactosidase expression. The residual (3-galactosidase activity was not affected by nutritional conditions. The promoter-less lacZ gene in pNTX expressed minimal (3- galactosidase activity. Plasmids pNTX-400 and pNTX-100 which retained the

76 Figure 24. Primer extension analysis of tpr transcription in P. gingivalis W83. A, G, T and C are abbreviations for adenosine, guanosine, thymidine and cytidine, respectively. PE, primer extension. Oligonucleotide tpr64 was used as the primer. The positions of primer extension products are indicated as two A-T base pairs.

77 • o o -o o

OJ on OJ OJ •00 -OJ on on o o O O m ± 2 O CD CQ o o O o CD CO cz + + » Q. + + + 2O. E Bl o^ ' CD O O ZJ 03 ro ro ZJ r* cn ro Q- CD o o ro CD to to "o Q. 03 TZO CO CL to , ZJ CD A T3 CD Q. o H —I o Q. 2 X X X X OJ i x I i o =>' CD CQ on ro OJ 5" o o OJ ro o o oJ> 3 <9. 3' o o o —1 "D CD O CQ > o o sz —I ZJ o N 03 ZJ to I CD ZJ 03 Co" ZD 03 to on o 1 I ?. ZJ o o O on on — 00 on CL cr ZJ ro 3 CD CD to on CO o o /—N —V s > m CD CD CL Pi ro 1 —1 ro —Ji OJ ro cn o O o On cn O i. a o bo O CD ro O ZJ c± CD •—' £CO 0^ 3 -CIL £ to ' jcT • cz f—•- 03 OJ ro ro r- |w 03_ ZJ _ < p p o o O ro OJ 03 a CL J> OJ CD 03 ro bo on CD —\ o o to S -co o o —» 00 00 cn —» o 03 o" OJ ro CD O ZJ °- CQ *— •—- ^— CL 00 ro to < 0y3. w " ' •—• SL o CL 01 O if OJ N to to CD O CL CL to a ZJ 03 CD 5" 03 5' to 8.9 ( > OJ .~0 cr _ R, o J> O o ro o CD bo ro o o p OJ ^—s p —s o Q <'

78 promoter region of tpr expressed (3-galactosidase activity and were regulated by the nutritional conditions. The regulatory pattern corresponded well to the expression of tpr in the wild-type strain. This result supported the primer extension analysis showing that the tpr promoter was 215 bp upstream of the coding region. The effect of deleting the three direct repeats in the 5' region of fprwas shown in pNTX-100A. In this case there was expression of (3-galactosidase activity but the expression was not regulated by nutritional status as was the case for the parent strain. Cells grown in 0.5TYE had only about 1.5 fold (3-galactosidase activity compared to cells grown in BHI. This suggested the three direct repeats were directly involved in tpr regulation.

The effect of deletion mutations in tprr.lacZ reporter gene constructs were also analyzed in E. coll XL-1 cells. As shown in Figure 25, E. coll cells expressed (3- galactosidase activity in pTXZ19R-400, pTXZ19R-100, pTXZ19R-30, pTXZ19R20 and pTXZ19R52, but not in pTXZ19R137 or pTXZ19R. This expression was not enhanced by the addition of IPTG nor was it affected by the orientation of the insert (these constructs were also made in pTZ18R). Generally, cells grown in Minimum A medium had 2-3 times (3-galactosidase activity compared to cells grown in LB broth. This regulation was different from that in P. gingivalis W83. To clarify the discrepancy, primer extension analysis was conducted to see if there was a second promoter recognized by E. coli on the tpr sequence. Primer extension analysis (Figure 26) of

RNA from E. coli XL-1 /pTXZ19R-400 indicated the transcription started at a C residue 68 nucleotides upstream of tpr translation start codon. There was high homology of the putative -10 region to E. coli promoter -10 region and the putative -35 region lies within the last direct repeat. The deletions into the repeats abolished lacZ expression in E. coli. This suggested E. coli recognized a different promoter region of fprthan P. gingivalis. This also explained the discrepancy of (3-galactosidase expression in P. gingivalis and in E. coli.

79 A G TC1 2

Figure 26. Primer extension analysis of tpr transcription in E. coli XL-1 /pTXZ19-400. A, G, T and C are abbreviations for adenosine, guanosine, thymidine and cytidine, respectively. 1 and 2, primer extension of RNA from E. coli grown in LB and Minimum A media, respectively. Oligonucleotide tpr170 was used as the primer. The position of primer extension product is indicated as a C-G base pair.

80 7. Growth factors that regulate tpr expression

By using the tprr.lacZ reporter gene, we were able to analyze tpr expression under various nutrient growth conditions. An allelic exchange P. gingivalis mutant, clone 55, was named P. gingivalis W83//acZ and analyzed in the following assays. In a time course study (Figure 27), P. gingivalis W83//acZ expressed the highest (3- galactosidase activity when grown in 0.5TYE in stationary growth phase. (3- galactosidase activity was repressed when grown in BHI, TYE, or 0.5TYE supplemented with BSA or gelatin throughout the growth. Of interest was the observation that addition of Casamino acids to 0.5TYE repressed lacZ expression during log phase, but repression was overcome during stationary phase. To analyze whether control of the tpr expression is related specifically to the presence of free amino acids in the medium, P. gingivalis W83//acZ was grown in BHI broth to mid-log phase (OD660=0-5) and resuspended in pre-reduced 0.5TYE or 0.5TYE supplemented with individual amino acids. After overnight growth, the cells were harvested and (3- galactosidase activity determined (Figure 28). Addition of 5 mM free amino acids to 0.5 TYE did not significantly reduce the [3-galactosidase expression. This result and the effect of Casamino acids supplementation suggested that small peptides rather than free amino acids were involved in the regulation of tpr expression.

Since BHI greatly repressed lacZ expression, we fractionated BHI by molecular size on a Sephadex G-10 column and analyzed individual fractions for their ability to suppress tpr expression. BHI fractions of low molecular weight (<700 dalton) suppressed tpr expression to various degrees (data not shown). We analyzed the amino acid composition of several of the BHI fractions, and found the fraction rich in phenylalanine, proline and alanine had the most repressive effect on tpr expression.

The effect of a number of peptides and other chemicals on lacZ expression is shown in

Figure 29. The di-peptide phenylalanyl-phenylalanine repressed lacZ expression 10- fold compared to cells grown in 0.5TYE without supplementation. Di-

81 OD660

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83 80

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0 T I T i ea o 0> u £ Two o o 1/1 Z U Q. I Q. a. Q. • • H • • z es 6 et 6 et o o Z u "OJD u CS u o. I i a. Q. K z 3 Q. in o • a Z o 3 CS z

Figure 29. Effect of peptides and some other chemicals on lacZ expression in P. gingivalis W83//acZ. Cells were grown in BHI broth to mid-log phase and resuspended in 0.5TYE or 0.5TYE supplemented with 5 mM peptides or 40 mM of the other chemicals,incubated overnight and p-galactosidase activity analyzed. One Unit of p-galactosidase activity equals 1 nanomole ONPG hydrolyzed per minute per mg of cellular protein.

84 peptides phenylalanyl-alanine and alanyl-proline also repressed lacZ expression but to a lesser extent. NH4-acetate, (NH4)2S04, NH4NO3, sucrose succinate-Na and

NaCI addition to 40 mM did not greatly influence lacZ expression.

8. Effect of other environmental conditions on tpr expression To analyze whether tpr expression was regulated by heat shock or other stress conditions, (3-galactosidase activity was measured in heat shocked cells, cells incubated at pH 5.5 and pH 8.5, or hemin starved cells. Under these stress conditions, the growth of P. gingivalis is affected to various extents (118, 133, 134). Heat shock at 42°C for 4 hours did not affect cell growth, whereas the pH change resulted in loss of viability. Hemin starved cells also stopped growing (183). The results suggested tpr expression was not affected by pH changes or hemin starvation, its expression was repressed by heat shock to a small extent (Figure 30).

85 125

IOOH

50 H

25 ^

i c u o £ s •o Q.

Figure 30. Effect of hemin limitation, heat shock and pH on p-galactosidase expression i n P. gingivalis W83//acZ. Cells were grown in 0.5TYE, 0.5TYE without hemin supplementation or incubated in 0.5TYE at 42°C, pH 5.5 or pH 8.5 for 4 hours. One Unit of p-galactosidase activity equals 1 nanomole ONPG hydrolyzed per minute per mg of cellular protein.

86 IV. DISCUSSION

In this investigation, a protease gene was cloned from P. gingivalis W50. The nature of the proteolytic activity, size of the recombinant protein, immunoblot analysis and Southern blot analysis, indicated that it was the tpr protease gene. The recombinant proteinase has similar characteristics to the product of the cloned tpr gene of P. gingivalis W83 (169), though it reacted more strongly to antibodies against

P. gingivalis whole cell and membrane. There was rearrangement in the plasmid DNA as indicated by the altered restriction pattern of the vector. The reason is unclear.

Since the auto-proteolytic activity is very strong in the E. coli clone, one hypothesis is that the bacterial cells rearranged the plasmid to decrease the expression of the protease to ensure its survival. Another possibility is that there was modification of the clone during DNA manipulations.

Although there have been numerous reports that P. gingivalis proteases are regulated by the growth environment (23, 126, 133, 184), little is known about how this regulation is achieved at the genetic level. In this study, we made a first attempt to address this question. The finding that tpr proteolytic activity (Pz-peptidase) was influenced by nutritional growth conditions made it an interesting subject of investigation in gene regulation (170). Northern blot analysis indicated that tpr expression was regulated by nutritional growth conditions and this regulation occurred at the transcriptional level. P. gingivalis W83 cells grown in nutrient limited medium (0.5TYE) produced more tpr mRNA than cells grown in nutrient rich medium (BHI). Two kinds of regulation could be responsible: repression or induction. Our data suggested the former because tpr expression was repressed when nutrients were added to the nutrient limited medium.

A variety of nutrient sources, including BHI, TYE, BSA, gelatin, and to a lesser degree, Casamino acids, repressed tpr expression. These nutrients share the common property of being rich in proteins and peptides. Since P. gingivalis posesses

87 a large number of proteases and has the ability to degrade proteins to short peptides and transport them into the cells for metabolism, it is not surprising that the expression of certain proteases is regulated by the availability of nutrients. Our results suggested none of the 20 amino acids had a significant effect on tpr expression. In a time course growth and reporter lacZ expression study, Casamino acids supplementation repressed lacZ expression in log growth phase, but this repression was abolished at stationary growth phase. Since Casamino acids is an acid hydrolysate of casein in which free amino acids and small peptides are present in a ratio of 82 to 18%, respectively (according to the manufacturer), it suggests that the repressive effect may due to the small percentage of peptides, and when they were used up by P. gingivalis as nutrients after growth, the repression effect was abolished. Taken together, these studies suggested peptides, rather than free amino acids repressed tpr expression. Since BHI broth was the most effective tpr repressor, we fractionated BHI by molecular size on a Sephadex G-10 column and analyzed individual fractions for their ability to suppress tpr expression. BHI fractions of low molecular weight all suppressed tpr expression to various extents at aconcentration of 20 mg/ml. Analysis of the amino acid composition of some of the fractions indicated that the fraction rich in phenylalanine, alanine and proline had the most repressive effect on tpr expression. The attempt to further fractionate this sample to identify the specific factor that repressed tpr expression was not successful probably due to the inability to obtain sufficient quantities of a particular peptide. Based on the BHI results, a number of commercially available peptides were examined and the di-peptide phenylalanyl- phenylalanine had the most repressive effect when supplemented to 5 mM. The supplementation of other peptides, nitrate, ammonium, succinate, sucrose, NaCI or heat shock, pH change and hemin limitation had small or no effect on tpr expression (Figure 29 and 30), suggesting this is not a response to nitrogen, heat shock or a stress response. The exact identity of the factor that represses tpr expression is not

88 certain, however, based on our studies, it suggests a short peptide with certain amino acid composition, especially phenylalanine, may be responsible. Marugg et al. (127) studied the transcriptional regulation of two proteinase genes in Lactococcus lactis.

They found the proteinase genes prtP and prtM were regulated at the transcriptional level by the peptide content of the medium and, to a less extent, by the growth rate.

The expression of prtP and prtM was high in whey permeate medium with relatively low concentrations of peptides, while in increased peptides media the expression was repressed. The effect of individual amino acids was not significant. They speculated that specific peptides in the growth medium played an important role in the regulation of the prt genes. They tested 11 dipeptides and 1 tripeptide and found the addition of the di-peptide leucylproline or prolylleucine to the growth medium specifically repressed prtP expression. There is a possibility that growth rate was responsible for tpr regulation. P. gingivalis grown in TYE had a generation time of 3.5 hours while cells grown in 0.5TYE had a generation time of 4.1 hours (166). If lower growth rate results in higher expression, then cells grown in BSA or gelatin supplemented 0.5TYE would have the highest level of tpr expression since with BSA or gelatin supplementation, the generation time decreased to 4.6 h and 4.2 h, respectively (166). Our Northern blot analysis showed that tpr expression was repressed when BSA or gelatin was added to growth media (Figure 14). The results indicate that growth rate is not the sole cause of tpr regulation. However, a role of growth rate in tpr expression could not be ruled out. A chemostat in which the growth rate can be controled would be a useful way to address this question.

Stationary phase cells had a higher level of tpr expression than cells of log phase

(Figure 17 and 27). This is similar to the E. coli rpoS regulon expression in which 30 or more genes are expressed when nutrients become limiting during the transition to stationary phase (115). Proteins in this regulon enhance long-term survival in nutrient-

89 deficient medium and have diverse functions including protection against DNA damage, the determination of morphological changes, the mediation of virulence, osmoprotection, and thermotolerance. Differential expression of families of genes within this regulon is affected by supplementary regulatory factors, working individually and in combination to modulate activity of different as dependent promoters. It has been difficult to assign a consensus sequence for as-dependent promoters because of the similarity to o7^ promoters and because sequence variations arising from the involvement of additional regulatory factors. Therefore, most as-controlled genes were identified by comparing their expression in wild type and as-deficient mutant strains.

Presently, we don't know if this regulon exists in P. gingivalis and no such mutants have been identified. The size of tpr mRNA (1.7 kb) is consistent with the result of the primer extension analysis showing that transcription started 215 bp upstream of the coding region and terminated at a potential rho-independent termination sequence approximately 1.7 kb downstream. Since the coding region of tpr is approximately 1.45 kb, it suggests tpr is transcribed monocistronically. The transcription start site 215 bp upstream of tpr coding region is not unusual since in the few studies of transcription in P. gingivalis genes, the detected transcription start sites were all further upstream of their coding regions. Karunakaran et al. (88) found the transcription started at a C residue 240 bp upstream of the coding region of hemR gene. Madden et al. (120) determined the prfT gene transcription start site to be of an A residue 510 bp upstream of coding region.

More studies are needed to determine if this phenomenon is common in P. gingivalis or if these examples are exceptions.

This 1.7 kb transcript could be detected in P. gingivalis cells grown in nutrient limited medium (0.5TYE) but not in nutrient rich medium (BHI). This is probably due to the low level of tpr expression in BHI. Although tpr was expressed in P. gingivalis

W83, no transcript could be detected in P. gingivalis ATCC33277, a type strain for this

90 species. This result is consistent with the finding by Park and McBride (169) that the tpr gene appeared to be different in strain ATCC33277. The different organization of the tpr gene among P. gingivalis strains and the regulated expression of it may explain why it was not detected in some studies (56, 177). We observed different P. gingivalis strains, different growth medium, and different stages of growth could all affect the expression of the tpr gene. To facilitate our study of the transcriptional regulation of tpr expression, we used a lacZ reporter gene to analyze the expression of tpr. lacZ is a well-studied reporter gene and has been used on numerous occasions in the isolation and analysis of promoters for various species of bacteria and eukaryotic cells. P. gingivalis is asaccharolytic and does not produce p-galactosidase. The use of a tprr.lacZ reporter gene construct made the analysis of tpr regulation much easier.

The lacZ gene lacks a promoter and ribosome binding site, therefore, its expression is under the control of tpr promoter and directly reflects the level of tpr expression. By analyzing p-galactosidase activity in P. gingivalis, one can follow the expression of the tpr gene. This is especially useful in a bacterial strain that possesses numerous proteases that make the analysis of proteolytic activity difficult.

The finding that lacZ expression was regulated in the same way in the single crossover and double crossover mutants indicated that the Tpr protein was not involved in its own regulation since in the allelic exchange mutant, no Tpr protease could be produced. The results also showed that a plasmid borne tprr.lacZ and a chromosome integrated tprr.lacZ exhibited the same kind of regulation pattern, suggesting that multiple copy shuttle vectors can be used to analyze the effects of 5' deletion mutations on tpr expression.

tprr.lacZ reporter analysis indicated that tpr promoter lies within the 618 bp 5' non- coding region. Further deletion analysis indicated that tpr promoter was located between nucleotides 315 to 400 (Figure 23). Computer analysis of this region did not

91 find significant homologous sequences to E. co//o70 promoter consensus sequences

(73). A putative set of -10 and -35 promoter regions was identified based on the relative position to the transcription start site (+1) and the optimal distance between them. The putative -35 region (5'-J_TCAGG) has 3 nucleotides identical to E. coli promoter -35 box (5'-TTGACA). The putative -10 region (5'-GCTCTT) has no homology to E. coli promoter Pribnow box (5'-TATAAT). The position of tpr promoter region was confirmed by deletion analysis of TX-30, in which the deletion to the -30 region abolished tpr expression (Figure 25).

In recent years, there have been a number of papers reporting the cloning and expression of P. gingivalis genes in E. coli. Some of these genes appeared to be transcribed from their own promoters and putative promoter regions (-10 and -35 regions) homologous to E. coli promoters were identified (16, 27, 32, 84, 86, 148, 179).

On the other hand, the DNA sequence of some other P. gingivalis genes, prflH (44), prtR (93), prpRI (3) and hemR (88) did not appear to have -10 and -35 promoter consensus sequences. Comparison of these putative promoter regions does not reveal a consensus sequence for a P. gingivalis promoter (Table 3). Furthermore, the majority of these promoter regions were identified based on similarity to E. coli promoters.

Klimpel and Clark (94) used antisera to the E. coli core enzyme and o70 to examine the RNA polymerase of P. gingivalis and found no cross-reacting proteins in P. gingivalis extracts with either antisera. This suggests that E. coli RNA polymerase may not be sufficiently similar to P. gingivalis RNA polymerase and the transcription apparatus in E. coli and P. gingivalis may be different. The recombinant Tpr was expressed regardless of the orientation of the insert on the vector (16, 169), suggesting that fprgene was transcribed from its own promoter in E. coli. (Figure 2).

However, our primer extension analysis indicated the transcription of tpr started further upstream. The finding that E. coli recognized a different segment of DNA for tpr

92 transcription supported the hypothesis that E. coli and P. gingivalis have different promoter and/or promoter recognition factors. It suggests that DNA regions homologous to E. coli promoters do not necessarily represent the P. gingivalis promoters. This finding is supported by this study, even though some cloned P. gingivalis genes appeared to be under the control of their own promoters, they may actually be transcribed from a DNA sequence that was recognized by E. coli as promoter but is not the true promoter in P. gingivalis. More studies on the transcription of P. gingivalis genes are needed to develop a better understanding of the P. gingivalis promoter structure. We identified 3 identical direct repeats between the transcription start site and the coding region of tpr. Deletion of these repeats abolished much of gene regulation even though the potential promoter region remained intact. This suggests that they are involved in tpr regulation. The involvement of direct repeats in gene regulation has been reported. Simon et al. (204) found the regulatory region of the E. coli torCAD operon, which encodes the anaerobically expressed trimethylamine N-oxide (TAMO) reductase respiratory system, contained four direct repeats of the decameric sequence 5'-CTGTTCATAT. They designated these repeats as tor boxes, and found they were the specific targets of a regulatory protein TorR involved in the torC expression. Progulske-Fox et al. (180) identified 4 direct repeats of 41 nucleotides in the promoter region of hagB gene in P. gingivalis. The function of these direct repeats is not known.

Since the hagB gene expression is regulated by growth conditions, they may play a role in gene regulation. Madden et al. (120) analyzed the prtT protease gene of P. gingivalis ATCC 53977, they found six regions of overlapping dyad symmetry down stream from the stop codon of the prfl gene, these may represent a rho-dependent transcription terminator (111, 176). Interestingly, within the putative transcription termination region of prfT, there are three 12 bp direct repeats, 5'-CTATATAGCTTT. This region is also flanked by one pair of long direct repeats of approximately 110

93 nucleotides with 92% identity. The function of these repeat are unknown at this time, they may represent a binding site for a regulatory protein. Most regulatory proteins bind to the 5' region of the promoter to exert effects on RNA transcription. The location of the 3 direct repeats indicated that they are not involved in the initiation of transcription. However, if the 3 direct repeats represent a regulatory protein binding site, it could regulate tpr mRNA synthesis at the transcript elongation and termination stage. The genes in the TyrR regulon (175) are regulated by the TyrR protein in conjunction with any one of the three aromatic amino acids tyrosine, phenylalanine, and tryptophan. In the aroP gene regulation of E. coli, the binding site for the TyrR repressor is located down stream and outside of the putative RNA polymerase binding site. There is some uncertainty about the mechanism of TyrR-mediated repression of aroP. One hypothesis is that in this case, the binding of the TyrR repressor to the TyrR boxes interferes with RNA chain elongation rather than initiation. He et al. (74) found the operator (PurR binding site) of the purB gene of E. coli was 242 bp downstream of the transcriptional start site and overlapped codons 62 to 67 in the structural gene. Further studies indicated that the PurR-mediated repression of purB occurred by a transcriptional roadlock mechanism. They identified a truncated purB mRNA species in a Northern assay. It was hypothesized that this truncated RNA was formed when RNA polymerase met the PurR repressor bound to the purB operator. In our Northern blot, only a 1.7 kb tpr mRNA was detected in all growth conditions. Due to the detection limit and small size of truncated RNA, we did not detect small size tpr messenger. Furthermore, our Northern blot analysis of the insertional mutant of tpr, W83/PM, suggested that truncated tpr mRNA was not stable (Figure 15). So we can not rule out the possibility that there was aborted tpr mRNA species.

Another explanation is that the identified promoter is not the sole promoter for the expression of tpr and an additional promoter is located downstream of the 3 direct

94 repeats. Although our primer extension experiment resulted in a cluster of primer extension products which vary in size by increments of a single nucleotide near the expected position, these incomplete extension products were probably caused by difficulties encountered by the reverse transcriptase to transcribe near the end of messenger RNA. Our primer extension analysis indicates there is only one primer extension product, suggesting there is only one tpr promoter. The 3 direct repeats are also short open reading frames, the first repeat is part of a short open reading frame of 19 codons, N-Met-Arg-Asp-Leu-Ser-Lys-His-Leu-Asn- Ser-Leu-Val-Phe-Leu-Tyr-Thr-Asn-Ala. The other two repeats are in one short open reading frame of 16 codons, N-Met-Arg-Asp-Leu-Ser-Lys-Lys-Met-Arg-Asp-Leu-Ser- Lys-Ser-Phe-Pro. There is a phenylalanine residue in each of the short open reading frames. Since we found that a phenylalanine dipeptide modified tpr expression, these open reading frames may act as transcriptional attenuating sequences similar to E. coli trp and S. typhimurium his operon expression. In these operons, transcriptional attenuation is achieved by coupling transcription and translation within a regulatory leader region in which the position of a translating ribosome dictates the formation of either an RNA secondary structure that causes transcriptional termination or an alternative secondary structure that precludes termination. However, our computer analysis of tpr 5' region did not reveal sequences that could form secondary structures that were required for the formation of alternative hair-pins and no p-independent transcriptional terminator sequences were identified.

It would be interesting to knock out each of the direct repeats one at a time to define the region that is involved in tpr regulation.

The mechanism of nutrient regulation of tpr expression in P. gingivalis W83 remains to be determined. Our current state of knowledge of tpr expression suggests the following possibilities: transcription of the tpr gene begins 215 bp upstream of the coding region. Initiation of transcription appeared not to be regulated. Regulation

95 appears to be at the transcript elongation and termination stages. The 3 direct repeats are involved in tpr regulation. There may be a regulatory trans-acting factor that directly or indirectly senses the amount of short peptides of certain amino acid composition in the cell and acts on the 3 direct repeats. Another possibility is that the 3 short open reading frames are attenuator sequences and the availability of certain amino acids in the cell will affect the translation rate of these leader sequences and result in the regulation of tpr. Clearly, more research is needed to have a better understanding of gene regulation in P. gingivalis.

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