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A study of a membrane-associated protease of porphyromonas gigngivalis W83 Park, Yoonsuk 1995

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A STUDY OF A MEMBRANE-ASSOCIATED PROTEASE OF PORPHYROMONAS GINGIVALIS W83 by YOONSUKPARK B.Sc, Yonsei University, Seoul, Korea, 1986 M.Sc, Yonsei University, Seoul, Korea, 1988 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE STUDIES (Department of Microbiology and Immunology) We accept this thesis as conforming to the required standard. THE UNIVERSITY OF BRITISH COLUMBIA February 1995 © Yoonsuk Park, 1995 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. (Signature) Department of M"^»fh~oU^ij > ^ Xm^M4ifCo( The University of British Columbia Vancouver, Canada Date 4^ ^ • ^^ DE-6 (2/88) ABSTRACT A protease gene (tpr) of Porphyromonas gingivalis W83 was cloned and expressed in Escherichia coli. The recombinant protease with an apparent molecular mass of 90 kDa was detected as a proteolytic band on bovine serum albumin (BSA)-substrate zymogram and characterized as a thiol-dependent protease. Western immunoblot analysis and in-vitro translation showed that the tpr gene encodes a 50-kDa polypeptide, suggesting that the 90-kDa active protease may be composed of a subunit having a molecular mass 50 kDa. The recombinant protein was expressed at high levels using a T7 RNA polymerase/promoter system. Antiserum raised to the recombinant protein was reactive to a native P. gingivalis protein with a molecular mass of 80 kDa. A specific protease (Tpr)-deficient isogenic mutant, P. gingivalis W83/PM, was generated by homologous recombination between P. gingivalis W83 chromosomal DNA and a suicide plasmid carrying the inactivated tpr gene in which a portion of the gene was replaced with an erythromycin resistance gene. Gelatin-substrate zymography and Western immunoblot analysis showed that W83/PM did not express the 80-kDa protein seen in the parent strain. Compared with W83, the mutant W83/PM showed a greatly reduced ability to hydrolyze the bacterial collagenase substrate, p-phenylazobenzyloxycarbonyl-L-prolyl-leucyl-glycyl-L-prolyl-D-arginine (Pz-peptide). The enzyme is associated with the cell membrane of W83. The Pz-peptidase activity of P. gingivalis W83 was characterized. No significant differences were found in the ability of W83 or W83/PM to hydrolyze collagen, or in their pathogenicity in a mouse model. To determine the role of Tpr in nutrient uptake by P. gingivalis, W83 and W83/PM were cultivated in media containing limiting amounts of amino acid/peptide nutrients and in the growth limiting media supplemented with BSA or gelatin. The growth of both strains was enhanced by BSA or gelatin. Cells of W83 grown in the nutrient limited media expressed higher levels of Tpr. ii A complementation study was performed by introducing the recombinant shuttle plasmid pBY3 containing the tpr gene into W83/PM. Expression of Tpr by the complemented strain was observed by gelatin-substrate zymography, Western immunoblot analysis and Pz-peptidase assay. i i i TABLE OF CONTENTS ABSTRACT ii TABLE OF CONTENTS iv LISTOFTABLES ix LIST OF FIGURES x ABBREVIATIONS, NOMENCLATURE AND SYMBOLS xii ACKNOWLEDGMENTS xv I. INTRODUCTION 1 1. Periodontal Disease and Porphyromonas gingivalis 1 1.1. Periodontal disease and periodontal microflora 1 1.2. Porphyromonas gingivalis 4 1) Classification of P. gingivalis 4 2) Nutrition of P. gingivalis 7 3) P. gingivalis in the etiology of periodontal disease 9 2. Porphyromonas gingivalis Virulence Factors 11 2.1. Adherence 11 1) Fimbriae 11 2) Hemagglutinating activities 12 3) Other adhesins 14 2.2. Capsules 15 2.3. Lipopolysaccharide (LPS) 16 2.4. Outer membrane vesicles 17 2.5. Superoxide dismutase and metabolic end-products 17 3. Proteases of Porphyromonas gingivalis 18 3.1. General properties of P. gingivalis proteases 19 1) Types of P. gingivalis proteases 19 iv (a) Trypsin-like protease 19 (b) Collagenase 22 (c) Glycylprolyl peptidase 23 2) Location of P. gingivalis proteases 23 3) Cloning of P. gingivalis protease genes 24 3.2. Potential roles of P. gingivalis proteases 26 1) Activities associated with colonization 26 2) Destruction of host tissue 28 3) Disruption of host defense mechanisms 29 4) Activities associated with growth 31 4. Purposes and Objectives of the Research 32 H. MATERIALS AND METHODS 33 1. Bacteria and culture conditions 33 2. DNA isolation and general recombinant DNA methods 33 3. Generation of a P. gingivalis genomic library and screening of clones 34 4. Southern hybridization 34 5. Electrophoretic techniques and zymography 35 6. Antiserum 36 7. Affinity purification of monospecific antibodies 37 8. Western blot immunoassay 37 9. In vitro translation 38 10. Synthetic oligonucleotides and PCR 38 11. Construction of pT7-YS and high-level expression of tpr 40 12. NH2-terminal amino acid sequence 40 13. Construction of pBY2-IN 41 14. Construction of pBY3 41 15. Conjugation 41 v 16. Cellular fractionation 44 17. Pz-peptidase assay and activity against other protease substrates 45 18. Effects of protease inhibitors and metal ions on Pz-peptidase activity 46 19. Effects of pH on Pz-peptidase activity 46 20. Collagenase assay with [14C]-labeled soluble collagen 46 21. Hemagglutinating activity 47 22. Virulence test 47 HI. RESULTS 49 1. Cloning of the tpr Gene 49 1.1. Isolation of a protease positive clone 49 1.2. The recombinant plasmid 51 1.3. S outhern hybridization 51 2. Identification of the Cloned Gene Product 54 2.1. BSA-substrate zymography 54 2.2. Western immunoblot analysis 54 2.3. In vitro translation 57 3. Characterization of the Cloned Gene and its Gene Product 57 3.1. Subcloning of the insert 57 1) Localization of the cloned gene 57 2) Zymography and Western immunoblot 59 3.2. Effects of protease inhibitors and substrate specificity 59 3.3. The tpr gene 63 3.4. Sequence homology with calpains 65 4. High-Level Expression of the Recombinant Tpr 65 4.1. Construction of pT7-YS 65 4.2. High-level expression of tpr 69 4.3. NH2-terminal amino acid sequence of Tpr 69 vi 4.4. Western immunoblot analysis 71 4.5. Antiserum raised against the inclusion bodies 71 4.6. Tpr in other P. gingivalis strains 74 5. Generation of a Tpr-Negative Mutant 74 5.1. Conjugation 74 5.2. Isolation of a Tpr-deficient mutant 76 5.3. Southern hybridization 76 5.4. Western immunoblot and zymography 76 5.5. P. gingivalisW83/PM 80 5.6. Pz-peptidase activity of P. gingivalis W83 and W83/PM 80 5.7. Identification of Tpr in P. gingivalis 83 6. Complementation 83 6.1. Isolation of the complemented strain 83 6.2. Stability of the plasmids in P. gingivalis WS3/PM 88 6.3. Pz-peptidase activity of the complemented strain 88 7. Characterization of Tpr 90 7.1 Effects of reducing agents 90 7.2 Effect of protease inhibitors and metal ions 90 7.3 Optimum pH 94 7.4 Effect of temperature 94 8. Determination of the Role of Tpr 99 8.1. Role of Tpr in collagen hydrolysis 99 1) Collagen hydrolysis 99 2) Effect of temperature and protease inhibitors 99 8.2. Role of Tpr in growth or nutrient uptake of P. gingivalis 102 1) Growth of P. gingivalis W%3 and W83/PM 102 2) Induction of Pz-peptidase activities under growth-limiting conditions 102 vii 8.3. Tpr and pathogenicity of P. gingivalis 106 IV. DISCUSSION 107 1. Molecular Genetics of P. gingivalis 107 2. Characterization of the tpr Gene and its Gene Product I l l 3. Characterization of Tpr and Determination of its Role 116 V. REFERENCES 122 viii LIST OF TABLES TABLE PAGE 1.1 P. gingivalis proteases 20 III. 1 Pz-peptidase activities and trypsin-like protease activities of P. gingivalis W83 and W83/PM 82 III.2 Pz-peptidase activity of the complemented strain, P. gingivalis W83/PM (pBY3). 91 III. 3 Effect of protease inhibitors on the Pz-peptidase activity of the crude membrane fraction of P. gingivalis W&3 93 III. 4 Effect of metal ions on the Pz-peptidase activity of the crude membrane fraction of P. gingivalis W83 95 III. 5 Generation time and final optical density of P. gingivalis W83 and W83/PM 104 ix LIST OF FIGURES FIGURE PAGE 1.1 Tissue comprising the periodontium in the healthy and disease state 2 1.2 Taxonomic history of the gram-negative, anaerobic black-pigmented rods 5 1.3 Nucleotide sequence of the cloned tpr gene from P. gingivalis W83 25 11.1 Construction of pT7-YS 39 11.2 Construction of pBY2-IN 42 11.3 Construction of pBY3 43 III. 1 Detection of proteolytic activity of the clone 50 111.2 Restriction endonuclease map of the recombinant plasmid, pYS307 52 111.3 Southern hybridization 53 111.4 Detection of proteolytic activity by BSA-substrate zymography 55 111.5 Identification of the cloned gene product by Western immunoblot analysis 56 111.6 Autoradiogram of the proteins expressed in the in vitro translation system 58 111.7 Localization of the cloned gene 60 111.8 Detection of proteolytic activities of the subclones by BSA-substrate zymography 61 111.9 Western immunoblot of the subclones 62 III. 10 Kyte-DoolMe hydropathy plot of the amino acid sequence deduced from the open reading frame of the P. gingivalis tpr gene 64 III. 11 Amino-acid sequence homologies between Tpr and calpains 66 III. 12 Comparison of the amino-acid sequences of Tpr and domain II of the large subunits of several calpains 67 III. 13 Polymerase chain reaction product 68 III. 14 Coomassie blue-stained SDS-PAGE and Western immunoblot of Tpr from a high-level expression system 70 III. 15 Western immunoblot of P. gingivalis W83 probed with the antibody purified against the native 90-kDa recombinant protein 72 III. 16 Western immunoblot of P. gingivalis W83 probed with anti-rTpr 73 III. 17 Immunological and enzymatic assays for Tpr in a number of strains of P. gingivalis 75 III. 18 Possible homologous recombinations between pBY2-IN and P. gingivalis chromosome 77 III. 19 Southern blot to identify the disrupted tpr gene 78 111.20 Western immunoblot and gelatin-substrate zymogram of P. gingivalis W83/PM. 79 111.21 Growth curves of P. gingivalis W83 and W83/PM 81 111.22 Two-dimensional SDS-PAGE analysis of the cellular fractions of P. gingivalis W83andW83/PM 84 III. 23 Western immunoblot of the cellular fractions of P. gingivalis W83 and W83/PM 87 III. 24 Western immunoblot and gelatin-substrate zymogram of the complemented strain, P. gingivalis W83/PM (pBY2) 89 111.25 Effect of reducing agents on P. gingivalis W83 Pz-peptidase activity 92 111.26 Effect of pH on the Pz-peptidase activity of P. gingivalis W83 96 111.27 Effect of temperature on the Pz-peptidase activity of P. gingivalis W83 97 111.28 Effect of temperature on the stability of the Pz-peptidase activity of P. gingivalis W83 98 111.29 [14C]-labeled collagen-hydrolyzing activities of the crude membrane fractions of P. gingivalis W83 and W83/PM 100 III. 30 Effect of temperature and inhibitors on the collagenolytic activity of P. gingivalis W83andW83/PM 101 III. 31 Growth of P. gingivalis W83 and W8 3/PM in TYE limited medium and protein supplemented medium 103 III. 32 Pz-peptidase and trypsin-like activities of P. gingivalis W83 and W83/PM 105 xi ABBREVIATIONS, NOMENCLATURE AND SYMBOLS 2D-PAGE anti-rTpr Ap APMA ATCC BApNA BCIP BHI pME BPB BSA DTT EDTA Em G+C GApNA GCF Gn h IL IS KDO Km LB two-dimensional polyacrylamide gel electrophoresis antiserum raised to the inclusion bodies containing the recombinant protein ampicillin 4-aminophenylmercuric acetate American Type Culture Collection a-2V-benzoyl-L-arginine p-nitroanilide 5-bromo-4-chloro-3-indolyl phosphate brain heart infusion P-mercaptoethanol black-pigmented Bacteroides species bovine serum albumin dithiothreitol ethylenediamine tetraacetate erythromycin guanine+cytosine glycyl-L-arginine p-nitroanilide gingival crevicular fluid gentamycin hours interleukine insertional sequence 2-keto-3-deoxyoctonate kanamycin Luria-Bertani medium LPS lipopolysacchari.de min minutes NBT nitroblue tetrazolium NEM N-ethylmaleimide NR nonreduced-nondenatured protein samples for SDS-PAGE analysis CD optical density PAGE polyacrylamide gel electrophoresis PBS phosphate-buffered saline pCMB p-chloromercuribenzoate PCR polymerase chain reaction PGE2 protaglandin E PMNL polymorphonuclear leukocyte PMSF phenylmethylsulfonyl fluoride Pz-peptide p-phenylazobenzyloxycarbonyl-L-prolyl-L-leucyl-glycyl-L-prolyl-D-arginine R reduced-denatured protein samples for SDS-PAGE analysis r resistant rTpr the recombinant gene product of tpr SAAPpNA iV-succinyl-L-alanyl-L-alanyl-L-prolyl-L-phenylalaninep-nitroanilide SDS sodium dodecyl sulfate SOD superoxide dismutase SSC NaCl-Sodium citrate buffer (3 M NaCl, 0.3 M Sodium citrate, pH 7.0) TBS Tris-buffered saline Tc tetracycline TLCK iV-a-p-tosyl-L-lysine chloromethyl ketone Tp trimethoprim TPCK iV-tosyl-L-phenylalanine chloromethyl ketone Tris Tris[hydroxymethyl]aminomethane xiii TYE Trypticase peptone-Yeast Extract medium X-gal 5-bromo-4-chloro-3-indolyl pVD-galactopyranoside ACKNOWLEDGMENTS I would like to express thanks to my supervisor Dr. Barry C. McBride for his supports and guidance during the whole project. It has been an honor for me to work in the best working environment. I also appreciate the support and advice of my thesis committee members, Drs. R.A.J. Warren, G. Weeks and V.-J. Uitto. My thanks to all the members of the laboratory. Especially, Angela Joe and Pauline Hannam for their friendship, encouragement and endless correction of my language skills, and Carol Mazur for her assistance. Finally, I would like to express heartfelt thanks to my parents and family for their endless love, and to my lovely wife Chanyoung for her love, understanding, encouragement and great support. xv I. INTRODUCTION 1. Periodontal Disease and Porphyromonas gingivalis. 1.1. Periodontal disease and periodontal microflora. The periodontium, surrounding and supporting the teeth, consists of gingiva, periodontal ligament, root cementum and alveolar bone (Fig. 1.1, A). Periodontal diseases refer to pathological conditions of the periodontium, and can be grouped broadly into gingivitis and periodontitis. Gingivitis is inflammation of the gingiva without an affect on the attachment apparatus of teeth. Periodontitis denotes destruction of connective tissue attaching the tooth and adjacent alveolar bone. Almost all adults in all parts of the world experience gingivitis and some degree of periodontitis. The advanced forms of periodontitis with extensive loss of tooth-supporting connective tissue and bone occur in approximately 7% to 15% of dentate adults. A healthy periodontium is associated with a 1 to 3 mm gingival sulcus adjacent to the tooth (Fig. 1.1, A). In this state, the gingiva contains numerous collagen fibers that ensure close proximity of gingiva to tooth. There is little infiltration of inflammatory cells. Gingivitis is characterized by large numbers of polymorphonuclear leukocytes (PMNLs) migrating through the gingival sulcus and junctional epithelium in response to challenge by plaque microorganisms. The inflammatory response of the gingiva results in tissue redness, edema, bleeding and tenderness. Gingival tissue swelling leads to an increase of gingival sulcus depth (4 to 5 mm) and the formation of periodontal pockets. Gingivitis can persist for prolonged periods without significant progression, or can lead to periodontitis. Periodontitis lesions exhibit gingival inflammation as well as deepening periodontal pockets, and apical migration of the junctional epithelium from crown to root surfaces, as a result of destruction of periodontal connective tissue (Fig. 1.1, B). Ulceration of the epithelium lining periodontal pockets is characteristic of active periodontitis. 1 A. Healthy B. Diseased gingival sulcus periodontal ligament alveolar bone cementum Periodontal pocket area of bone resorption Junctional epithelium FIGURE 1.1. Tissues comprising the periodontium in the healthy (A) and diseased (B) state. 2 Virtually all forms of periodontal disease are inflammatory infections that appear to be initiated by bacterial plaque (160). The first step in determining the etiologic agents of different forms of periodontal disease is to compare the microbiota present in sulci of healthy individuals, and in periodontal pockets of individuals with clinical symptoms. The study of the microflora associated with periodontal diseases is quite difficult; complications include defining the clinical status of the diseases, obtaining samples of the microflora representative of the disease status without contamination, and properly identifying and quantitating the large number of different species within the sulcus. However, a number of studies have revealed some important characteristics of subgingiva microflora (for a review, see reference 280). Human periodontal disease is associated with complex microflora in which more than 350 microbial species can be found (270). Many bacterial species can initiate gingival inflammation if bacterial cells are present in high numbers, associated with poor oral hygiene. The production of sufficient noxious microbial compounds induces an inflammatory response in the host. The development of gingivitis, thus, probably constitutes a nonspecific infection. In contrast, a more limited number of species appears to be responsible for the conversion of a gingivitis lesion to a progressive periodontitis lesion with periodontal tissue destruction and loss of alveolar bone. The development of destructive periodontitis often seems to be the result of a specific infection (161). The healthy gingival sulcus harbors a microflora dominated by gram-positive organisms and facultative anaerobic species (270), in which Actinomyces and Streptococcus species each account for about 40% of total isolates. With increasing severity of disease, the proportions of anaerobic, gram-negative, and motile organisms increase significantly (55, 270-272, 315). While some of these microorganisms may be present in low numbers as members of the indigenous microflora, it appears that an increase in their numbers and activity is correlated with degradation of periodontal connective tissue (316). It has been also shown that many of the anaerobic gram-negative isolates from periodontitis lesions belong to the group of microorganisms formerly known as black-pigmented Bacteroides (BPB) (181,274, 338, 360). 3 Among a limited number of microorganisms present in elevated proportions in many cases of adult periodontitis, Porphyromonas gingivalis is believed to be closely associated with advanced adult periodontitis and seems to be one of the numerically most important pathogens in the disease (for a review, see references 181,279). This species is generally not detected in the healthy gingival sulcus. The following sections are devoted to a description of P. gingivalis and studies which provide evidence of its role in adult periodontitis. 1.2. Porphyromonas gingivalis. 1) Classification of P. eineivalis. Many of the isolates from periodontitis lesions have been shown to be gram-negative, nonmotile, nonsporeforming anaerobic rods that produce black pigment when grown on blood agar plates. Originally, most of these microorganisms were thought to be a single species, Bacteroides melaninogenicus. It is now clear that the black-pigmented strains represent 3 different genera and 10 different species, 8 of which can be isolated from the human oral cavity. Collectively they are known as black-pigmented Bacteroides (BPB). There has been confusion in the interpretation of the literature prior to 1980, because of numerous changes in taxonomy of BPB species (Fig. 1.2). The changes in the taxonomy of BPB are reviewed in this section. In 1921, Oliver and Wherry (215) isolated small anaerobic gram-negative rods, which produced black-pigmented colonies when grown on blood agar plates. These microorganisms can be isolated from a variety of sites, including the oral cavity, urine, human feces, and respiratory tract, as well as from postsurgical infections. Because the pigment was considered to be melanin, the bacterium was named Bacterium melaninogenicum. This bacterium was classified in the third edition of "Bergey's Manual of Determinative Bacteriology" (19) as Haemophilus melaninogenicus because it grew better on solid medium containing the X (protoporphyrin IX or protoheme) and V (nicotinamide adenine dinucleotide [NAD] or NAD phosphate [NADP]) growth factors characteristic of the genus Haemophilus. Subsequently, 4 ly> f B. melaninogenicus subsp. melaninogenicus Bacterium melaninogenicum \ Haemophilus melaninogenicus \ Bacteroides melaninogenicus \ B. melaninogenicus subsp. intermedius J * } J } B. melaninogenicus B. loescheii B. denticola B. intermedius B. corporis , , , , ; P. melaninogenica P. loescheii P. denticola P. intermedia P. corporis N S 1 B. melaninogenicus subsp. asaccharolyticus \ B. asaccharolyticus J * ) *harolyticus B. gingivalis B. endodc I * I Genus Prevotella P. asaccharolytica P. gingivalis P. endodontalis \ / Genus Porphyromonas FIGURE 1.2. Taxonomic history of the gram-negative, anaerobic black-pigmented rods. Roy and Kelly (234) reclassified this organism to the genus Bacteroides, and named it Bacteroides melaninogenicus. Although biochemical and immunological heterogeneity was shown among strains of B. melaninogenicus (42, 245), only one species of BPB was recognized until 1970. Increased understanding of their physiology, particularly their fermentative capability of glucose, led to division of B. melaninogenicus into three subspecies (103); B. melaninogenicus subspecies melaninogenicus (strongly fermentative), B. melaninogenicus subspecies intermedius (weakly fermentative), and B. melaninogenicus subspecies asaccharolyticus (nonfermentative). Finegold and Barnes (62) showed that the biochemical and genetic characteristics of the saccharolytic and asaccharolytic strains were sufficiently different to justify the elevation of the asaccharolytic subspecies to the species level, B. asaccharolyticus. Further, the studies (43, 261, 332) showing the genetic heterogeneity between oral and nonoral asaccharolytic B. melaninogenicus species led Coykendall et al. (43) to propose the new species B. gingivalis for asaccharolytic BPB species isolated from oral sites. B. asaccharolyticus was retained for asaccharolytic strains isolated from nonoral sites. The DNA base content of B. gingivalis varied from 46.5 to 48.4 mol% G+C, while that of B. asaccharolyticus varied between 49.2 and 53.6 mol% G+C (43). In 1984, a third asaccharolytic BPB species, B. endodontalis, was described (337). Although originally classified as B. asaccharolyticus, B. endodontalis had little or no DNA homology with either B. asaccharolyticus or B. gingivalis. Shah and Collins (252) found that, based on the biochemical and chemical evidence, the asaccharolytic BPB species, B. asaccharolyticus, B. gingivalis and B. endodontalis differ markedly from the type species of the genus Bacteroides, B.fragilis, and proposed that these species should be reclassified into a new genus Porphyromonas as shown in Fig. 1.2. The weakly saccharolytic and strongly saccharolytic species are now classified into a new genus Prevotella, with Prevotella melaninogenica (B. melaninogenicus) as the type species (Fig. 1.2) (251). 6 In addition to P. gingivalis, several other members originated from human oral cavity in subgingival sites. These include Prevotella melaninogenica, Prevotella denticola, Prevotella loescheii, and Prevotella intermedia. In addition to 8 species shown in Fig. 1.2, Bacteroides macace (43) and B. levii (122) were isolated from a periodontal site in the macaque monkey Macaca aractoides and from bovine rumen and cattle horn abscess, respectively, and were included in the group formerly as BPB. In this dissertation, BPB will occasionally be used to refer to the collection of organisms which were previously grouped under this name. 2) Nutrition of P. gingivalis. P. gingivalis is an obligate anaerobe, and requires hernin and vitamin K (menadione) for growth (78). For specific isolation of P. gingivalis, a medium containing bacitracin, colistin and nalidixic acid as selective agents has been developed (109). The roles of hemin and vitamin K in the growth of P. gingivalis are not completely understood, although there has been a proposal that these function as electron carriers in electron transport (78). Vitamin K has been found to stimulate synthesis of phospho-sphingolipids in the cell envelope, suggesting a possible role in membrane structure (158). Gibbons and MacDonald (78) suggested that the dark pigment produced by P. gingivalis is a mechanism for storage of hemin. This idea was supported by the observation of Rizza et al. (227) who showed that, when P. gingivalis was grown in a medium containing a high concentration of hemin, the cells were able to divide 8 to 10 times after transfer to a medium without hemin, supporting the previous suggestion. Recently, the dark pigment was characterized as protohemin with traces of protoporphyrin (250), which is possibly a storage form of hemin. Succinate can replace hemin as a growth factor for P. gingivalis (82, 85, 182). It was shown that the growth of P. gingivalis can be supported by succinate supplied by Treponema denticola, another suspected periodontopathogen (82). Mayrand and McBride (182) also demonstrated that succinate produced by other bacteria affect the infectivity of P. gingivalis, 1 as well as its growth. The P. gingivalis strain studied was not infectious when inoculated subcutaneously as a pure culture into guinea pigs. However, coinoculation with succinate-producing microorganisms resulted in a rapid progression of disease. Further, incorporation of hemin or succinate into the inoculum was sufficient to replace the need for a second microorganism. P. gingivalis is unable to ferment sugars (261). In addition, P. gingivalis showed a very poor growth response to increased concentration of glucose, and [14C]-glucose was not converted to metabolic end products (253, 260). The low levels of [14C]-glucose taken up were incorporated into cellular components (260). P. gingivalis can obtain energy by fermenting amino acids or peptides (75,249,260,340). The addition of various protein hydrolysates, such as trypticase and proteose peptone, to a basal medium or a defined medium enhances the growth of P. gingivalis (249,260). Gharbia and Shah (75) recently studied the utilization of peptides by P. gingivalis using of fluorescamine labeling techniques. The studies showed that, compared to high-molecular-weight oligopeptides, low-molecular-weight peptides (10 to 14 residues) are preferentially taken up by P. gingivalis. P. gingivalis also appeared to prefer peptides as energy source rather than amino acids. Although P. gingivalis has limited ability to uptake free amino acids, a few amino acids appear to be taken up as readily as peptides (75, 259, 260). The dicarboxylic amino acids aspartate and glutamate appeared to be significant catabolic amino acids. The periodontal pocket of the human oral cavity, the major site of colonization of P. gingivalis, is bathed in protein-rich gingival crevicular fluid (GCF). P. gingivalis is known to produce proteases capable of hydrolyzing these components as well as collagen (discussed in section 3.1). The proteases make P. gingivalis ideally suited for colonization in the periodontal pocket (site of tissue degradation), since the activity of these enzymes must release peptides for the growth of the microorganism. In addition, GCF contains other factors 8 that are necessary for the growth of some subgingival microorganisms, including hemin which is an essential growth factor for P. gingivalis. 3) P. gingivalis in the etiology of periodontal disease. Unlike gingivitis, the development of destructive periodontitis appears to be the result of a specific infection. The following is devoted to a discussion of the evidence for the involvement of P. gingivalis in the pathogenesis of periodontitis. Several studies have shown a direct relationship between the appearance of P. gingivalis as a dominant cultivable species in subgingiva microflora and the occurrence of periodontitis (37, 39, 52, 164, 275, 315, 316, 343, 359). Tanner et al. (315) first demonstrated a positive correlation between clinical inflammation and P. gingivalis. P. gingivalis appeared to be only detectable in periodontal pockets of patients with severe gingival inflammation, but not in healthy gingival sulci (343). It was also shown that sites associated with recent bone loss contained a higher proportion of P. gingivalis in the subgingival microbiota than inactive sites (316). Christersson et al. (39) examined the prevalence of P. gingivalis in a large patient population. 94% of the 53 patients with severe adult periodontitis harbored P. gingivalis. However, P. gingivalis was detected in only 9.9% of the reference group comprising a large number of general dental patients (1559 subjects). Case reports have provided additional evidence, implying that the elimination of P. gingivalis can arrest the disease. Loesche et al. (163) reported that P. gingivalis constituted 30 to 40% of the subgingival microflora of severe periodontitis patients, and that treatment with metronidazole eliminated the organism and resolved the disease. A case of severe, recurrent periodontitis was reported by Van Dyke et al. (330) in which P. gingivalis was the major microorganism (approximately 70%) in cultivable microflora. Success in controlling the disease was correlated with the elimination of cultivable P. gingivalis from the sites after intensive therapy. Immunological studies provide additional support for the role of P. gingivalis in periodontal disease. Serum antibody levels to P. gingivalis were higher in patients diagnosed 9 with periodontal diseases (48,56, 58,192,339). Several studies reported that antibody levels to P. gingivalis were higher in GCF than in serum, suggesting a local immune response to this organism (57,59,124,318). Direct evidence was obtained in the early studies which showed that asaccharolytic BPBs including P. gingivalis play an important role in the pathogenesis of transmissible mixed anaerobic infections in animal models (85,167,182, 303). Combinations of bacteria isolated from various sources (plaque, necrotic dental pulp and periodontal pockets) were tested for their capacity to induce abscess formation and transmissible infections when injected subcutaneously in guinea pigs. A synergistic effect was commonly observed. Individual bacterial species did not induce an infection, whereas mixtures of two or more bacteria produced an infection. The inclusion of P. gingivalis in several different mixed-culture inocula resulted in the development of necrotic lesions, and produced a transmissible infection, suggesting that P. gingivalis plays a central role in the pathogenesis of periodontal disease. Experiments with both mice and guinea pigs showed that P. gingivalis can infect by itself (73, 88, 128, 198, 335). Kastelein et al. (128) demonstrated that F. gingivalis can produce lesions when injected intradermally or subcutaneously in the animals. Histological examination of the tissue showed evidence of invasion of the connective tissue. This animal model has been used as a simple virulence test for P. gingivalis. Genco et al. (73) developed a mouse subcutaneous chamber model that allows for the examination of host-parasite interactions as well as the determination of gross pathology with P. gingivalis challenge. When inoculated into stainless-steel chambers implanted subcutaneously in mice, P. gingivalis induced a systemic IgG response, and caused phlegmonous, ulcerated, necrotic lesions, and death. Several research groups have examined experimental infections in periodontal tissues of animals other than mice or guinea pigs to observe the function of P. gingivalis in in-vivo systems which mimic the human periodontium (20, 104, 115, 133, 211). Holt et al. (104) 10 reported that the introduction of P. gingivalis into the periodontal microbiota of monkeys resulted in an increase in systemic levels of antibody to the microorganism, and rapid and significant bone loss. This indicated that P. gingivalis is capable of functioning as a primary pathogen in periodontal disease. 2. Porphyromonas gingivalis Virulence Factors. For an organism to cause periodontal disease, it must be able to (i) colonize the gingival sulcus, (ii) resist host defenses, and (iii) produce factors which result in tissue damage. While the involvement of P. gingivalis in the pathogenesis of periodontal disease has been clearly demonstrated, little is known about specific virulence factors. However, a number of putative virulence factors of P. gingivalis have been identified. 2.1. Adherence. The ability of periodontopathic bacteria to adhere to surfaces in the oral cavity is a prerequisite for both colonization and disease initiation. Colonization of bacteria in the oral cavity is facilitated by binding to host tissue or other bacteria. Bacterial cell surfaces have specific adhesins which are responsible for attachment to specific host receptors. These adhesins include type-specific pili or fimbriae and surface-binding proteins. P. gingivalis possesses a variety of adhesion factors which may enable colonization of the gingival sulcus. 1) Fimbriae. The fimbriae of P. gingivalis have been isolated and purified, and their morphological, immunological and chemical properties characterized (156, 212, 289, 356, 357). They are heat stable, thin, curly filaments, approximately 5 nm in width. Fimbrillin subunits produced by different strains of P. gingivalis are heterogeneous (156). The fimbriae of P. gingivalis have been thought to confer hemagglutinating activity (213). However, pure fimbrial preparations did not exhibit hemagglutinating activity, or hemagglutination inhibitory activity (357). Boyd and McBride (25) also demonstrated that removing fimbriae from P. gingivalis 381 had no effect on the hemagglutinating activity of whole cells, suggesting no correlation 11 between the presence of fimbriae on the cell and hemagglutinating activity. In contrast to these studies, a recent report by Ogawa and Hamada (208) demonstrated that P. gingivalis fimbriae caused the agglutination of erythrocytes from different sources. Fimbriae extracted from P. gingivalis were found to attach to hydroxy apatite coated with collagen, a major component of the gingival connective tissue, suggesting that P. gingivalis fimbriae play an important role in colonization of this organism (195). Lee et al. (155) have also shown that P. gingivalis, as well as the purified fimbriae, binds to saliva-coated hydroxyapatite in a concentration-dependent manner. This binding was inhibited by the purified fimbriae and by several synthetic peptides having sequences corresponding to the carboxy-terminal 1/3 of the predicted sequence of the fimbrillin subunit. Recently, two separate research groups (93, 169) reported the generation of a nonfimbriated P. gingivalis mutant by inactivating the gene (fimA) encoding the fimbrial subunit protein of P. gingivalis 381, fimbrillin, and demonstrated that the FimA protein is essential for the interaction of the organism with human gingival tissue cells, as well as with saliva-coated hydroxyapatite. The nonfimbriated mutant was significantly less able to cause periodontal bone loss in gnotobiotic rats (169). P. gingivalis fimbriae stimulated bone resorption in vitro (130). P. gingivalis fimbriae induced gene expression and production of some inflammatory cytokines in human gingival fibroblasts and mouse peritoneal macrophages (94). In addition, P. gingivalis fimbriae appeared to induce migration of monocytes, which in turn may contribute to either host defense or tissue damage during the pathogenesis of periodontal disease (208). These results suggested that P. gingivalis fimbriae may be involved in the alveolar bone loss observed in adult periodontitis, as well as in colonization of the organism. 2) Hemagglutinating activities. Hemagglutinating activity may be a marker of bacterial cell attachment to mammalian cells. P. gingivalis agglutinates erythrocytes isolated form various animals (25, 183, 213, 214, 277, 278, 310). In fact, this property is an important taxonomic character which can distinguish P. gingivalis from P. endodontalis and P. asaccharolytica (179, 183, 277, 337). 12 Although hemagglutinating activity has been observed for years, attempts to purify and characterize hemagglutinating factors are rather recent P. gingivalis capsular polysaccharide or lipopolysaccharide (LPS) did not exhibit hemagglutinating activity, and the extracts did not inhibit hemagglutination (212). Okuda and Kato (210) showed that, although LPSs from various species in subgingival microflora possess the ability to adhere to erythrocytes, P. gingivalis is one of the bacterial species whose LPS does not cause hemagglutination. In contrast, Ogawa and Hamada (208) recently demonstrated that P. gingivalis LPS could agglutinate erythrocytes from many animal species. Boyd and McBride (25) have also shown that the hemagglutinating activity was associated with low-molecular weight fraction containing LPS, protein, and loosely bound lipid. Mouton et al. (190) identified a cell-bound hemagglutinin, and designated it HA-Ag2. HA-Ag2 did not react with antiserum raised to P. gingivalis fimbrillin (191). In addition, immuno-gold labeling revealed HA-Ag2 to be a surface-associated amorphous material distinct from the fimbriae (51). These findings indicated that the hemagglutinating activity of P. gingivalis is not related to fimbrillar structure. It differs from the observation of Ogawa and Hamada (208). Several groups have isolated hemagglutinin(s) of P. gingivalis (113, 214) from extracellular material. The purified hemagglutinin(s) consisted of at least two protein components which might contain loosely bound lipid components. Interestingly, the inhibition of the hemagglutinating activity by a low concentration of L-arginine was observed. It is possible that arginine functions as a contact residue between the bacterial cell receptor and its counterpart on the erythrocyte during agglutination. A cell surface-associated hemagglutinin of P. gingivalis was isolated by Singh (268). The native molecule had an apparent molecular mass in excess of 103 kDa, and was composed of subunits with a molecular mass of 43 kDa. Since purified P. gingivalis hemagglutinins exhibit different characteristics, it is possible that more than one component may function as a hemagglutinin. 13 It was shown that the hemagglutinating and trypsin-like enzyme activity of P. gingivalis were inhibited by the same chemical reagents, suggesting that both biological activities may exist on the same molecule (202, 203). However, a report by Shah et al. (258) attributes hemagglutinating activity and proteolytic activity to separate molecules. They proposed that these are probably closely associated on the outer membrane of P. gingivalis and may be functionally related. Pike et al. (221) isolated separate proteases with arginine and lysine specificity from a high molecular mass fraction of the P. gingivalis culture supernatant. These proteases were found to be noncovalently complexed with hemagglutinins. A hemagglutinin gene of P. gingivalis 381 has been cloned and expressed in Escherichia coli (224). The gene product was found to be located on the P. gingivalis cell surface. 3) Other adhesins. The initial stages of colonization by P. gingivalis may involve attachment to other bacterial species in dental plaque within the gingival sulcus, particularly as P. gingivalis colonizes preexisting plaque. P. gingivalis coaggregates with several bacteria commonly found in the indigenous microflora of the oral cavity, including various species of Streptococcus and Actinomyces (278). Boyd and McBride (25) indicated that a bacterial aggregating component isolated from the outer membrane of P. gingivalis was composed of protein, carbohydrate, and a high-molecular-weight LPS fraction. A number of research groups have studied coaggregation of P. gingivalis with other oral microorganisms, including Treponema denticola (79) and Fusobacterium nucleatum (136,138). Close contact between these species strongly associated with active disease would favor the establishment of pathogenic microbial population in the periodontal pocket. Several studies have shown that P. gingivalis binds to human epithelial cells (36, 116, 117, 212, 278) and to hydroxyapatite coated with host substances such as collagen, serum, saliva, or crevicular fluid (41, 194). The adhesin molecules that mediate binding have not been identified. The involvement of P. gingivalis fimbriae in attachment to epithelial cells and saliva-coated hydroxyapatite has been suggested (117,155). P. gingivalis has also been 14 known to bind human fibrinogen (144,147,148) and fibronectin (116,146). It was suggested that adherence to fibrin and fibronectin may facilitate colonization of host tissue, and lead to degradation of gingival tissue or interference with tissue repair (147,150). 2.2. Capsules. Bacterial capsules have various functions: they can serve as physiochemical barriers for the cell, they provide protection against desiccation by binding water molecules, can promote attachment, and they can be antiphagocytic in that they act to avoid engulfment by PMNLs and to prevent hydrolytic degradation of microorganisms engulfed by PMNLs (for a review, see reference 30). The first electron microscopic observation of BPBs (311) showed the presence of capsular structures. Mansheim and Kasper (172) observed an electron-dense layer external to and associated with the outer membrane of P. gingivalis strains. van Steenbergen et al. (336) showed that the virulent strains of P. gingivalis, W83 and W50, were more resistant to killing by human serum and PMNLs than those strains which were less virulent. The virulent strains had a thicker capsule and were more hydrophilic than the less virulent strains. However, a clear correlation between the presence of capsules on P. gingivalis and virulence of the bacteria in the mouse model was not established. It was shown that, while encapsulation and resistance to phagocytosis in vitro were features of the invasive strains in experimental infections, these factors alone were not determinants for bacterial virulence (301). The exact role of capsules in the pathogenicity off. gingivalis is, therefore, still not clear. Grenier and McBride (92) recently reported that the cells of a P. gingivalis recovered from an experimental infection had a thicker and much denser extracellular layer than cells grown under laboratory conditions. Neiders et al. (198) have found that freshly isolated P. gingivalis strains were invasive in the mouse pathogenicity model while laboratory strains were not. It has been suggested that the observed difference in virulence is due in part to a difference in capsular structures. 15 2.3. Lipopolysaccharide (LPS). Bacterial LPS or endotoxin exhibits a wide variety of biological activities in vitro and effects the physiology of living organisms in a number of ways (345). P. gingivalis LPS has an unusual chemical composition (101, 102, 143, 172, 173, 193) with low levels of both heptose and 2-keto-3-deoxyoctonate (KDO) (26,121,140). Kiley and Holt (134) have suggested that this may contribute to weak linkages between the lipid A and core units, and could result in the release of free lipid A and polysaccharide moieties into the environment. They also showed that free LPS is capable of penetrating the crevicular epithelial layer to basal and parabasal cells near the basement membrane. Classical endotoxin assays show that P. gingivalis LPS has little endotoxic activity (101, 172, 304, 305). Apart from its low endotoxin activity, P. gingivalis LPS appears to be capable of stimulating a wide range of in vitro responses in almost every type of cell found in the periodontium (for a review, see reference 345). It stimulates bone resorption (110,186, 193, 306), and inhibits bone collagen formation in a rat calvarial system (186). The ability of P. gingivalis LPS to inhibit bone formation as well as to stimulate bone resorption suggests that it could be an important mediator of the bone loss accompanying periodontal disease. Recently, Loomer et al. (165) suggested that P. gingivalis might contribute to the bone loss in periodontal diseases not only by stimulating resorption, but also by inhibiting bone formation directly. P. gingivalis LPS stimulates both gingival fibroblasts (308, 313, 351) and macrophages (26, 95) to produce several inflammatory cytokines, including interleukines (IL-1, IL-6 and IL-8), prostaglandin E (PGE2), and tumor necrosis factor. Hopps and Sismey-Durrant (107) reported that rat periosteal fibroblasts stimulated with P. gingivalis LPS induced bone resorption in the mouse calvarial bone resorption assay. The extent of the resorption was much greater than that stimulated by P. gingivalis LPS alone, implying that the LPS was inducing fibroblasts to secrete a bone resorption factor(s), such as PGE2 and IL-1. 16 Larjava et al. (149) reported that P. gingivalis LPS inhibited growth of human gingival fibroblasts, the cells primarily responsible for maintaining the structural integrity and functions of the periodontium. 2.4. Outer membrane vesicles. Gram-negative bacteria, including P. gingivalis (178, 286), have been shown to form membranous extensions or outgrowths of the outer membrane during in vitro growth, and to release the fragments as vesicles (blebs) into the growth environment. A number of biological activities have been reported for the extracellular vesicles of P. gingivalis. The protein profiles of outer membrane and vesicle fractions are very similar (87, 344), suggesting that vesicles contain at least some of the binding receptors present on whole cells. P. gingivalis vesicles attach to hydroxyapatite surfaces coated with saliva, serum or crevicular fluid (41, 269). Vesicles increase the attachment capability of Streptococcus sanguis to serum-coated hydroxyapatite by about 10-fold (269). P. gingivalis vesicles stimulate coaggregation of some oral bacteria, including Eubacterium saburreum and Capnocytophaga (87), and Actinomyces strains (24,60). Hydrolytic enzymes including trypsin-like protease, collagenase, glycylprolyl dipeptidase, alkaline phosphatase, Af-acetyl-P-glucosaminidase, are associated with cell-free vesicles (87, 287), suggesting that vesicle formation might be an important mechanism whereby cells can export hydrolytic enzymes into the extracellular environment (187). While the role of the vesicles in pathogenesis is unclear, characterization of their biological activities could serve to amplify and extend the pathogenic potential of P. gingivalis. Their small size (50 to 150 nm diameter) may allow them to cross epithelial barriers that are impermeable to whole cells; indeed, they have been found in the interstitial space between epithelial cells (220). 2.5. Superoxide dismutase and metabolic end-products. 17 Phagocytosis of bacteria by PMNLs is accompanied by enhancement of PMNL oxidative metabolism. Oxygen metabolites such as superoxide anion (02~), hydrogen peroxide (H202), and hydroxy radical (OH") are major components of the bactericidal activity of PMNLs. Many anaerobic pathogenic microorganisms produce enzymes, such as superoxide dismutase (SOD), catalase, and peroxidase, which protect the organism from these toxic reactive products (16). Among BPBs, P. gingivalis has the highest SOD activity, and is the most tolerant to the presence of oxygen (7). Amano et al. (4) also demonstrated that exogenous SOD purified from P. gingivalis (5,6) inhibited bacterial killing of PMNLs. P. gingivalis produces a thermostable protein which interferes with the bactericidal activity of PMNLs by modulating the generation of reactive oxygen species (352). Rotstein et al. (232, 233) suggested that succinic acid, a metabolic end-product of P. gingivalis, is responsible for inhibiting 02~ production by PMNLs. Several of the metabolic end-products of P. gingivalis may contribute to the virulence of this organism. Butyrate and propionate, the characteristic end-products of asaccharolytic metabolism, are potent inhibitors of various cultured human or animal cell lines (86,267, 323, 334). Volatile sulfur compounds produced by P. gingivalis are also believed to be important cytotoxins because they influence the permeability of oral mucosa (200), and reduce collagen synthesis (322). Other potentially toxic end-products of P. gingivalis include indole and ammonia (166). 3. Proteases of Porphyromonas gingivalis. Proteolytic enzymes are important pathogenic determinants in a number of diseases (18, 27, 63, 108, 126, 139, 174, 204, 238, 239, 314). Slots and Dahlen (276) suggested that virulence factors of P. gingivalis mainly involve proteases with potential to interfere with host defenses and to disintegrate periodontal tissue. P. gingivalis has been distinguished from other periodontopathic microorganisms by its thiol-dependent collagenolytic and proteolytic enzymes (151, 152, 273, 319, 321). This section will discuss proteases produced by P. gingivalis, and their possible role in the pathogenicity. 18 3.1. General properties of P. gingivalis proteases. 1) Types of P. eineivalis proteases. For a considerable length of time, the proteolytic activity of P. gingivalis was classified into two groups: (i) enzymes which specifically degraded collagen, and (ii) "trypsin-like" proteases hydrolyzing the synthetic peptide, a-N-benzoyl-L-argininep-nitroanilide (BApNA). P. gingivalis proteases purified and characterized were summarized in Table 1.1. Analysis of those proteases suggested that they could be grouped into three major types, trypsin-like protease, collagenase, and glycylprolyl protease. (a) Trypsin-like protease. Since Yoshimura et al. (355) first reported trypsin-like extracellular protease in 1984, most studies to date have supported the idea that the general proteases of P. gingivalis are trypsin-like in nature. A number of studies have reported on the purification and characterization of the enzymes (17, 34, 68-70, 255, 256, 281, 283, 285, 294, 325). The activity of all the purified enzymes was enhanced by reducing agents such as cysteine, pV mercaptoethanol (fJME), and dithiothreitol (DTT), indicating that these enzymes belong to the thiol protease group. Shah et al. (255,256) provided more definitive evidence for the cysteine protease nature of the P. gingivalis proteolytic enzymes by using a method of isolation specific for thiol-containing proteins and a type of time-dependent inhibition that is highly selective for thiol groups. The reactivity of the P. gingivalis enzymes was characteristic of those reported for well-characterized cysteine proteases. BApNA hydrolysis by P. gingivalis vesicles/supernatant was abolished by reaction with 2.2'-dipyridyl disulfide (a two-hydronic state thiol inhibitor and reactivity probe) (28, 29), and reactivated by (3ME, indicating that the enzymes belonged to the cysteine protease group. Purified trypsin-like proteases of P. gingivalis were active against a variety of native proteins including bovine serum albumin, gelatin, and casein. Some were capable of hydrolyzing host defense proteins and structural proteins (17, 61, 217, 244, 285, 294). The molecular mass of the enzymes ranged from 35 to 300 kDa. Some of them may represent the 19 TABLE 1.1. P. gingivalis proteases. Type or Name (Strain) Trypsin-like protease (ATCC 33277) Trypsin-like protease (ATCC 33277) Trypsin-like protease (381) Trypsin-like protease (W50) Gingivain (W83) Gingipain-1 (H66) Trypsin-like protease (381) Collagenase (381) Collagenase(llOl) Collagenase (ATCC 33277) Source Culture supernantant Cell extract Cell extract Vesicles Supernatant/ Vesicles Culture supernantant Culture supernatant Cell envelope extract Cell extract Cell surface M.W. (kDa) 300 (gel filtration) 53 (gel filtration) 45 (gel filtration) 50 (SDS-PAGE/100°C)b 58 (SDS-PAGE/100°C) 95 (SDS-PAGE/25°C) 50(SDS-PAGE/100°C) 44 (gel filtration) 55 (SDS-PAGE/100°C) 90 (zymography) 94 (SDS-PAGE/37°C) 120 (SDS-PAGE/25°C) 50,60 (SDS-PAGE/100°C) Thiol-dependency ND + + + + + + + + -Substrate BApNA, azoalbumin, azocasein, asocoll BApNA, gelatin, type IV collagen BApNA, BSA, casein, gelatin BApNA, type I collagen, fibronectin BApNA BApNA BApNA, collgen (type I, III, IV and V), C3, fibrinogen, fibronectin gelatin, type I, n and III collagen type IV collagen, gelatin type I collagen, Pz-peptide, kininogen, transferrin PMSF ND SI(l) ND ND MI NI (10) NI (10) NI(1) WI (10) ND Effect of inhibitors3 (mM) TLCK SI (0.01) ND SI (0.5) ND ND SI(1) SI (5) MI(1) M(l) ND TPCK ND ND NI(5) ND ND SKD NI (10) M(l) ND ND EDTA SI (1.0) SI (10) MI (10) ND WI(5) SI(1) ND SI (5) WI (10) ND pCMB SI (0.01) WI (1.5) NI (0.5) ND ND ND ND ND ND ND Ref. (68) (294) (325) (285) (254) (34) (17) (21) (154) (290) Glycyprolyl protease Outermembrane 29 (SDS-PAGE/37°C) - Gly-pro pNA, gelatin, MI (4) SI (2) NI(2) NI MI (4) (90) (ATCC 33277) 20 (SDS-PAGE/100°C) azocoll, azocasein (20) TABLE 1.1. (continued) Type or Name (Strain) Glycylprolyl aminopeptidase (381) Glycylprolyl peptidase (A7A1-28) Lysine-specific protease (ATCC 33277) Lys-gingivain (ATCC 33277) Lys-gingipain (H66) Argingipain (381) Source Cell extract Cell extract Culture supernatant Cell envelope extract Culture supernatant Culture supenatant M.W. (kDa) 75 (SDS-PAGE/100°C) 80 (SDS-PAGE/100°C) 75 (gel filtration) 48 (SDS-PAGE/100°C) 68 (gel filtration) 70 (SDS-PAGE/100°C) 60 (SDS-PAGE/100°C) 44 (SDS-PAGE/100°C) Thiol-dependency - ND ND + + + Substrate Gly-pro MCA Gly-pro pNA Tosyl-gly-pro-lys pNA, BSA,IgG,IgA Pi-lys containing synthetic peptide, fibrinogen, kininogen Z-lys-pNA Z-phe-arg-MCA, IgG, Collagen (type I & IV) PMSF MI (0.5) SI (5) NI(1) NI NI (10) WI(1) Effect of inhibitors8 (mM) TLCK ND NI(1) SI(D ND NI (10) SI(1) TPCK ND ND ND ND ND SI(1) EDTA NI(1) NI(5) Act0 NI NI (10) SKD pCMB WI(1) ND ND ND ND ND Ref. (188) (15) (71) (248) (221) (123) NI, no inhibition (80-100% of initial activity); WI, weak inhibition (50-80% of initial activity); MI, moderate inhibition (25-50% of initial activity; SI, strong inhibition (0-25% of initial activity); ND, not determined. Protein samples for SDS-PAGE were prepared at temperature indicated on the parenthesis. c  Enzyme activity was induced. same enzyme in an aggregated form or in complexes with other cellular components. Fishburn et al. (64) showed that the extent of serum protein proteolysis by culture supernatants of P. gingivalis correlated with the thiol-dependent, trypsin-like enzyme activity. It was also shown that trypsin-like activity of P. gingivalis was stimulated by glycine-containing peptides (33). This unusual property may have evolved in response to the glycine-rich environment which occurs when gingival collagen is degraded, (b) Collagenase. The ability of what is now known as P. gingivalis to degrade collagen was first demonstrated by Gibbons and MacDonald (77). To date, this is the only BPB organism with this capability (180,299, 331). Type I, II, HI, and IV collagens are degraded by P. gingivalis. Grenier and Mayrand (88) found that, with one exception, virulent strains of P. gingivalis have higher-levels of collagenolytic activity that non-virulent strains. Toda et al. (321) have demonstrated that P. gingivalis collagenases are active in a reduced environment, and suggested that this would be in keeping with a role in the degradation of collagen fibrils in the anaerobic environment of a periodontal pocket. Birkedal-Hansen et al. (21) classified a collagenolytic enzyme of P. gingivalis as a true collagenase, if it could dissolve reconstituted collagen fibrils at 35°C and attack the helical domain of interstitial collagens at 22°C. The P. gingivalis enzyme meeting these requirements had a molecular mass of 90 kDa, was dependent on free thiol groups and on metal ions, and was associated with the bacterial cells. Several investigators have reported purification and characterization of the collagenolytic enzymes (17, 154, 290, 294). Lawson et al. (154) purified a P. gingivalis collagenase which exists as an active precursor protein of molecular mass 94 kDa. Biotin-labeled collagen bound specifically to the 94 kDa form of the protein and to its cleavage products in ligand blots, suggesting a role of this enzyme not only in collagen degradation but also in adhesion to collagenous substrata. A collagenase purified by Sojar et al. (290) was capable of hydrolyzing rat plasma low-molecular-weight kininogen and transferrin, as well as salt-22 solubilized type I collagen from rat skin, suggesting that the enzyme has specificity for the Pro-X-Gly sequence found in those proteins, (c) Glycylprolyl peptidase. A statistically significant correlation has been found between the level of glycylprolyl (gly-pro) peptidase activity in crevicular fluid and the clinical severity of periodontal disease (8). Furthermore, Suido et al. (297) found a significant correlation between gly-pro peptidase activity in crevicular fluid and the number of P. gingivalis. They also demonstrated the usefulness of measurement of this enzyme activity in the diagnosis of P. gingivalis -associated periodontal disease. Gly-pro peptidase activity of P. gingivalis has been reported by several research groups (3, 15, 90, 91, 131, 188, 296). Four enzymes which are capable of hydrolyzing gly-pro dipeptide have been purified and characterized (3, 15, 90, 188). All the enzymes have been shown to be serine-group proteases, based on their sensitivity to specific inhibitors. In contrast to the trypsin-like and coUagenolytic enzymes, the gly-pro peptidase is not activated by reducing agents. Given the high levels of glycine and proline in collagen, such gly-pro peptidases may act as helper enzymes following an initial cleavage of the collagen by specific collagenases. A gly-pro peptidase purified by Abiko et al. (3) cleaves peptides of partially digested type I collagen, suggesting that the enzyme may be involved in degradation of native collagen with the cooperation of true collagenase. 2) Location of P. gingivalis proteases. Many investigators have successfully isolated and characterized P. gingivalis proteases from culture supernatant (3,17, 34, 69,71,99,217, 254, 285) and different cellular fractions, including cell extracts (15, 70, 188, 248, 294, 296, 325), cell envelope (68, 69, 90, 91, 145, 290, 355) and vesicles (45, 69, 256, 281, 283). By using polyacrylamide gels containing covalently bound bovine serum albumin (BSA), Grenier et al. (84) determined the proteolytic profiles of various cellular fractions of P. gingivalis. Eight distinct bands of proteolytic activity were detected with apparent molecular weights varying from 29 to 100 kDa. Seven 23 proteolytic bands were detected in cell extracts, outer membranes and vesicles, whereas the culture supernatant contained four. No band was detected in membrane-free extracts, suggesting that the proteases were associated with the cell envelope. Studies by Fujimura and Nakamura (69), and Smalley and his coworkers (282,284) have also shown that the vesicles are responsible for most of the protease (trypsin-like) activity in the culture supernatants. Therefore, it is believed that the role of P. gingivalis proteases is likely to be related to their cellular location (cell envelope and vesicles). Smalley et al. (284) examined the distribution of trypsin-like activity in batch cultures of P. gingivalis W50 and an avirulent variant (W50/BE1). The results suggested that the difference in the distribution of the trypsin-like activity between the particulate (cells and vesicles) and the soluble enzyme fractions may be of greater significance than the absolute levels of the enzyme as pathogenic determinant. 3) Cloning of P. gingival™ protease genes. The use of molecular biological techniques will be useful in determining the number of proteases and in defining the role of each in the virulence of P. gingivalis. Recently, several protease genes of P. gingivalis have been cloned and characterized (10, 23, 65, 129,168,196, 216, 309). Each of the protease genes cloned is genetically distinct. Arnott et al. (10) showed that the recombinant clones producing proteolytic clear zones on skim-milk plates were seen only when the plates were incubated anaerobically in the presence of P-mercaptoethanol, indicating that the recombinant protein was a thiol-dependent protease. Bourgeau et al. (23) reported a similar result. In their study, the clone contained a 3.0-kb insert that encoded for a protease with an apparent molecular mass of 64 kDa in BSA-substrate zymography. Sequencing part of the 3.0-kb DNA fragment revealed an open reading frame encoding a protein of 482 amino acids with a molecular mass of 62.5 kDa (Fig. 1.3). A comparison of the amino acid sequence with sequences listed in the GenEMBL Sequence Data Library, the GenNBRF Protein Sequence Data Library, and the SwissProtProtein Sequence library revealed the presence of a 23-amino-acid consensus sequence (amino acids 406 to 430) near the C terminus that had a degree of homology of 60 24 90 GTTAACGGTTTTTCATGCGTGATTTGTCCAAATTGCACCTTAATTCGTT^^ - 3 5 - 1 0 1 8 0 CGTGATTTGTCCAAGTCTTTTCCTTAATATATCCCTTCATTTGTGT^^ SD 2 7 0 TAAMTTTTTCAATTATGGAAAAGAAATTAGTACCGCAATCCATTTC&AA^^ M E K K L V P Q S I S K E R L Q K L E A Q A T L T 3 6 0 CCTCAACAAGAGGAAGCGAAAGCCCGTAAAATCGAAAGAGAGAAAGCCAGACTAAAAGAACTGAACATTCCTACCGAATCTAAAGAATCC P Q Q E E A K A R K I E R E K A R L K E L N I P T E S K E S 4 5 0 AAAGATTGCAGCCCTGCAGGGATGATCAATCCATATGCACTTACTGAAGTCATTTTAGAAAGACCATTGGATTGGAGCAACCCACGGACT K D C S P A G M I N P Y A L T E V I L E R P L D W S N P R T 5 4 0 ACCGATATAGTAGAACGCGTCTrGGGTrCAAGCATGCAAGATCTATCCAAAGGCGACTCTCTATTAAGAGCCGGAAGAGACCAAAATGCT T D I V E R V L G S S M Q D L S K G D S V L R A G R D Q N A 6 3 0 GAGGTCAAAATCGTGGATTCAGTTITGACTAAGACCCAAAGAGGGCAGGACGGTCTGGAGAGAA E V K I V D S V L T K T Q R G Q D G L E R I L E S P N D T D 7 2 0 ATGCCTCCCGAAGAAAAAGAAGAAGCTGCTCCAAAGGCAAAAAAAGCAGCCCAAAAACTCGACATCGACGACCTCAGAGAGCAAGCACTG M P P E E K E E A A P K A K K A A Q K L D I D D L R E Q A L 8 1 0 TCATCCACTACTATCACAAAGGAGATCAGCAAGATCATCCTTCCGACAAAAAATTTAAGAGATGATAATAATACAGTACATCAGTACAGA S S T T I T K E I S K I I L P R K N L R D D N N T V H Q Y R 9 0 0 GAAGTCGGCTTCCAAAGCAATGGAGCACACAACTTGTGGGACACAGTAGTTCAAGGAATCGCTGGCGAT^ E V G F Q S N G A H N L W D T V V Q G I A G D C Y M L A A L 9 9 0 TCGGCTATAGCTTGGGTATGGCCCGCTCTATTGAATATGGATCTCGACATCATGTCTAATCAAGATGAATGGCGACTCTATCGCTACTTC S A I A W V W P A L L N M D V D I M S N Q D E W R L Y R Y F 1 0 8 0 ATAGGGCGCTCTAAACAGACATATGCCAACCGGCCGTCGGGGTCAGGTACCTCCACCAATGAAATTCTTCAGGAAGGATATTACAAAGTT I G R S K Q T Y A N R P S G S G T S T N E I L Q E G Y Y K V 1 1 7 0 CCGATCTTTGCAAGGAGTCGCTATTCGTTCAACGGAGAATACTGGCCGGCTCTTTTCGAACAAGCATATGCCAATTGGAAATTCCC^ P I F A R S R Y W F N G E Y W P A L F E Q A Y A N W K F P N 1 2 6 0 GATTCCAAATACAACGCAATCCTACAGATTGGAGGTGGCTGGCCTGAGGAAGCACTTTGCG^ D S K Y N A I L Q I G G G W P E E A L C E L S G D S W F T S 1 3 5 0 TCGGGAAAACTCATCCTTTCTTCTTTCACAGATCTGTCATTGCTGAATT^ S G K L M L S S F T D L S L L N F M K S M C Y S W K T I K P 1440 ATGGTGATTGTAACCCCATGCTGGGACCCTCTACCTCCTATGATGCCCGGAATTGCAGCATACCATGCCTATACAGTTTTGGGATATACA M V I V T P C W E P L P P M M P G I A A T H A Y T V L G Y T 1530 GTTTCCAATGGAGCCTATTACCTGATTATCCGCAATCCATGGGGAGTGACTGAGCCAACAGGAGATGGAGTGCTAAGCAAAAGAGATTGG V S N G A Y Y L I I R N P W G V T E P T G D G V L S K R D W 1 6 2 0 GTTATCCACTTCGATAATATCAAGTGGTTCAATCTATCCAAAGACGATGGCATTTTC^ V I H F D N M K W F N L S K D D G I F A L R L D K V R E N F 1 7 1 0 TGGTACATCGCATATATGTATTGATATCAGATTATATTATACGGTTTGAGAACGATGAGTCGGCCCGGTAGAGCGGGCAGCCTCTjpTCT W Y I A Y M Y * 1 8 0 0 TTTCTGTACACGTCAGGCAACAGGCTCJMCCATGGATCTATCTGCCCAATTCCGAATCGACAAAGCTCCATATCGGAGCATACACT FIGURE 1.3. Nucleotide sequence of the cloned tpr gene from P. gingivalis W83 (23). The sequence is derived from the 1.8-kb fragment between MHCII mdHindUl sites of the insert (see Fig. III.7). 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. 25 to 65% with thiol or cysteine proteases from a wide range of species (Fig. 1.3, underlined amino acid sequence). The recombinant enzyme was found to hydrolyze azocoll, casein and bovine serum albumin (BS A), and was inhibited by TLCK, TPCK, pCMB and EDTA Kuramitsu and his coworkers have cloned two protease genes, prtT (216) and prtC (129, 309) from P. gingivalis ATCC 53977, which encode a thiol-dependent, trypsin-like protease and a collagenase, respectively. The complete nucleotide sequence of the prtT gene was determined, and the deduced amino acid sequence of the enzyme corresponded to a 53.9-kDa protein. Gelatin-substrate zymography also indicated a similar molecular size for the protease. The prtC gene appeared to encode the 35-kDa collagenase, which degraded soluble and reconstituted fibrillar type I collagen, heat-denatured type I collagen, and azocoll but not gelatin or the synthetic collagenase substrate. The result of Northern blot analysis suggested that the gene may be transcribed as part of a polycistronic mRNA. Most recently, Fletcher et al. (65) reported the cloning of a protease gene (prtH) from P. gingivalis W83. Casein-substrate zymography revealed a protease with an apparent molecular mass of 97 kDa. This gene encodes an enzyme which hydrolyzes the human C3 complement protein. An open reading frame encoding a 110-kDa protein of 992 amino acids was identified in the nucleotide sequence. 3.2. Potential roles of P. gingivalis proteases. P. gingivalis proteases may play numerous roles in the pathogenesis of periodontal disease, for example, (i) in colonization, (ii) in destruction of the host tissue, (iii) in inactivation of host defense mechanisms, and (iv) in bacterial nutrition during infection. 1) Activities associated with colonization. Proteases may be involved in bacterial adherence via two mechanisms: (i) exposure of hidden binding sites, and (ii) direct binding involving the active sites of the enzymes. Gibbons (76) has postulated a mechanism by which the hidden segments of molecules, cryptitopes, become exposed and are available to participate in adhesion. Pretreatment of 26 epithelial cells and fibronectin-collagen complexes with trypsin significantly enhanced the adherence of P. gingivalis (31, 35, 36,194). Naito and Gibbons (194) suggested that elevated levels of proteases in the gingival sulcus, associated with poor oral hygiene and gingivitis, might remove fibronectin and expose collagen molecules in the basement membrane, thereby promoting the attachment of P. gingivalis cells and facilitating their invasion into gingival tissues. P. gingivalis collagenase appears to be located on the bacterial cell surface (296, 321), and could serve as the adhesin responsible for promoting binding to collagen. Naito et al. (194) provided the evidence supporting this possibility by showing that binding to the collagenous substrata was inhibited by the serum protease inhibitors, and by inhibitors of thiol proteases. Several research groups have presented evidence for the involvement of the cell-bound P. gingivalis trypsin-like activity in adherence to host cells and other oral bacteria (45, 80, 106, 159, 202, 203). On the basis of inhibition experiments, Nishikata et al. (203) proposed that the hemagglutinating and proteolytic activities of P. gingivalis are exhibited by the same molecule, and that the same site of the molecule participates in the binding to erythrocytes and to substrate. More recently, a hemagglutinin possessing proteolytic activity was purified from the membrane fraction of P. gingivalis by the same research group (202), confirming the previous proposal. Supporting evidence was recently provided by Curtis et al. (45) who showed that an active-site-directed inhibitor of trypsin-like enzyme, tyrosyl-alanyl-lysyl-arginine chloromethyl ketone, inhibited P. gingivalis whole cell hemagglutination. Li et al. (159) demonstrated that, compared with the parent strain, P. gingivalis mutants with low-level protease activity have a decreased ability to coaggregate with Actinomyces viscosus. Hoover et al. (106) found that a P. gingivalis mutant deficient in trypsin-like protease activity exhibited reduced hemagglutinating activity. Grenier (80) reported that P. gingivalis with high levels of cell-associated trypsin-like activity attach in higher numbers to human epithelial cell than strains with low-level trypsin-like activity. It was suggested that 27 adherence of young P. gingivalis cells involves trypsin-like protease, whereas old cells possess additional attachment mechanisms. 2) Destruction of host tissue. Proteolysis of host tissue is a characteristic of chronic inflammatory periodontal disease (218, 243). Collagen (mostly type I), an important and major constituent of the gingival connective tissue, is one of the constituents degraded in adult periodontitis (67). Although collagen, a triple-helix protein, is resistant to a wide variety of proteolytic enzymes, it is degraded by bacterial and tissue collagenases, both of which exist in the mouth (72, 162). In addition to the true collagenase(s), some of purified trypsin-like proteases of P. gingivalis showed collagenolytic activity (17, 285, 294). The collagenolytic activity of P. gingivalis may account for at least some of the destruction of gingival connective tissue observed in periodontitis. P. gingivalis may cause collagen hydrolysis by two mechanisms; directly through cleavage by P. gingivalis enzymes, and indirectly by proteolytic activation of a latent host collagenase (329) and induction of the secretion of collagenase activity by human gingival fibroblasts (22). The main role of the basement membrane in the periodontium is to maintain tissue integrity, to provide a support for normal epithelial cell functions, and to serve as a barrier regulating the passage of cells and particles between tissue compartments. Destruction of the basement membrane may enable pathogenic microorganisms to invade host tissues. Uitto et al. (326) have reported the degradation of basement membrane collagen (type IV) by proteases from human gingiva and dental plaque, and suggested that the collagenolytic activities in periodontal disease may originate from both plaque bacteria and human leukocytes. P. gingivalis has been shown to be able to attach to a basement membrane-like matrix and to degrade it (348). In addition, cellular extracts of P. gingivalis have been shown to degrade basement membrane components, including type IV collagen (154, 327) and fibronectin (150, 285). Thus, P. gingivalis proteases may enable the organism to damage the tissue barrier, and/or to invade the subepithelial basement membrane and gain access to the 28 connective tissues. Saglie and his co-workers (235-237) have identified P. gingivalis in connective tissue histological sections, indicating the bacterial invasion during periodontal disease. Uitto et al. (328) have shown that a trypsin-like protease from P. gingivalis degraded cell surface components of human gingival fibroblasts. Furthermore, protease activity of P. gingivalis was implicated in the detachment of squamous epithelial cells (257). These results suggested that the proteases can damage the epithelial cells and loosen the structure of the epithelial tissues. 3) Disruption of host defense mechanisms. P. gingivalis produces proteolytic enzymes that degrade most of the serum proteins involved in defense against bacterial infections, including immunoglobulin, complement factors, protease inhibitors and transferrin. Kilian (135) first demonstrated the ability of P. gingivalis to hydrolyze IgA and IgG to small peptides. Other research groups have further characterized the immunoglobulin hydrolyzing activity of the organism (66, 71, 89, 244, 300). Grenier et al. (89) showed that P. gingivalis produces several proteases which non-specifically cleave immunoglobulins, and suggested that the degradation occurs in two stages; the molecules are first degraded into large fragments which are subsequently degraded into small peptides. Degradation of immunoglobulins by P. gingivalis proteases may protect suspected periodontal pathogens by interfering with the immunoglobulin-mediated host defense system. P. gingivalis may protect itself from the antibacterial effects of complement by proteolytically inactivating the key proteins of the complement system. Several studies have shown the degradation of complement factors C3, C4, C5 and C5a (246, 247, 298, 300, 302, 347). Sundqvist et al. (302) showed in~vivo degradation of the C3 protein using teflon cages subcutaneously implanted in guinea pigs. Recently, Cutler et al. (46) reported that inhibition of C3 and IgG proteolysis by P. gingivalis enhanced phagocytosis of the organism. These findings support a role of degradation of immunoglobulins and complement factors by P. 29 gingivalis in its resistance to phagocytosis. Wingrove et al. (347) demonstrated that a protease isolated from P. gingivalis, gingipain-1 (34), is capable of generating biologically active chemotatic factor C5a from C5. C5a, if generated at the site of infection, could result in a significant leukocytic infiltration and help to explain the rapid onset of the inflammatory response associated with progressive periodontitis. A trypsin-like enzyme, isolated from P. gingivalis vesicles, inactivated the bactericidal activity of human serum (81), supporting the previous observation by Grenier and Belanger (83) that P. gingivalis vesicle-associated proteases protected Capnocytophaga ochracea and Prevotella loescheii against the bactericidal action of human serum. A study by Lala et al. (142) found that trypsin-like protease(s) secreted from P. gingivalis attenuated the bactericidal activity of neutrophils by cleaving the formyl-peptide receptor molecule involved in the stimulation of neutrophil activation. Cleavage of the receptor molecule may alter the ability of neutrophils to defend against pathogenic bacteria. Additionally, a trypsin-like protease of P. gingivalis inhibits PMNL phagocytosis (346). A trypsin-like enzyme isolated from P. gingivalis has been shown to be able to degrade lysozyme from PMNLs (61, 217). The level of lysozyme in saliva of patients with severe periodontitis was significantly reduced when compared with that of healthy controls (175). Inflammation plays an important role in the host defense against microorganisms invading the tissues. In human plasma, there are four highly integrated systems (kininogen, complement, clotting, and fibrinolytic) which produce inflammatory mediators. These systems are characterized by specific consecutive proteolytic reactions (proteinase amplification cascades). The roles of the plasma proteases of the inflammatory cells include digestion of bacteria, enhanced locomotion through connective tissue, demarcation of the site of infection, and tissue remodeling. The plasma proteinase cascades are modulated by plasma proteinase inhibitors such as CI-inhibitor, o^-antiplasmin, anti thrombin, and c^-macroglobulin (324). Uncontrolled release of proteases in inflammation by the inactivation of the inhibitors causes self-digestion, greater tissue destruction, and a rapid progression of 30 disease, due to enhancement of the inflammatory response, vascular permeability and fibrinolysis. P. gingivalis possesses specific protease(s) capable of degrading human plasma proteinase inhibitors (17, 31, 100, 201, 248). P. gingivalis proteases may rapidly inactivate the important protease inhibitors in human plasma, and thus paralyze the host defenses against invading microorganisms. In addition, P. gingivalis proteases, isolated from a culture supernatant, were able to enhance vascular permeability in a dose-dependent manner when injected into guinea pigs (112, 125). Imamura et al. (112) recently showed that a cysteine protease gingipain-1 (34) is the major vascular permeability enhancement factor of P. gingivalis. Scott et al. (248) purified a membrane-associated thiol lysyl-protease that preferentially cleaves the Pl-Lys position of the chromogenic tripeptide substrates. This enzyme, named as lys-gingivain, cleaves both kininogen and fibrinogen rapidly destroying the functional activity of both. Kininogen and fibrinogen are multifunctional plasma proteins participating in various phases of the inflammatory process and the major protein substrate of the coagulation cascade, respectively. Lys-gingivain also appeared to be the most potent kininogenase and fibrase (fibrinogen-digesting enzyme) to be described to date. 4) Activities associated with growth. P. gingivalis is an asaccharolytic organism whose metabolism is dependent on small peptides and amino acids (75, 249, 259, 340), and, therefore, degradation of host proteins is an essential requirement for survival. Grenier et al. (89) demonstrated that growth of P. gingivalis was stimulated by supplementing growth limiting media with IgG. Hemin is an essential growth factor for the growth of P. gingivalis. In the host, this compound is found in complexes with the plasma proteins, such as albumin, hemopexin, haptoglobulin and transferrin, which are involved in the transport of body iron and in preventing undue loss of iron through urinary excretion (225). P. gingivalis is capable of degrading these proteins (32), and, as reported by Inoshita et al. (114), the organism is able to 31 utilize transferrin as a source of iron. The degradation of the plasma proteins by P. gingivalis may fulfill the requirement of iron, as well as hemin, for bacterial growth. In addition, the hemolytic activity of P. gingivalis has been characterized by several research groups (40, 132, 258). It was suggested that P. gingivalis may meet its hemin requirement by using hemoglobin released from red blood cells. Shah and Gharbia (254) also demonstrated the direct involvement of a secreted cysteine protease of P. gingivalis in lysis of red blood cells. 4. Purposes and Objectives of the Research. Despite intensive studies for a decade, the role of P. gingivalis proteases in pathogenesis still remains to be determined. The purpose of this study was to characterize a P. gingivalis protease, and to determine the potential role of the enzyme. The general objectives of this study were as follows; i) Clone a gene encoding a P. gingivalis protease and characterize the gene product, ii) Generate an isogenic mutant of P. gingivalis lacking the protease, iii) Use the isogenic mutant to determine the role of the protease in growth, metabolism and virulence. 32 H. MATERIALS AND METHODS 1. Bacteria and culture conditions. Porphyromonas gingivalis W83, ATCC 33277, W50 (ATCC 53978), W12, P. asaccharolytica ATCC 25260, Prevotella corporis ATCC 33547, Prevotella denticola ATCC 33135, Prevotella intermedia ATCC 15032, Bacteroides levii ATCC 29147, Prevotella loescheii ATCC 15930, Prevotella melaninogenica ATCC 25845 were grown in brain heart infusion (BHI) broth (Difco Laboratories, Detroit, Mich.), supplemented with hemin (5 |ig/ml) and vitamin K (0.5 |j,g/ml), at 37°C in an anaerobic chamber (Coy Manufacturing, Ann Arbor, Mich.) containing an atmosphere of N2, H2 and C02 (85:10:5). Human blood and agar were added to BHI broth to final concentrations of 5% (vol/vol) and 1.5% (wt/vol), respectively, to prepare BHI-blood agar plates. Erythromycin (Em), gentamycin (Gn) and tetracycline (Tc) at final concentrations of 10 (Xg/ml, 200 |J.g/ml and 10 |a.g/ml, respectively, were added to the media when required. In some studies, P. gingivalis was grown in TYE broth (1.7% Trypticase peptone, 0.3% Yeast extract, 0.5% NaCl, 0.25% KH2P04) supplemented with hemin (5 |ig/ml) and vitamin K (1 (ig/ml). For a medium containing growth limiting amount of amino acid/peptides, the trypticase peptone was reduced to 0.5% (0.5TYE broth) (89). Escherichia coli was grown aerobically in Luria-Bertani (LB) broth (242) at 37 °C. LB agar plates were prepared by adding agar at final concentration of 1.5%. Ampicillin (Ap), kanamycin (Km) and trimethoprim (Tp) were added when required, at a final concentrations of 100 |J,g/ml, 50 |ig/ml and 200 |ig/ml, respectively. Growth of P. gingivalis strains was determined by measuring the increase in optical density at 660 nm (OD660) using an ICM colorimeter (ICM, Hillsboro, Oreg.). 2. DNA isolation and general recombinant DNA methods. 33 Plasmid DNA was isolated by the alkali-lysis method and further purified when necessary by equilibrium centrifugation in cesium chloride-ethidium bromide gradients (242). For general purposes, plasmid DNA was isolated by the miniprep method described by Rodriguez and Tait (230). The method described by Silhavy et al. (266) was used to prepare chromosomal DNA of P. gingivalis W83 for generation of a genomic library. Small scale chromosomal DNA preparation was done by the miniprep method (11). All recombinant DNA procedures were carried out as described by Sambrook et al. (242). 3. Generation of a P. gingivalis genomic library and screening of clones. Purified chromosomal DNA from P. gingivalis W83 was partially digested with endonuclease Sau3AI, and fractionated on a 10-40% (wt/vol) sucrose density gradient to yield size-selected DNA fragments from 2 to 10 kb. Size-selected fragments were ligated with dephosphorylated pTZ18R (185) linearized with BamHI using T4 DNA ligase. The ligation mixture was used to transform E. coli JM83 made competent by CaCl2 treatment (242). A representative P. gingivalis genomic library was obtained by selecting ampicillin-resistant white E. coli colonies from LB agar plates containing 5-bromo-4-chloro-3-indolyl |3-D-galactopyranoside (X-gal). Pro tease-positive clones were detected in LB agar plates supplemented with 1% (wt/vol) powdered skim milk (Carnation). Transformants were transferred onto LB agar plates containing skim-milk and incubated overnight at 37 °C. After exposure to chloroform vapor for 1 h, the plates were incubated at 37°C and then examined for areas of clearing around the colonies. 4. Southern hybridization. To prepare probe DNA, the plasmid was digested with the appropriate restriction enzyme(s), and the desired DNA fragment was extracted from an agarose gel after electrophoresis by the methods described by Silhavy et al. (266). The DNA fragment was 34 labeled with biotin-7-dATP using the BRL Nick Translation System (BRL, Gaithersburg, Md.). Southern hybridization was performed using the non-radioactive nucleic acid detection system (BluGENE™, BRL, Gaithersburg, Md.) with slight modifications of the recommended protocol. Briefly, test samples of plasmid and chromosomal DNA were digested to completion with the appropriate restriction enzyme(s), and electrophoresed in a 0.8% agarose gel. DNA in the gel was transferred to nitrocellulose by capillary transfer using 20XSSC (3 M NaCl, 0.3 M sodium citrate, pH 7.0) as described by Sambrook et al. (242). Hybridization was done overnight at 65°C with the probe in hybridization buffer comprising 6XSSC, 50 mM Tris-HCl (pH 7.5), heat-denatured salmon sperm DNA (100 ng/ml), 0.5% (wt/vol) SDS and 1% Denhardt's solution. The blot was washed two times in 2xSSC containing 0.1% SDS at room temperature, two times in 0.2XSSC containing 0.1% SDS at room temperature, and two times in 0.16XSSC containing 0.1% SDS at 55°C. After a brief rinse in 2XSSC at room temperature, hybridized bands were detected by incubation of the blot with streptavidin-alkaline phosphatase followed by a substrate solution containing 5-bromo-4-chloro-3-indolyl phosphate (BCIP) and nitroblue tetrazolium (NBT) (Sigma, St. Louis, Mo.). 5. Electrophoretic techniques and zymography. Sodium dodecyl sulfate (SDS)-10% polyacrylamide gel electrophoresis (PAGE) was carried out by the Laemmli method (141), using a Mini PROTEAN II cell (0.075 cm thick) (BioRad Laboratories, Richmond, Calif.). Protein samples were prepared in two different ways. For nonreduced-nondenatured (NR) samples, whole cells or cellular fractions were solubilized in sample solubilization buffer (4% [wt/vol] SDS, 20% [vol/vol] glycerol, 0.125 M Tris-HCl [pH 6.8]) for 30 min at 37°C. Reduced-denatured (R) samples were incubated in the same buffer containing 10% |3-mercaptoethanol (|3ME) for 5 min at 100°C. Electrophoresis was conducted at constant 200V. Bovine serum albumin (BSA)-substrate zymography was performed by preparing the gels with BS A-polyacrylamide conjugate as described by Grenier et al. (84) . For gelatin-substrate 35 zymography, the gels were prepared by copolymerizing the acrylamide with non-conjugated gelatin (final concentration, 1 mg/ml) described by Banda et al. (13). After electrophoresis of NR samples, the gel was gently shaken in 100 mM Tris-HCl (pH 7.5) containing 2% (vol/vol) Triton X-100 for 30 min, rinsed twice with water, and then shaken for a further 30 min in 100 mM Tris-HCl (pH 7.5). The gel was then transferred into development buffer, which unless noted otherwise, was 100 mM Tris-HCl (pH 7.5) containing 50 mM cysteine. The gel was incubated at 37°C for 2 h and stained for protein with Coomassie brilliant blue R-250. After the gel was destained, proteolytic activity was visualized as a clear band against a dark blue background. To study the effect of protease inhibitors on the recombinant protease activity, each inhibitor was added into the development buffer at the final concentration as follows: iodoacetate (8 mM), A/-a-p-tosyl-L-lysine chloromethyl ketone (TLCK) (4 mM), N-tosyl-L-phenylalanine chloromethyl ketone (TPCK) (4 mM), phenylmethylsulfonyl fluoride (PMSF) (4 mM), EDTA (40 mM), CaCl2 (10 mM), MgCl2 (10 mM), and ZnCl2 (10 mM). For two-dimensional PAGE (2D-PAGE), samples were prepared by boiling in 5% (vol/vol) Nonidet P-40 for 5 min, followed by solubilizing in lysis buffer containing urea and Nonidet P-40 as described by O'Farrell (205). Proteins were separated in the first dimension by isoelectric focusing in mini-tube gels containing 4% ampholytes in the pH range 5 to 8 and 1% ampholytes in the pH range of 3 to 10, and in the second dimension by SDS-PAGE using 10% separation gels as described in the BioRad mini-2D instruction manual (BioRad Laboratories, Richmond, Calif.). Proteins were detected by staining with silver nitrate (206). 6. Antiserum. Polyclonal antisera to whole cells of P. gingivalis and to the inclusion bodies in the E. coli clone containing the high-level expressed recombinant protein were generated by immunizing female New Zealand white rabbits. The first dose of antigen was administered with complete Freund's adjuvant, and 2 weeks later with a second dose in incomplete Freund's adjuvant. Subsequent injection of antigen was done without adjuvant. An IgG fraction was isolated from whole serum by affinity chromatography on a protein A Sepharose CL column (242). 36 7. Affinity purification of monospecific antibodies. Monospecific antibodies to the cloned protease were isolated by affinity purification with antigen immobilized on a nitrocellulose membrane in a modification of the method described by Smith and Fisher (288). Cell lysate of E. coli DH5cc (pT7-YS/pGPl-2) containing highly expressed recombinant protein (see section 11) was resolved by SDS-PAGE (10% polyacrylamide). Separated proteins were transferred electrophoretically to nitrocellulose membranes, and a strip from each side of the membrane was stained for proteins with Coomassie brilliant blue R-250. The strips were then realigned with the remaining membrane and used as templates to excise a horizontal strip containing the immobilized recombinant protein. The strip containing antigen was treated with blocking buffer (2% [wt/vol] BSA in Tris-buffered saline [TBS; 50 mM Tris-HCl (pH 8.1), 150 mM NaCl]) containing 0.02% (wt/vol) sodium azide and then incubated for 2 h at room temperature with antiserum to P. gingivalis whole cells. Following several washes, the bound antibodies were recovered by incubation with 1 ml of elution buffer (50 mM glycine, 0.5 M NaCl, 0.5% [vol/vol] Tween 20,100 fig of BSA per ml [pH 2.3]). The elute was immediately neutralized by the addition of an equal volume of 100 mM Na2HP04. 8. Western blot immunoassay. Western blot immunoassays were done by the method of Renart and Sandoval (226). Proteins resolved by SDS-PAGE were transferred electrophoretically to nitrocellulose. The blots were incubated for 1 h with blocking buffer (3% BSA in TBS [20 mM Tris-HCl (pH 7.5), 0.5 M NaCl]), and then incubated for 2 h at room temperature or overnight at 4°C with the primary antibody diluted in 1% BSA-TBS. Following two 10-min washes with 0.05% (vol/vol) Tween 20 in TBS, the blots were incubated for 1 h at room temperature with the secondary antibody, goat anti-rabbit IgG (H+L) alkaline phosphatase conjugate (human-adsorbed) (BRL, Gaithersburg, Md.), diluted in 1% BSA-TBS. Immunoreactive bands were developed with a solution containing BCIP and NBT. 37 9. In vitro translation. In vitro translation (222) was done with a prokaryotic DNA-directed translation kit (Amersham Co., Arlington Heights, 111.). The transcription and translation of 5 |ig of the purified plasmid pYS307 were done using [35S]-methionine as the radiolabel, as described by the supplier. Incubations were at 37°C for 1 h with [35S]-methionine. Nonradiolabeled L-methionine was then added, and the mixture incubated for another 5 min to allow the completion of protein chains which may have been terminated prematurely due to the limited amount of radioactive methionine. The radiolabeled gene products were separated by SDS-PAGE (10% polyacrylamide) as described above, and the gels were exposed to X-ray film. 10. Synthetic oligonucleotides and PCR. The following single-stranded oligonucleotide primers were synthesized by the Oligonucleotide Synthesis Laboratory, University of British Columbia: primer ECO (5!-GCGAATTCTAGTACCGCAATCCATTTCA-3'), primer BAM C5'-GCGGATCCATGGTT-AGAGCCTGTTGCC-3'). The underlined nucleotides represent EcoRI and BcrniHl sites, respectively. Primer ECO conesponds to the region between nucleotides 209 and 228 in the sequence of the tpr gene (for the sequence, see Fig. 1.3); the 7 nucleotides on the 5' end provide an EcoRI site for the DNA fragment amplified by the polymerase chain reaction (PCR). Primer BAM is a sequence complementary to the region between nucleotides 1727 and 1750; the additional 3 nucleotides provide a BamUI site to the amplified DNA fragment (Fig. II. 1, B). The 1.9-kb HincR-Hindm fragment of pYS307-4(+), containing the protease gene, was used as a template. After digestion of pYS307-4(+) with Hindi and HindBI, the 1.9-kb fragment was purified from the agarose gel using glass beads (GeneClean; Bio 101, La Jolla, Calif.). PCR was performed with the Twin Block System (EriComp, Inc., San Diego, Calif.) as described by Sambrook et al. (242). The reaction mixture (100 |0.1) contained 100 pmol of the primers ECO and BAM, all four deoxynucleoside triphosphates (1.25 mM each), 50 mM KCl, 10 mM Tris-HCl (pH 8.0), 1.5 mM MgCl2, 0.01% gelatin, and 3 U of Thermus aquaticus (Taq) DNA polymerase (Applied Biosystems, Inc., Foster City, Calif.). 38 Hindi CGATTCGAACTTCTCGATTCGAACTTCTGATAGACTTCGAAATTAATACGACTCACTATAGGGAGACCACAACC3GTTTCCCTCTAGAAATAATT M A R I R A R G S S R V D L Q P K L I I D TTGTTTAAACTTTAAGAAGGAGATATACATATGGCTAGAATTCGCGCCCGGGGATCCTCTAGAGTCGACCTGCAGCCCAAGCTTATCATCGAT rbs EcoKl Bam HI (B) M E K K L V P Q S I S 5' TTATGGAAAAGAAATTAGTACCGCAATCCATTTCA---3 ' AATACCTTTTCTTTAATCATGGCGTTAGGTAAAGT- — 5 ' GCGAATTCTAGTACCGCAATCCATTTCA 3 ' Primer BAM Y M Y • 3' CCGTTGTCCGAGATTGGTACCTAG£CG 5' -TATATGTATTGA GGCAACAGGCTCTAACCATGGATC 3 ' - ATATACATAACT CCGTTGTCCGAGATTGGTACCTAG 5 ' Primer ECO pYS307-2(+) I PCR I L V P Q S Y M Y . 5' GCGAATTCTAGTACCGCAATCC TATATGTATTGA AACCATGGATCCGC 3' 3' CGCTTAAGATCATGGCGTTAGG ATATACATAACT TTGGTACCTAGGCG 5' EcoW. _ _ „ , Bam HI PCR product Digestion with Eco RI & BamHl Ligation with pT7-7 digested with Eco RI&BamHl M A R I L V P Q S Y M Y -5 ' ATGGCTAGAATTCTAGTACCGCAATCC TATATGTATTGA AACCATGGATCCTCTACA 3 ' 3 ' TACCGATCTTAAGATCATGGCGTTAGG ATATACATAACT TTGGTACCTAGGAGATGT 5 • EcoW BamHl pT7-YS FIGURE II.1. Construction of pT7-YS. In the DNA nucleotide sequence, boldface type represents Bam HI or EcoRI sites. The arrows indicate the direction of the transcription. (A) Structure of the T7 expression vector plasmid pT7-7. (B) Scheme for the construction of pT7-YS. 39 The PCR product was purified by chloroform extraction followed by ethanol precipitation and was identified on agarose gels after electrophoresis. 11. Construction of pT7-YS and high-level expression of tpr. The 1.5-kb DNA fragment amplified by PCR was digested with EcoRI andBamBI and cloned into the EcoRI-Bam HI region of the T7 expression plasmid pT7-7, constructed by Dr. S. Tabor, Dept. of Biological chemistry, Harvard Medical School, Boston, Mass. (see Fig. 11. 1, A). The subcloned plasmid, named pT7-YS, was transformed into E. coli DH5oc harboring pGPl-2 (307) . The pGPl-2 contains the gene for T7 RNA polymerase under the control of the Xp-^ promoter that is repressed by a temperature-sensitive repressor, cI857. The two compatible plasmids were maintained in the same cell by selection with kanamycin (pGPl-2) and ampicillin (pT7-YS). For high-level expression, E. coli DH5a (pT7-YS/pGPl-2) was grown to an ODgoo of 0.4 to 0.5 at 30°C and then incubated at 42°C for 30 min to induce expression of T7 RNA polymerase in pGPl-2. After induction, rifampicin (final concentration, 200 |i.g/ml) was added to the culture to inhibit the synthesis of E. coli proteins. The cells were incubated at 37°C for another 3 h to allow for the expression of the cloned gene regulated by T7 promoter. Whole-cell proteins were analyzed by SDS-PAGE and Western blot immunoassay. 12. NH2-terminal amino acid sequence. The NH2-terminal amino acid sequence of the gene product was determined by the method of Matsudaira (177). Briefly, a protein sample containing the remobinant protein (rTpr) from high-level expression system was separated by SDS-PAGE, and proteins were transferred electrophoretically to Immobilon-P (Millipore Co., Bedford, MA). After staining with Coomassie brilliant blue R-250, the membrane was destained, and washed with high-performance liquid chromatography grade distilled water. The protein band of interest was excised from the blot, dried, and then subjected to automated stepwise sequencing on an Applied Biosystems 474A gas-phase sequencer at the University of Victoria (BC, Canada). 40 13. Construction of pBY2-IN. A wide-host range cosmid vector pJRD215 (47), which can be mobilized into, but cannot replicate in P. gingivalis (262), was chosen as the suicide vector. The erythromycin resistance (Emr) gene of pNJR5 (262) (derived from Tn4400 of Bacteroides fragilis [229, 312]) was used as the selection marker for P. gingivalis transconjugants, respectively. A suicide plasmid containing the disrupted tpr gene, pBY2-IN was constructed as shown in Fig. n.2. The 2.4-kb BamHL-Hinffil fragment of pYS307-2(+) was cloned into pJRD215 to construct pBY2. Plasmid pBY407-EM was constructed by cloning the 1.0-kb BamUl-Pstl fragment of pYS307-2(+) and the 3.8-kb Emr fragment of pEM (pEM, 3.8-kb EcoW-SacI fragment of pNJR5 containing the Emr gene cloned into pUC18) into a plasmid vector pBluscriptll SK(-). The 4.8-kb BarnHl-Kpnl fragment of pBY407-EM was cloned into pBY2, resulting in the replacement of the 0.7-kb Pstl-Kpnl fragment in the middle of the tpr gene with the 3.8-kb Emr fragment. 14. Construction of pBY3. The shutde vector plasmid developed for Bacteroides spp., pNJR12 (262), was chosen as the base on which to construct the shuttle plasmid containing the tpr gene. This plasmid consists of a wide-host-range cosmid vector pJRD215, a Bacteroides spp. plasmid pB8-51 (264, 265) and 2.7-kb fragment containing tetracycline resistance (Tcr) gene from a conjugal element of Bacteroides fragilis. The 2.4-kb BamHl-Hindlll fragment of pYS307-2(+) was subcloned into pNJR12, and the recombinant plasmid named pBY3 (Fig. II.3). Expression of the recombinant Tpr in E. coli (pBY3) was determined by gelatin-substrate zymography and Western immunoblot analysis. 15. Conjugation. Conjugation between E. coli strains to introduce R751 into E. coli containing pBY2-IN or pBY3 was performed as described by Dr. N.B. Shoemaker (University of Illinois, Urbana) (personal communication). Cells were grown to OD6oo=0.2 to 0.3 in LB broth containing 41 FIGURE H.2. Construction of pBY2-IN. The black section and the stippled section represent the DNA insert of the original clone and the Emr fragment, respectively. Abbreviations for restriction endonucleases: Bm, BamFLI; He, HincE; Ps, Pstl; Sp, Sphl; Kp, Kpnl; Hd, HindBI; Sa, Sail; Xb, Xbal; Ec, EcoM; CI, Clal 42 FIGURE H.3. Construction of pBY3. The black section represents the DNA insert of the original clone and the shadowed section Tcr fragment containing the tetracycline resistant gene. Abbreviations for restriction endonucleases: Bm, BamHl; He, Hindi; Ps, Pstl; Sp, Sphl; Kp, Kpnl; Hd, Mncffll; Ss, Sstl; Xb, Xbal; Ec, EcoRI; CI, Clal 43 antibiotics, Tp (200 Ug/ml) for R751 and Km (50 jig/ml) for pBY2-IN and pBY3. 0.5 ml of the donor cells and 0.5 ml of the recipient cells were mixed, harvested by centrifugation, resuspended in 0.1 ml of LB broth, and then spotted on an LB agar plate without antibiotics. After overnight incubation, colonies were streaked on LB agar plates containing Tp (200 |i.g/ml) and Km (50 |J.g/ml). Isolated colonies were restreaked on the plate containing Tp and Km to be sure that transconjugants contained both plasmids. pBY2-IN was mobilized from E. coli to P. gingivalis W83 by conjugation using the broad host-range vector R751 to provide transfer function, following a modification of the method described by Maley et al. (171). Briefly, E. coli (R751/pBY2-IN) and P. gingivalis W83 were grown to early-exponential phase (OD6oo=0.2). 0.2 ml of E. coli DH5a (R751/pBY2-IN) and 1 ml of P. gingivalis were mixed, and the cell mixture was harvested by centrifugation at 9,600 Xg for 2 min. The cells were resuspended in 0.1 ml of LB broth. The cell mixture was spotted onto a sterile Millipore filter (HAWP 013 00, Millipore Co., Bedford, MA) on a BHI agar plate containing 5% human blood. The plate was incubated aerobically at 37°C for 2 h, and then anaerobically at 37°C for 72 h. The filter was removed from the agar plate, and cells on the membrane were resuspended in 3 ml of BHI broth. The cell suspension was plated onto BHI-blood agar plates containing 20 (J,g/ml erythromycin and 200 |J.g/ml gentamycin, and were incubated anaerobically at 37°C for 2 weeks. A modification of the method described above was used to introduce the shuttle plasmid pBY3 into P. gingivalis W83/PM. The cell mixture, prepared as described in the previous paragraph, was spotted onto the filter on a BHI-blood agar plate, incubated aerobically overnight, and then anaerobically for 24 h. The cells on the filter were resuspended in 3 ml of BHI broth and then incubated overnight under anaerobic conditions before plating onto the antibiotic-containing medium. 16. Cellular fractionation. 44 Whole cell extracts, soluble cell extracts and crude membrane fractions of P. gingivalis were prepared by French pressure cell disruption followed by differential centrifugation. Cells cultivated for 48 h were harvested and washed once with TM buffer (100 mM Tris-malate [pH7.5], 10 mM CaCl2, 10 mM NaCl). Cells were resuspended in TM buffer containing 10 mM EDTA, 10 mM MgCl2, 20 mg/ml RNase A and 20 mg/ml DNase I, and broken by five passes through a cold French pressure cell at 20,000 psi (Ambicon French Pressure Cell, SLM Instruments Inc., Urbana, 111.) The broken cell suspensions were subjected to low speed centrifugation (6,000xg for 10 min at 4°C) to pellet whole cells and cell debris. The supernatant was designated as the whole cell extract (WCE). Crude membrane fractions and soluble cell extract (SCE) were prepared by centrifugation of the WCE at 100,000Xg for 2 h at 4°C. Membrane fractions were resuspended in TM buffer. All fractions were stored frozen at -20°C. Protein concentration was determined with the BioRad protein assay (BioRad Laboratories, Hercules, Calif.). 17. Pz-peptidase assay and activity against other protease substrates. The Pz-peptidase assay was done using a bacterial collagenase substrate, p-phenylazobenzyloxycarbonyl-L-prolyl-L-leucyl-glycyl-L-prolyl-D-arginine (Pz-peptide) (350). 200 H-l of the enzyme preparation containing 200 jug of protein was added to 1 ml of assay solution (100 mM Tris-malate [pH7.5], 10 mM CaCl2, 10 mM NaCl, 60 mM cysteine, 1 mg/ml of substrate). After incubation at 37 °C for 1 h, the reaction was stopped by addition of 1 ml of a 0.5% (wt/vol) citric acid solution. 4 ml of ethyl acetate was then added, vigorously mixed for 15 seconds, and allowed to settle until clear. 3 ml of the upper ethyl acetate phase was taken and transferred to tubes containing 1 g of Na2S04. Enzyme activity was determined by measuring absorbance at 320 nm (A320). Boiled enzyme preparations were used as negative controls for each reaction. Hydrolysis of BApNA, glycyl-L-arginine p-nitroanilide (GApNA) and N-succinyl-L-alanyl-L-alanyl-L-prolyl-L-phenylalanine p-nitroanilide (SAAPpNA) was determined by 45 incubating samples at 37 °C with the chromogenic substrate (final concentration, 2 mM) in the reaction mixture described above. Enzyme activity was determined by measuring absorbance at 410 nm (A410) after the incubation. 18. Effects of protease inhibitors and metal ions on Pz-peptidase activity. The effects of various compounds on the Pz-peptidase activity were tested at final concentrations of: 0.5, 5 and 10 mM for TLCK; 0.5 and 5 mM for TPCK, PMSF, p-chloromercuribenzoate (pCMB), iodoacetate, 4-aminophenylmercuric acetate (APMA) and N-ethylmaleimide (NEM); 5 and 50 mM for EDTA; 5 and 50 |ig/ml for pepstatin and leupeptin. Various metal ions were tested at 1 to 20 mM. The enzyme preparation was added to the assay solution containing the appropriate compound, and enzyme activities determined as described above. The activity assay with metal ions was performed using a crude membrane fraction of P. gingivalis W83 which was prepared with TM buffer without added CaCl2. 19. Effects of pH on Pz-peptidase activity. The reaction mixture for determination of Pz-peptidase activity at different pH values consisted of 200 |il of enzyme preparation and 1 ml of an appropriate buffer (100 mM phosphate buffer for pH 6.0 to 7.0; 100 mM Tris-HCl buffer for pH 7.0 to 9.0; 100 mM glycine buffer for pH 9.0 to 10.5) containing 60 mM cysteine and 1 mg/ml of substrate. The enzyme assay was performed as described above. The pH value of each reaction mixture was remeasured after mixing all components. 20. CoIIagenase assay with [14C]-labeled soluble collagen. The collagenase assay was performed using biologically [14C] -labeled type I chick embryo collagen (kindly provided by Dr. J.-V. Uitto, Dept. of Oral Biology, U.B.C.) by a modifi-cation of the method originally described by Terato et al. (317). A standard reaction mixture consisted of 0.2 ml of [14C]-labeled collagen (2 mg/ml, approximately 10,000 dpm) in TM buffer supplemented with 50 mM dithiothreitol (DTT) and aliquots of the crude membrane fraction containing 40 |ig of proteins. The final volume was made up to 0.4 ml with the same 46 buffer. After incubation at 25°C for 2 h, the reaction was stopped by adding APMA (final concentration, 5 mM). Degradation products were extracted by mixing with 0.4 ml of dioxane. Undegraded collagen molecules were separated by centrifugation at 10,000xg for 20 min. Aliquots (0.3 ml) of the supernatant were transferred into scintillation vials containing liquid counting scintillant (Scinitisafe 30% for aqueous samples, Fisher Scientific, Fair Lawn, NJ). CoUagenolytic activity was determined by calculating the percentage of radioactivity extracted from reaction mixture. 21. Hemagglutinating activity. Hemagglutinating activity was assessed in U-bottom polystyrene microtitre plates. Human erythrocytes were washed with phosphate-buffered saline (PBS) and resuspended at a concentration of 2% (vol/vol) in the same buffer. Bacterial cells were harvested, washed, and resuspended in PBS to an optical density between 1 and 2, at 660 nm. Equal volumes (100 til) of erythrocytes and bacterial cells were added to the well of a microtitre plate. The samples were gently shaken for 20 min at room temperature, and allowed to sit for 1 h before scoring for hemagglutination. To determine a hemagglutination titer for a particular sample, the bacterial cell suspension was diluted in a two-fold series with PBS, and 100 ja.1 of each dilution was analyzed for hemagglutinating activity. The titer for a particular cell strain was taken as the reciprocal of the highest dilution showing positive agglutination. 22. Virulence test. Cultures in TYE broth were harvested by centrifugation (10,000 Xg, 30 min) and washed with PBS. The cells were resuspended in PBS to 5 X 1010 total cells per ml. Cell numbers were determined by measuring optical density at 600 nm after a 100 fold dilution and by using a standard equation ([X 108] cells/ml = 15.5 X OD6oo)- Viable cell numbers were determined after serial dilution on BHI-blood agar plates. A 0.3 ml suspension was injected subcutaneously in the abdominal area of mice. PBS without bacteria was used as a control inoculum. The mice were then examined three times per day for health status and the 47 presence of lesions at the injection site. Virulence tests were performed 3 times with a total of 20 mice for each strain. 48 HI. RESULTS 1. Cloning of the tpr Gene. 1.1. Isolation of a protease positive clone. A genomic library of P. gingivalis W83 was generated in E. coli JM83 as described in Materials and Methods. Screening for a protease positive clone involved looking for zones of clearing around colonies growing on skim-milk agar plates. Screening 2,000 recombinant clones yielded a single colony capable of hydrolyzing milk proteins. The zone of clearing appeared deep in the skim-milk agar plate only after 2 weeks of incubation at 37 °C. Plasmid isolated from this clone was transformed into E. coli JM83 to confirm that the proteolytic activity of the clone was expressed by a cloned gene. It was observed that colonies which grew deep within the agar developed zones of protein hydrolysis, while those inoculated on the agar surface did not. This suggested that the protease was only active under conditions where exposure to oxygen was minimized, i.e. conditions that would exist deep in the agar. To confirm the requirement of the reducing conditions for activity of the cloned protease, the transformants were incubated in an oxygen-free environment. 100 of the transformants cultivated on skim-milk-containing agar media were transferred into an anaerobic chamber after treatment with chloroform vapor, and incubated for several days. Under these conditions, clear zones appeared around all the colonies after 3 days. Fig. III. 1 shows the milk-protein hydrolyzing activity of the clone when incubated under different environments. Cells grown aerobically on the surface of skim-milk-containing agar media were treated with chloroform vapor and incubated under aerobic or anaerobic conditions. The clone incubated under anaerobic conditions produced a proteolytic clear zone (plate B), while the clone incubated under aerobic conditions did not (plate A). Even without treatment with chloroform vapor, the clone produced zones of clearing under anaerobic conditions (plate C). It was believed that the proteolytic activity in plate C was due to cell 49 FIGURE m. l . Detection of proteolytic activity of the clone on LB agar plates supplemented with skim milk. YS and TZ represent the protease positive clone and E. coli JM83 (pTZ18R), respectively. The plates were incubated aerobically overnight for cells to grow, and then incubated further under aerobic conditions (plate A) or under anaerobic conditions (plates B, C). Plates A and B were treated with chloroform for 30 min prior to the incubation. Plate C was not treated with chloroform. 50 lysis during incubation. E. coli JM83 containing pTZ18R did not show proteolytic activity under any conditions studied. 1.2. The recombinant plasmid. The recombinant plasmid pYS307 isolated from the protease-positive clone contained a 5.2-kb insert. A restriction endonuclease map was generated by analyzing the restriction patterns of pYS307 after single or double digestions with various restriction endonucleases (Fig. III.2). The DNA insert contained single cleavage sites for AccI, Hindi, Kpnl, Pstl, SacI and Sphl, and multiple cleavage sites for BamHI, EcoW and HindSI. 1.3. Southern hybridization. The origin of the cloned DNA fragment was confirmed by Southern hybridization analysis using the 3.2 kb-Hindlll fragment located in the middle of the insert of pYS307 as a probe. Chromosomal DNA isolated from/', gingivalis W83 digested with Hindm possessed a single 3.2 kb fragment which hybridized with the probe (Fig. III.3, lane 2), showing that the cloned protease gene could be found in the genomic donor. Chromosomal DNA samples of several different P. gingivalis strains and other species formerly known as BPB were assayed by Southern hybridization to observe the species-specificity of the cloned DNA. As shown in Fig. III.3, chromosomal DNA isolated from P. gingivalis ATCC 33277, W50 (ATCC 53978) and W12 hybridized with the probe (lanes 3 to 5). In the case of P. gingivalis ATCC 33277 (lane 3), the probe hybridized with a 5-kb fragment, unlike the other strains which produced the 3.2-kb hybridizing band as P. gingivalis W83. No hybridizing band was detected with chromosomal DNA samples of other BPB species: P. asaccharolytica, Prevotella corporis, P. denticola, P. intermedia, P. loescheii, P. melaninogenica, and Bacteroides levii (lanes 6 to 12). This suggests that the HindM fragment containing the cloned protease gene is species-specific, when assayed under high stringency washing conditions. 51 FIGURE m.2 . Restriction endonuclease map of the recombinant plasmid, pYS307. The black thick arc represents the insert DNA fragment, and the remainder represents the vector plasmid pTZ18R. 52 FIGURE m.3. Southern hybridization. A 3.2-kb HindEI fragment of pYS307 was used as a probe. Lane 1 is pYS307 digested with Mndin. All of the chromosomal DNA samples in lanes 2 to 12 were digested with HindJR. Lanes: 2, P. gingivalis W83; 3, P. gingivalis ATCC 33277; 4, P. gingivalis W50 (ATCC 53978); 5, P. gingivalis W12; 6, P. asaccharolyticus ATCC 25260; 7, Prevotella corporis ATCC 33547; 8, P. denticola ATCC 33185; 9, P. intermedia ATCC 15032; 10, B. levii ATCC 29147; 11, P. loescheii ATCC 15930; 12, P. melaninogenica ATCC 25845. 2. Identification of the Cloned Gene Product. 2.1. BSA-substrate zymography. Protein profiles of cell lysates in Coomassie blue-stained gels did not reveal significant differences between E. coli JM83 (pTZ18R) and the recombinant clone E. coli JM83 (pYS307) (data not shown). However, the enzymatic activity associated with the cloned gene was detected by BSA-substrate zymography. A proteolytic band migrating with an approximate molecular mass of 90 kDa was identified in the recombinant clone (Fig. IH.4, lane YS). Activity was observed when whole cell lysates were prepared in the absence of fJME. No proteolytic activity could be detected with sample prepared in the presence of 10% (5ME in the solubilization buffer (data not shown). In lanes containing cell lysates of E. coli JM83 (pTZ18R), several host proteins appear as Coomassie blue-stained bands (lane TZ). These bands disappeared in lanes containing cell lysates of the clone (lane YS). This suggests that the cloned protease degrades E. coli proteins during the lysis of whole cells in sample solubilization buffer. The proteolytic band at 90 kDa was detected only when a reducing agent, cysteine (50 mM), was added to the assay system following electrophoresis. This result suggests that the cloned protease is thiol-dependent, and confirms the results observed in the colony screening assay. 2.2. Western immunoblot analysis. The identity of the cloned P. gingivalis protein was determined by immunoblotting with antiserum raised against P. gingivalis W50 whole cells (Fig. III.5). The antiserum reacted with several E. coli proteins; however, the recombinant protein could be identified by comparing the clone and the control strain containing pTZ18R. A single reactive band of 90 kDa was detected with nonreduced-nondenatured sample of the clone (panel NR, lane YS). The position of the reactive band was identical to the proteolytic band on the BSA-substrate zymogram shown in Fig. III.4. A different result was observed with a reduced-denatured sample. In this case, the antibody reacted with a peptide of 50 kDa and a 38-kDa peptide 54 FIGURE m.4 . Detection of proteolytic activity by BSA-substrate zymography. Lane TZ, E. coli JM83 (pTZ18R); lane YS, E. coli JM83 (pYS307). Samples were nonreduced-nondenatured whole-cell lysate. After electrophoresis, SDS was removed from the gel by incubating with 100 mM Tris-HCl (pH 7.5) containing 2% Triton X-100. The gel was washed, and incubated at 37°C for 2 h in 100 mM Tris-HCl (pH 7.5) containing 50 mM cysteine. The incubated gel was stained with Coomassie brilliant blue R-250 and destained. 55 i R NR TZ YS YS TZ FIGURE m.5. Identification of the cloned gene product by Western immunoblot analysis. Samples in panels R and NR were reduced-denatured, and nonreduced-nondenatured whole-cell lysates, respectively. Nonreduced-nondenatured and reduced-denatured samples were solubilized in sample solubilization buffer (see Materials and Methods) for 30 min at 37 °C and in buffer containing 10% (3ME for 5 min at 100°C, respectively. Lanes: TZ, E. coli JM83 (pTZ18R); YS, E. coli JM83 (pYS307). Antiserum raised against P. gingivalis W50 whole cell was used as a primary antibody. 56 (panel R, lanes YS). The 38 kDa was not always detected, and is believed to be a degradation product of the recombinant protein. These results suggest that the recombinant 90-kDa active protease may be composed of a subunit having a molecular mass 50 kDa. 2.3. In vitro translation. Western immunoblot analysis suggested that the recombinant 90-kDa protease contained a subunit having a molecular mass of 50 kDa. To confirm this suggestion and identify the cloned gene product, in vitro translation was done and the electrophoresed products were analyzed by autoradiography. As shown in Fig. III.6, in the in vitro translation system, pYS307 specifically expressed a protein having a molecular mass of 50 kDa (lanes R-YS and NR-YS). The 90-kDa protein was not detected in nonreduced-nondenatured samples. In the in vitro system, the protein was apparently not processed to the proteolytically active 90-kDa form. The vector plasmid, pTZ18R, expressed two proteins with approximate molecular masses of 32 kDa and 20 kDa (lanes R-TZ and NR-TZ). The 32-kDa protein is believed to be p-lactamase (Apr gene product) and the 20-kDa protein is the a fragment of (3-galactosidase (lacZ' gene product). No 20-kDa protein could be observed in pYS307, due to the inactivation of lacZ' gene by the inserted DNA fragment. Lower-molecular-mass bands around the proteins were observed, and were believed to be produced by premature termination of translation and/or aberrant initiation at the internal AUG codon (11,222). 3. Characterization of the Cloned Gene and its Gene Product. 3.1. Subcloning of the insert. 1) Localization of the cloned gene. Subclones were constructed on the basis of the restriction map of pYS307 (Fig. III. 2). The plasmid vector pTZ19R (185) was used to construct subcloned plasmids which had the insert in a particular orientation with respect to the lac promoter. pTZ19R differs from pTZ18R only in the orientation of the multiple cloning site. Expression of protease activity 57 R NR kDa 2 2 4 -1 0 9 -7 2 -46 -TZ YS TZ YS 1 8 -1 5 -FIGURE m.6. Autoradiogram of the proteins expressed in the in-vitro translation system. Lanes: TZ, proteins encoded in the vector plasmid pTZ18R; YS, proteins encoded in the recombinant plasmid pYS307. R represents reduced-denatured protein samples; NR represents nonreduced-nondenatured samples. 58 by the subclones was determined by observing proteolytic clearing in skim-milk agar plates. As shown in Fig. III.7, the cloned gene was localized to the 1.9-kb DNA fragment between the Hindi and Mndffl sites (the subcloned plasmids, pYS307-4(+) & -4(-)). The orientation of the 1.9-kb fragment did not influence expression of the protease suggesting that the DNA fragment contains a promoter region. 2) Zymography and Western immunoblot. The subclones showing protease activity on skim-milk plates also produced a proteolytic band with a molecular mass of 90 kDa in BSA-substrate zymography which is identical to that of the original clone (Fig. 111.8). The protease-positive subclones had the same immunoblot patterns as the original clone, when assayed against the antiserum raised to P. gingivalis W50 whole cells (Fig. III.9). The antiserum reacted with a single 90-kDa protein in the nonreduced-nondenatured samples (panel NR) and a 50-kDa protein in the reduced-denatured samples (panel R). The subcloned plasmid, pYS307-3(+) and -3(-), expressed a 30-kDa protein which reacted with the antiserum (lane 3(+) and 3(-)). The protein was believed to be a truncated form of the 50-kDa protein, suggesting that the promoter of the cloned gene is localized to the region between Hindi and Pstl sites, and the gene is transcribed in the direction as shown in Fig. m.7. The E. coli strain containing pYS307-5(+) or -5(-) did not produce any band reacting with the antiserum (data not shown). 3.2. Effects of protease inhibitors and substrate specificity. Several protease inhibitors of known specificity were incorporated into the BSA-substrate zymography assay to determine their effect on the 90-kDa protease. The cloned protease was inhibited by iodoacetic acid, TLCK, TPCK, PMSF, EDTA and ZnCl2, but not by CaCl2 and MgCl2 (data not shown). Substrate specificity was assessed by measuring the hydrolysis of synthetic peptides including BApNA, SAAPpNA, GApNA and Pz-peptide. The source of enzyme was the cell lysate of the clone E. coli JM83 (pYS307). The lysates did not hydrolyze any of substrates, 59 pYS307 pYS307-l(+) & -l(-) Bm Hd Ec Ac Bm Bin He Ps Kp Hd Sc Bm 3d Sp Ec pYS307-2(+) & -2(-) pYS307-3(+) & -3(-) pYS307-4(+) & -4(-) (30kl)a) + pYS307-5(+) & -5(-) |< l k b >1 Protease Activity FIGURE m.7. Localization of the cloned gene. The lines represent the DNA fragments inserted into the vector plasmids, pTZ18R or pTZ19R. (+) and (-) indicate the right and opposite orientations, respectively, with respect to the lac promoter. The orientation of the original insert of pYS307 was defined as the right orientation. Abbreviations are as follows: Ac, AccI; Bm, BamBI; Ec, Eco RI; He, HincU; Hd, //indm; Kp, Kpnl; Ps, Pstl; Sc, Sad; Sp, Sphl. A hatched arrow line represent the possible location and transcription direction of the gene. 60 FIGURE HL8. Detection of proteolytic activities of the subclones by BS A-substrate zymography. Lanes: TZ, E. coli JM83 (pTZ18R); YS, E. coli JM83 (pYS307); 1(+), E. coli JM83 (pYS307-l(+)); 1(-), E. coli JM83 (pYS307-l(-)); 2(+), E. coli JM83 (pYS307-2(+)); 2(-), £. coli JM83 (pYS307-2(-)); 3(+),E. co/i JM83 (pYS307-3(+)); 3(-), E. coli JM83 (pYS307-3(-)); 4(+), £. co/i JM83 (pYS307-4(+)); 4(-), E. coli JM83 (pYS307-4(-)). 61 4 FIGURE in.9. Western immunoblot of the subclones. NR represents nonreduced-nondenatured protein samples; R represents reduced-denatured protein samples. Lanes: see the legend of Fig. IH.8. Antiserum raised against P. gingivalis W50 whole cell was used as the primary antibody. 62 although the enzyme did hydrolyze casein, BSA and gelatin. Casein hydrolyzing activity was detected by observing a clear zone on skim-milk plates. BSA- and gelatin-substrate zymography studies showed the capability of the recombinant protein to hydrolyze these proteins. 3.3. The tpr gene. Subsequent to our initial study [Y. Park and B.C. McBride (1992) FEMS Microbiol. Lett. 92: 273-278], Bourgeau et al. (23) reported the cloning, expression, and sequencing of a protease gene (tpr) of P. gingivalis W83. The restriction endonuclease map of the tpr gene was identical to the results obtained in this study. The location of the cloned gene in our restriction map was also identical to the open reading frame region of the tpr gene (Fig. 1.3). The sensitivities of both gene products to protease inhibitors were similar to each other. These suggested that both research groups had cloned the same gene; thus, the cloned protease gene will henceforth be called tpr. The nucleotide sequence of tpr reported by Bourgeau et al. (23) was analyzed further using the nucleic acid and protein sequence analysis software systems, DNA Strider™ 1.0 (Commissariat a L'Energie Atomique, France) and PC/Gene (University of Geneva, Switzerland). The molecular mass of the protein encoded by tpr was shown to be 55 kDa when calculated from the amino acid sequences deduced from the nucleotide sequence by using both sequence analysis systems. This was similar to the molecular mass of the denatured recombinant protein determined in SDS-PAGE analysis. However, it differed from the molecular mass of 62.5 kDa reported by Bourgeau et al. (23). A hydropathic plot of the translated protein was obtained by Kyte-Doolittle analysis (Fig. III. 10). Approximately 30 residues in the middle of the deduced amino-acid sequence formed a hydrophobic region. The same region (residues 224 to 246) was also identified as a potential transmembrane helix by the secondary structure analysis of the PC/Gene system. A region between residues 386 and 406 was also predicted to be a membrane associated helix. 63 ON 4*. i I I I | 100 200 300 I I I I | I I I I | I I I I | I I I I | I I I I | I I I I I 400 I I I I | I I I I | I I I FIGURE m.10. Kyte-Doolittle hydropathy plot of the amino acid sequence (482 residues) deduced from the open reading frame of the P. gingivalis tpr gene. Hydrophobic regions are above, and hydrophilic regions below the central line. Numbers on the horizontal axis represent amino-acid residue number of the protein. The regions marked with "*" indicate potential membrane associated helices predicted by the PC/Gene sequence analysis system. 3.4. Sequence homology with calpains. The amino acid sequence deduced from the DNA sequence (Fig. LI) was analyzed to search for conserved sequences by the BLAST network services (National Center for Biotechnology Information, NIH). In addition to the consensus sequence of cysteine proteases found by Bourgeau et al. (23), three additional regions shown in Fig. IH.11 which were highly conserved (58 to 87%) to regions in the mammalian calpains (thiol-dependent calcium activated proteases) were identified. All of the conserved regions were located on domain II which is responsible for the thiol-dependent proteolytic enzyme activity of the calpains. The C terminal region (amino acid 219 to 445) of Tpr was aligned with the calpains shown in Fig. III. 11 using the computer sequence analyzing program (GeneWorks, IntelliGenetics, Inc. Mountain View, Calif.). As shown in Fig. III. 12, the C terminus region of Tpr could be aligned with the amino acid sequences of the domain II (cysteine protease domain) of various calpain large subunits without any significant deletions or insertions. The amino acid sequence comparison of Tpr in the region (amino acid 219 to 445) and the rat calpain (nCL-2) (292) showed 22% identity and 53% similarity. Tpr showed a high degree of homology in the regions suggested to contain active site residues (amino acids C, H and N, marked with arrow in Fig. III. 12). These support the results described earlier that Tpr is a thiol-dependent (cysteine) protease. 4. High-Level Expression of the Recombinant Tpr. 4.1. Construction of pT7-YS . The original recombinant clone E. coli DH5a (pYS307) expressed low levels of proteolytic activity and levels of protein which could not be detected by Coomassie-blue in SDS-PAGE. In order to increase the level of expression, the T7 RNA polymerase/promoter expression system (307) was adapted. The 1.5-kb DNA fragment containing the open reading frame of the tpr gene was amplified by PCR using the primers, ECO and BAM (Fig. III. 13, lane PCR), and subcloned into the T7 expression vector plasmid pT7-7 to construct pT7-YS, 65 Tpr Rat nCL-2 Rat nCL-2' RatnCL-1 Rat calpain II Human muCL Human muCL Chicken mCL Sol protein Tpr Rat nCL-2 Rat nCL-2' RatnCL-l(p94) Rat calpain II Human muCL Human muCL Chicken mCL Tpr RatnCL-2 RatnCL-2' RatnCL-l(p94) Rat calpain II Human muCL Human muCL Chicken mCL Sol protein 221 WQGIAGDCYMLAALSAI 238 345 IRQGGLGDCWLLAAIASL 392 331 IRQGGLGDCWLLAAIASL 378 662 ICQGDLGDCWLLAAIACL 715 401 ICQGALGDCWLLAAIASL 454 419 ICQGALGDCWLLAAIASL 472 462 ICQGALGDCWLLAAIASL 515 335 ICQGALGDCWLLAAIGSL 388 3483 ICQGVLGNCWLLSALAVL 3539 308 EYWPALFEQAYA 319 573 EFWSALLEKAYA 608 559 EFWSALLEKAYA 594 896 EFWSALLEKAYA 931 635 EFWSALLEKAYA 670 653 EFWSALLEKAYA 688 696 EFWSALLEKAYA 731 569 EFWSALLEKAYA 604 425 IRNPWGVTEPTG 436 900 LRNPWGEVEWSG 935 886 LRNPWGEVEWSG 921 13 67 LRNPWGQVEWNG 1402 962 IRNPWGQVEWTG 997 980 IRNPWGEVEWTG 1015 1023 MRNPWGEVEWTG 1058 896 IRNPWGQVEWTG 931 4059 LRNPWGHYSWRG 4094 Identity 50% 50% 44% 44% 44% 44% 44% 42% 66% 66% 66% 66% 66% 66% 66% 58% 58% 58% 75% 75% 67% 75% 50% Similarity 87% 87% 77% 83% 83% 83% 77% 84% 83% 83% 83% 83% 83% 83% 83% 75% 75% 67% 75% 75% 75% 75% 58% FIGUREIII.il. Amino-acid sequence homologies between Tpr and calpains. These include rat calpains (nCL-2 [D14479] and -2' [D14480]) (292), rat nCL-1 [J05121] (291), rat calpain II [L09120] (50), human muCL's [M23254 & X04366] (9, 111), chicken mCL [X01415] (209), and Drosophila melanogaster Sol protein [M64084] (49). The numbers in square parentheses refer to the GenBank accession number. The numbers at each end of the sequence represent residue numbers of amino acid or nucleotide. 66 Tor 219 DTWQGIAGDCYMLAALSAIAWVWPALL Consensus D * QG GDC *LAA***** LL Rat nCL-2 & nCL2• VWKRPTEL-CPNPQFIVGGATRT D-IRQGGLGDCWLLAAIASLT-LNEKLL Human muCL KWKRPTEL-LSNPQFIVDGATRT D-ICQGALGDCWLLAAIASLT-LNDTLL Chicken mCL EWKRPSEL-VDDPQFIVGGATRT D-ICQGALGDCWLLAAIGSLT-LNEELL Rat nCL-1 (p94) VWKRPPEI-CENPRFIIGGANRT D-ICQGDLGDCWLLAAIACLT-LNERLL D. melanogaster Sol protein QWRRPHEINCDGGAYPPWAVFRTPLPSD-ICQGVLGNCWLISALAVLA-EREDLV (from residue 1046) » NMDVDIMSNQDEWRLYRYFIGRSKQTYANRPSGSGTSTNEILQ-EGYYKVPIFARSRYWFNGEYWPALFEQAYANWKFPNDSKYN ******* ***Q **** ***** L * ** *p *g **E*W*AL E*AYA K ***y* YR VLPRDQSFQ--KDYAGIFHFQGWQYGEWVEWIDDRLP-TKNGQL-LFIHSEE--GNEFWSALLEKAYA--KL--NGSYE HR WPHGQSFQ- -NGYAGIFHFQLWQFGEWVDWVDDLLP - IKDGKL-VFVHSAE- -GNEFWSALLEKAYA- -KV- -NGSYE HR WPHGQSFQ- -EDYAGIGHGQIWQFGEWVDVWDDLLP-TKDGEL-LFVHSAE- -CTEFWSALLEKAYA- -KL- -NGCYE FR VIPHDQSFT--ENYAGIFHFQFWRYGDWVDWIDDCLP-TYNNQL-VFTKSNH--RNEFWSALLEKAYA--KL--HGSYE KE VLVTKEXCGQ GAYQVRLCKDGKWTTVLVDDLLPCDKRGHL-VYSQAKR--KQ-LWVPLIEKAVA--KI--HGCYE AILQIGGGWPEBALCELSGDSWFTSSGKLMLSSF--TDLSLLNFMKSMCY--SWK TIKPMVIVTPCWEPLPPMMPGI A** GG E* * *G* * K ** * * K* * *I * *** * ** G ALV GGSTIEGFEDFTGGISEFYDLKKPPENL- -YYIIQKALRKGSLL- -GC SIDVSTAAEAEATTRQKLVKG-^ ALS GGSTSEGFEDFTGGVTEWYELRKAPSDL- -YQIILKALERGSLL- -GC SIDISSVLDMEAITFKKLVKG-SLS GGSTTEGFEDFTGGVAEMYDLKRAPRNM- -GHIIRKALERGSLL- -GC SIDITSAFDMEAVTFKKLVKG-ALK GGNTTEAMEDFTGGVTEFFEIKDAPSDM--YKIMRKAIERGSLM--GC SID- [ISl] -ETRMACGLVKG-ALV SGRAIEGLATLTGAPSESIPLQASSLPMPSEDELDKDLIWAQLLSSRCV-RFLMGASCGGGNMKVDEEEYQQKGLRPR-• • AAYHAYTVLGYTVSN--GA--YYLI-IRNPWGVTEPTGDGVLSKRDW 445 HAY*V G * *N G LI *RNPWG *E *G* * * *W HAYSVTGVEEVNFHGRPEK-LIRLRNPWGEVEWSGAWSDNAPEWNYIDPRRKEELDKKA-EDGEFVMSFSDFL HAYSVTGAKQVNYRGQWS-LIRMRNPWGEVEWTGAWSDSSSEWNNVDPYERDQLRVKM-EDGEFWMSFRDFM HAYSVTAFKDVNYRGQQEQ-LIRIRNPWGQVEWTGAWSDGSSEWDNIDPSDREELQLKM-EDGEFWMSFRDFM HAYSVTGLEEALFKGEKVK-LVRIRNPWGQVEWNGSWSDGWKDWSFVDKDEKARLQHQVTEDGEFWMSYDDFV ---HAYSVLDVKDI--QGHR LLKLRNPWGHYSWRGDWSDDSSLWT--DDLRDALMPHGASE-GVFWISFEDVL • A o\ ISl Sequence Of Rat nCL-1: DGTNMTYGTSPSGLNMGELIARMVRNMDNSLLRDSDLDPRASDDRPSRTIVPVQY FIGURE m.12. Comparison of the amino-acid sequences of Tpr and domain II of the large subunits of several calpains. Amino acids marked with arrow (C, H and N) represent putative active site residues. Star mark (*) in "Consensus" indicates amino acids of Tpr which are homologous with the consensus sequence of the calpains. M E K K L V P Q S I S 5 ' TTATGGAAAAGAAATTAGTACCGCAATCCATTTCA 3 • AATACCTTTTCTTTAATCATGGCGTTAGGTAAAGT 5 ' GC GAATTCTAGTACCGCAATCCATTTCA 3 ' Primer ECO Y M Y * -TATATGTATTGA--ATATACATAACT-Primer BAM 3' CCGTTGTCCGAGATTGGTA CCTAGG££ 5' -GGCAACAGGCTCTAACCATGGATC 3 ' -CCGTTGTCCGAGATTGGTACCTAG 5 ' pYS307-2(+) I PCR I L V P Q S 5' GCGAATTCTAGTACCGCAATCC-3 ' CGCTTAAGATCATGGCGTTAGG-EcoRl Y M Y * -TATATGTATTGA--ATATACATAACT-Bam HI -AACCATGGATCCGC 3 ' -TTGGTACCTAGGCG 5 ' PCR product FIGURE m.13. Polymerase chain reaction product. Lanes: A-Hd, the molecular size marker, Mndin-digested ADNA; 2(+)-Hd/Hc, pYS307-2(+) digested with Mndlll and HincU; PCR, an end product of PCR. 68 as described in Materials and Methods (Fig. HI). The recombinant plasmid pT7-YS was used to transform E. coli JM83 containing pGPl-2. It was shown that E. coli JM83 containing pT7-YS/pGPl-2 could be induced to produce a proteolytic clear zone around colonies on skim-milk plate and the proteolytic band of 90 kDa in BSA-substrate zymography (data not shown). 4.2. High-level expression of tpr. The transformant, E. coli (pT7-YS/pGPl-2) was used for high-level expression of the recombinant tpr as described in Materials and Methods. SDS-PAGE analysis of reduced-denatured cell lysate samples showed the presence of a large amount of a 50-kDa protein (Fig. III. 14, A). The molecular mass of the highly expressed rTpr was the same as the product expressed in in-vitro translation study with pYS307 (Fig. III.6). The protein could be separated from the whole cell lysate by low speed centrifugation, suggesting that it was in the form of inclusion bodies. Inclusion bodies in E. coli were not visible by phase-contrast microscopy. Although the denatured 50-kDa protein could be seen on a Coomassie blue-stained SDS-PAGE gel only after induction by the T7 RNA polymerase/promoter system, the increase in the amount of the protein was not as significant as expected. In addition, the recombinant active 90-kDa protein was not detected by Coomassie-blue staining in nonreduced-nondenatured cell lysate samples (data not shown), although the protein was detected as an antibody reactive band in Western immunoblots (Fig. III. 14, B), and as a proteolytic band in BSA-substrate zymography (data not shown). These may have been due to autodegradation of the protein during incubation, since degradation products of the protein were detected on a Western immunoblot (Fig. III. 14, B). 4.3. NH 2 -terminal amino acid sequence of Tpr. The recombinant Tpr (rTpr) from high-level expression system was extracted from SDS-PAGE gel, and the NH2-terminal amino acid sequence determined. The NH2-terminus of the 69 R R NR UN IN UN IN UN IN ^ 2 9 _ kDa A B FIGURE HI.14. Coomassie blue-stained SDS-PAGE gel (A) and Western immunoblot (B) of Tpr from a high-level expression system. In the Western immunoblot analysis, antiserum raised against P. gingivalis W50 whole cell was used as a primary antibody. Lanes UN and IN are the whole-cell lysates of E. coli DH5cc (pGPl-2/pT7-YS) before and after induction, respectively. R represents reduced-denatured samples, and NR nonreduced-nondenatured samples, respectively. 70 protein was "ARILVPQSISKERLQKLEAQAT". With the exception of the first 3 amino acids, the sequence was identical to that deduced from the DNA sequence of the tpr gene. The first 3 amino acids (ARI) were identical to the amino acids encoded immediately upstream of the EcoRI site of pT7-7 (Fig. II. 1). They were inserted during generation of pT7-YS. 4.4. Western immunoblot analysis. Western blots were used to explore the relationship between the 50-kDa protein and the 90-kDa protease and to identify the protease in P. gingivalis which correlated with the recombinant protease. More specific antibodies were affinity purified against the denatured 50-kDa protein and the native 90-kDa recombinant protein. The anti-50-kDa antibody reacted with the 50-kDa protein as expected, but not with the 90-kDa recombinant protein or with protein in P. gingivalis lysates (data not shown). The antibody to the 90-kDa proteolytically active molecule reacted with an 80-kDa protein in nonreduced-nondenatured lysates of P. gingivalis W83, as well as the recombinant 90-kDa protein (Fig. III. 15). The discrepancy in reactivity suggested that there is a difference between the epitopes of the denatured protein and the native protease. The 80-kDa P. gingivalis protein which reacted with the anti 90-kDa antibody migrated to a position identical to an active protease as seen in gelatin-substrate zymography (see Fig. III. 17), suggesting that the conformation of Tpr is different in P. gingivalis. This could be due to differences in post-translational processing in the recombinant clone. 4.5. Antiserum raised against the inclusion bodies. Antiserum reactive with the P. gingivalis Tpr was generated by immunizing rabbits with the inclusion bodies containing rTpr. The antiserum reacted with an 80-kDa antigen in Western immunoblots of nonreduced-nondenatured samples of P. gingivalis W83 cell lysate (Fig. III. 16, lane NR). However, no reactive band was detected with reduced-denatured protein samples of P. gingivalis (Fig. III. 16, lane R). A 50-kDa antigen believed to be the 71 TZ YS W83 224 -109 -72 -46 -kDa FIGURE 111.15. Western immunoblot of P. gingivalis W83 probed with the antibody purified against the native 90-kDa recombinant protein. Lanes: TZ, lysate of E. coli JM83 (pTZ18R); YS, lysate of E. coli JM83 (pYS307-2(+)); W83, lysate of P. gingivalis W83. The samples are nonreduced-nondenatured. 72 TZ-R YS-R NR R FIGURE HL16. Western immunoblot of P. gingivalis W83 probed with anti-rTpr. Lanes: TZ-R, reduced-denatured sample of E. coli JM83 (pTZ18R); YS-R, reduced-denatured sample of E. coli JM83 (pYS307-2(+)); NR, nonreduced-nondenatured sample of P. gingivalis W83; R, reduced-denatured sample of P. gingivalis W83. 73 denatured form of rTpr was detected in the clone, but not in E. coli strain containing pTZ18R (Fig. IH.16, lanes TZ-R & YS-R). 4.6. Tpr in other P. gingivalis strains. The presence of Tpr in various P. gingivalis strains was determined by Western immunoblotting with anti-rTpr as the primary antibody. The strains known to be virulent in animal studies, e.g. P. gingivalis W12 and W50 (ATCC 53978), produced an 80-kDa protein which reacted with the anti-rTpr (Fig. in. 17, A). Avirulent P. gingivalis ATCC 33277 did not produce any antibody reactive band. This result indicated that P. gingivalis ATCC 33277 did not express Tpr, at least at levels detectable in Western immunoblot analysis. Gelatin-substrate zymography was performed with the same protein sample used in the Western immunoblot analysis to observe differences in the proteolytic-band patterns between P. gingivalis ATCC 33277 and the other P. gingivalis strains used above. All the strains used, except ATCC 33277, showed a major gelatinolytic band of molecular mass 80 kDa identical in mass to the antiserum reactive protein in Western immunoblot (Fig. III. 17, B), suggesting that Tpr is the major gelatinolytic enzyme detected under the zymography conditions. 5. Generation of a Tpr-Negative Mutant. 5.1. Conjugation. The suicide plasmid pBY2-IN prepared as described in Fig. II.2 was mobilized from E. coli (pBY2-IN/R751) to P. gingivalis W83 by conjugation. In the initial conjugation study, the cell mixture on the filter (see Materials and Methods) was incubated for 72 h under anaerobic conditions before plating onto the selection media, as suggested by Dr. N. B. Shoemaker (University of Illinois, Urbana). However, no transconjugant was obtained. Therefore, the method was modified by incubating the mixture aerobically at 37°C for 2 to 4 h followed by anaerobic incubation for 72 h. 110 erythromycin-resistant transconjugants were obtained after 2 weeks of anaerobic incubation. 74 FIGURE EH.17. Immunological and enzymatic assays for Tpr in a number of strains of P. gingivalis. Western immunoblot probed with anti-rTpr (A) and gelatin-substrate zymogram (B). Lanes: W83, P. gingivalis W83; 33277, P. gingivalis ATCC 33277; W12, P. gingivalis W12; W50, P. gingivalis W50 (ATCC 53798). The samples are nonreduced and nondenatured. 75 5.2. Isolation of a Tpr-deficient mutant. Of 110 transconjugant colonies, 77 continued to grow when subcultured onto erythromycin-containing BHI-blood agar plates. The chromosomal DNA isolated from each transconjugant was digested with Hindlll, and subjected to Southern hybridization. The 0.7-kb Pstl-Kpnl fragment of the tpr gene was used as a probe. An isogenic mutant, in which the tpr gene was inactivated by replacement of a portion of the gene with the Emr fragment from pBY2-IN was identified by screening for colonies whose chromosomal DNA failed to hybridize to the 0.7-kb Pstl-Kpnl fragment. The objective was to find a clone in which there had been a double crossover event as described in Fig. III. 18. A single strain lacking the 0.7-kb fragment was isolated (Fig. III. 19, lanes 2 and 4). This strain was named P. gingivalis W83/PM. To test stability, 100 colonies of the mutant were subcultured on blood agar plates without erythromycin. All the colonies remained resistant to erythromycin through 10 subcultures. 5.3. Southern hybridization. Disruption of the tpr gene was confirmed by Southern hybridization using a 3.2-kb HindSR fragment of pYS307 and 3.8-kb Emr fragment of pEM as a probe. As shown in Fig. III. 19, 1.4- and 2.9-kb fragments of the //fndlll-digested mutant chromosome hybridized with the 3.2-kb Hindlll probe (lane 6), suggesting that a portion of the tpr gene had been replaced with the Emr fragment (lane 8). The probe hybridized with a single 3.2-kb fragment of the parent chromosome (lane 5). 5.4. Western immunoblot and zymography. A specific protein/protease deficiency in the isogenic mutant, P. gingivalis W83/PM was confirmed by Western immunoblot and gelatin-substrate zymography. The 80-kDa antigen (Tpr) of the parent strain W83 was not detected in the isogenic mutant W83/PM in Western immunoblot probed with the anti-rTpr (Fig. in.20, A). When compared with the parent strain W83 (Fig. 111.20, B, lane 1), it could be seen that the mutant W83/PM (lane 2) had lost a 76 -J -J / " pBY2-IN I 6m He Ps Ec Hd Hd Hd Kp Hd "%.. Bn' ../-•' HdBm Bm He P8 Kp Hd tpr Chromosome of P. gingivalis W83 Double crossover Chromosome of the isogenic mutant, P. gingivalis W83/PM e crossover or Hd Hd Kp Hd FIGURE 111.18. Possible homologous recombinations between pBY2-IN and P. gingivalis chromosome. The thick black line and the hatched line represent the DNA fragment containing the tpr gene and the Emr  fragment, respectively. Abbreviations are as follows: Ac, Accl; Bm, BamUl; Ec, EcoW; He, Hindi; Hd, HindlE; Kp, Kpnl; Ps, Pstl; Sc, Sad; Sp, Sphl FIGURE m.19. Southern blot to identify the disrupted tpr gene. The chromosomal DNAs of P. gingivalis W83 (the odd-numbered lanes) and W83/PM (the even-numbered lanes) were digested with HindEl (lanes 1,2, 5 and 6) and Pstl-Kpnl (lanes 3,4,7 and 8). Three different DNA fragments were used as probes: the 0.7-kb Pstl-Kpnl fragment of pYS307 for lanes 1 to 4, the 3.2-kb Mrcdin fragment of pYS307 for lanes 5 and 6, and the 3.8-kb Emr fragment of pEM for lanes 7 and 8. Sizes are shown in kilobases. 78 i B FIGURE 111.20. Western immunoblot (A) and gelatin-substrate zymogram (B) of P. gingivalis W83/PM. Lanes: 1, lysate of P. gingivalis W83; 2, lysate of P. gingivalis W83/PM. All samples are nonreduced-nondenatured. In Western immunoblot analysis, anti-rTpr was used as a primary antibody. 79 major gelatinolytic protease band with an apparent molecular mass of 80 kDa, confirming the suggestion that Tpr was a major gelatinolytic protease of P. gingivalis W83. Disappearance of smaller proteolytic bands of W83 in the range of 50 to 70 kDa was also observed in W83/PM. 5.5. P. gingivalis W83/PM. The mutant P. gingivalis W83/PM still produced black pigment on blood-BHI agar plate, and appeared morphologically similar to W83. Growth curves of the parent strain and W83/PM are shown in Fig. 111.21; both strains were cultured in BHI broth. Comparison of growth rate showed W83/PM to have a generation time of 4.7 h whereas W83 divided every 3.5 h. The presence of erythromycin in the media did not affect the growth rate of the mutant. No differences between W83 and W83/PM were observed in the fatty acid profiles of the culture supernatants. In hemagglutination activity assay using human erythrocytes, W83/PM showed greater hemagglutinating activity with a titre of 16 for the parent strain and 64 for the mutant 5.6. Pz-peptidase activity of P. gingivalis W83 and W83/PM. The mutant strain created a smaller zone of protein hydrolysis (clearing) when grown on a skim-milk agar plate. However, no significant differences were observed in hydrolysis of the protease substrates, BApNA (Table III.l), SAApNA, and Azocoll (data not shown). In zymographic studies, it was shown that Tpr was most active against gelatin, indicating that it might be possible to use the chromogenic Pz-peptide as a substrate for quantifying the activity. As shown in Table III.l, the Pz-peptidase activity of the whole cell extract fraction (WCE) from the mutant was significantly reduced compared with that of the parent strain. This suggested that Tpr is the major P. gingivalis protease hydrolyzing the Pz-peptide, a bacterial collagenase substrate. The Pz-peptidase activity was associated with the cell membrane of P. gingivalis W83. 80 TIME (Hours) FIGURE m.21. Growth curves of P. gingivalis W83 and W83/PM. Organisms were grown in BHI broth. 81 TABLE III.1. Pz-peptidase activities3 and trypsin-like protease activities of P. gingivalis W83 and W83/PM. Whole cell extract Soluble cell extract Crude membrane Substrate Pz-peptide BApNA Pz-peptide BApNA Pz-peptide BApNA W83 1.6 1.3 0.5 0.7 1.8 1.8 W83/PM 0.3 1.5 0.3 0.8 0.4 2.1 a AA32o/h/mg of protein. The values are mean value of data obtained at 4 different times in Pz-peptidase assay. b A|j,mole of pNA/min/mg of protein. The values are mean value of data obtained at 2 times in the BApNA assay. 82 5.7. Identification of Tpr in P. gingivalis. Two dimensional (2D)-PAGE was used to identify the P. gingivalis Tpr protein. Cells for study were harvested from 0.5TYE broth. Proteins from the cellular fractions, whole cell extract (WCE), soluble cell extract (SCE) and crude membranes, of W83 and W83/PM were prepared and analyzed by 2D-PAGE as described in Materials and Methods. As shown in Fig. 111.22, a cellular component migrating to a position consistent with a molecular mass of 55 kDa was observed in WCE and crude membrane of W83. The same component was not seen in the mutant WCE and crude membrane, nor was it seen in SCE of W83 and W83/PM. The lack of the protein in W83/PM and the soluble extract of W83 was consistent with the enzymological data described in Table 111.1. The molecular mass of the protein was identical to that calculated from the amino-acid sequence of the recombinant Tpr (see section 3.3). An additional protein with an approximate molecular mass of 27 kDa (Fig. 111.22, marked by arrow) was seen in the WCE and crude membrane of W83. Western immunoblots were done to confirm that the native form of Tpr was present in the cellular fractions. Fig. 111.23 shows a Western immunoblot in which anti-rTpr was reacted with the nonreduced-nondenatured samples of the cellular fractions of W83 and W83/PM. All of the W83 fractions had antiserum-reactive bands of 80-100 kDa (lanes 1, 3, and 5). Compared with the other fractions, only a small amount of Tpr was present in the SCE fraction of W83. The polypeptide was missing in the cellular fractions of W83/PM. 6. Complementation. 6.1. Isolation of the complemented strain. The shuttle plasmid pBY3 was mobilized into P. gingivalis W83/PM using R751 as described in Materials and Methods. No transconjugants were detected using the same conjugation methodology which was successful in introducing pBY2-IN into P. gingivalis W83 (see section 5.1). Therefore, the protocol was modified as follows. The conjugation mixture on a BHI-blood agar plate was incubated aerobically overnight instead of for 2 h, and 83 W83 WCE oo W83/PM WCE r~TT FIGURE 111.22. Two-dimensional PAGE analysis of the cellular fractions of P. gingivalis W83 and W83/PM. Cells were cultivated in 0.5TYE broth. 12 \ig of proteins were used for each gel. WCE (A), SCE (B) and Env. (C) represent whole cell extracts, soluble cell extracts and crude membrane fractions, respectively. Proteins were detected by staining with silver nitrate. B W83 SCE P^ FIGURE IH.22. (continued) W83/PM SCE kDa —66-—45 — -36-24^ %± • <n "^.- •***^-r.s Hf. oo ON FIGURE IH.22. (continued) W83/PM Env. 4 J • FIGURE 111.23. Western immunoblot of the cellular fractions of P. gingivalis W83 and W83/PM probed with anti-rTpr. Cells were cultivated in 0.5TYE broth. 10 (ig of protein was applied to each lane. Lanes: 1, WCE of W83; 2, WCE of W83/PM; 3, SCE of W83; 4, SCE of W83/PM; 5, crude membrane fraction of W83; 6, crude membrane fraction of W83/PM. 87 i anaerobically for 24 h instead of 72 h. Prior to plating onto the selective medium, the mixture was suspended in fresh BHI broth and incubated anaerobically overnight, in order to enhance the viability of P. gingivalis. 5 transconjugants were isolated by this procedure. All 5 Tcr/Emr transconjugants were selected, then their plasmids isolated and analyzed by electrophoresis after digestion with Sstl. The restriction patterns of the plasmids were identical to that of pBY3 (data not shown). The plasmid isolated from a P. gingivalis transconjugant was used to transform E. coli. All of E. coli transformants produced the typical clear zone on skim-milk plates. These results showed that the transconjugants contained an intact autonomous plasmid, pBY3. One of the transconjugants was selected for further studies. 6.2. Stability of the plasmids in P. gingivalis W83/PM. The stability of pBY3 and pNJR12 in P. gingivalis W83/PM was measured by incubating the transconjugants on blood agar plates without erythromycin and tetracycline. After cultivation without either antibiotic, a high percentage (40 to 60%) of the transconjugants lost the ability to grow on the plates containing both antibiotics. Even after several transfers in BHI broth with Em and Tc, the growth of the transconjugants became slower, and eventually stopped. These results showed that both plasmids were unstable in P. gingivalis W83/PM. 6.3. Pz-peptidase activity of the complemented strain. Western immunoblots and gelatin-substrate zymography were performed to determine if pBY3 could restore the ability of W83/PM to express the protein Tpr. The results are shown in Fig. 111.24. The complemented strain produced a proteolytic band in zymography and an antiserum reactive band in Western immunoblot, which were identical with those produced by the parent strain W83. P. gingivalis W83/PM mobilized with the original shuttle plasmid pNJR12 (lacking the tpr gene) did not produce the 80-kDa proteolytic band or the antiserum reactive band on gelatin-substrate zymogram and Western immunoblot. 88 FIGURE HL24. Western immunoblot (A) and gelatin-substrate zymogram (B) of the complemented strain, P. gingivalis W83/PM (pBY3). Lanes: 1, cell extract of P. gingivalis W83; 2, cell extract of P. gingivalis W83/PM; 3. cell extract of P. gingivalis W83/PM (pBY3); 4, cell extract of P. gingivalis W83/PM (pNJR12). Cell extracts were prepared by sonication, as described in Materials and Methods. 10 \xg and 150 ng of proteins were used for Western immunoblot and zymography, respectively. All samples are nonreduced-nondenatured. In Western immunoblot analysis, anti-rTpr was the primary antibody. 89 The Pz-peptidase activity of whole cell extract of the complemented strain, P. gingivalis W83/PM (pBY3), was found to be approximately 2 times higher than the parent strain W83 (Table III.2). Mobilization of pNJR12 into W83/PM did not affect on the enzyme activity of the mutant. 7. Characterization of Tpr. Enzymatic properties of Tpr were determined, using the crude membranes obtained from W83 grown in 0.5TYE. These cells had a higher-level of Pz-peptidase activity than cells grown in complete growth media (discussed in section 8.2, Fig. 111.32). 7.1 Effects of reducing agents. Tpr has been characterized as a thiol-dependent protease (see section 2.1). The effect of reducing agents on the enzyme activity of Tpr was determined by measuring Pz-peptidase activity in the presence of cysteine, DTT and (3ME. Fig. 111.25 shows that reducing agents are essential for enzyme activity. DTT and (3ME stimulated the activity at concentration as low as 10 mM, while cysteine was required at concentrations higher than 20 mM to induce the activity. Concentrations of DTT and [3ME greater than 60 mM inhibited the reaction. 7.2 Effect of protease inhibitors and metal ions. The effect of inhibitors on W83 Pz-peptidase activity was determined by performing the enzyme assay in the presence of various inhibitors (Table III.3). The enzyme activity was strongly inhibited by the chymotrypsin-like protease inhibitor TPCK (0.5 mM) and the thiol -group blocking agent APMA (0.5 mM). High concentrations of TLCK (10 mM), EDTA (50 mM) or iodoacetate (5 mM) also inhibited the activity. Unlike the inhibition studies performed in BSA-substrate zymography with the recombinant clone (see section 3.2), the trypsin-like protease inhibitors, TLCK and PMSF, were only weakly inhibitory. The thiol-group blocking agents such as NEM and pCMB had little effect on enzyme activity. 90 TABLE III.2. Pz-peptidase activity of the complemented strain, P. gingivalis W83/PM (pBY3). Strains W83 W83/PM W83/PM (pBY3) W83/PM (pNJR12) Pz-peptidase activity (AA32o/h/mg of protein) 1.6 0.3 3.2 0.2 91 "3 o u a s o -< S >> • mm < Reducing Agent (mM) FIGURE ni.25. Effect of reducing agents on P. gingivalis W83 Pz-peptidase activity. 92 TABLE III.3. Effect of protease inhibitors on the Pz-peptidase activity of the crude membrane fraction of P. gingivalis W83. Inhibitors Concentration (mM) Relative Activity (%) None - 100 92 69 20 38 0 101 58 111 97 97 54 85 14 103 23 46 9 106 110 84 83 TLCK TPCK PMSF NEM pCMB EDTA Iodoacetate APMA Pepstatin Leupeptin * (i g/ml of concentration. 0.5 5 10 0.5 5 0.5 5 0.5 5 0.5 5 5 50 0.5 5 0.5 5 5* 50* 5* 50* 93 The effect of metal ions is shown in Table III.4. 20 mM Cd2+, Fe3+, Hg2+, and Zn2+ completely inhibited the enzyme activity. Zn2+ at levels as low as 5 mM was strongly inhibitory. Ca2+, Mg2+, and Mn2+ had litde effect on the enzyme activity. These are likely to be minimal inhibitory concentrations as it is possible that the metal ions complexed with the reducing agent. 7.3 Optimum pH. The effect of pH on the Pz-peptidase activity is shown in Fig. 111.26. The activity was highest between pH 7 and 9. The enzyme preparation showed higher activity in phosphate buffer at pH 7.0 than in Tris-HCl buffer at the same pH. 7.4 Effect of temperature. The optimum temperature of the Pz-peptidase activity is shown in Fig. 111.27. The activity increased with the increasing temperature up to 65°C, and then dropped dramatically at temperatures higher than 75°C. The enzyme showed approximately 2 times more enzyme activity at temperatures between 55 and 75°C than at 37°C. The membrane fraction of W83/PM always showed a very low level of activity regardless of temperature. The temperature stability of the Pz-peptidase activity was determined by pretreating the W83 membrane fraction at various temperatures for 30 min before the enzyme assay. Fig. 111.28 shows that the crude membranes of W83 retained 75% of their Pz-peptidase activity after incubation at 75°C, while only 30% remained after incubation at 85°C. Unlike the Pz-peptidase activity, trypsin-like enzyme(s) of W83 rapidly lost activity at temperatures above 55°C. 94 TABLE III.4. Effect of metal ions on the Pz-peptidase activity of the crude membrane fraction of P. gingivalis W83. Metal ions Concentration (mM) Relative activity (%) None 0 100 Ca2+ i 106 5 105 10 104 20 91 Mg2+ 1 103 5 103 10 108 20 100 Mn2+ 1 103 5 86 10 65 20 70 Rj3+ 1 64 5 51 10 48 20 0 Hg2+ 1 94 5 71 10 24 20 0 Cd2+ 1 91 5 61 10 49 20 0 Zn2+ 1 65 5 18 10 16 20 0 95 > — ' ••M 120-100-5 80-> "55 60-40-20-0- i 6 T" 7 T 8 pH - -a O Phosphate buffer •O Tris buffer • O— Glycine buffer T" 9 10 FIGURE 111.26. Effect of pH on the Pz-peptidase activity of P. gingivalis W83. 96 Temperature (°C) FIGURE III.27. Effect of temperature on the Pz-peptidase of P. gingivalis W83. 97 > o <! • ^ • p * en Pz-peptidase activity Trypsin-like activity 40 60 80 Temperature (°C) FIGURE 111.28. Effect of temperature on the stability of the Pz-peptidase activity of P. gingivalis W83. The crude membrane fraction was preincubated for 30 min at the given temperature and then the enzyme activity measured at 37°C. 98 8. Determination of the Role of Tpr. We attempted to determine the role of Tpr in collagen hydrolysis, in growth and nutrient uptake, and in in vivo pathogenicity. 8.1. Role of Tpr in collagen hydrolysis. 1) Collagen hydrolysis. The Tpr-deficient isogenic mutant W83/PM showed a greatly reduced ability to hydrolyze Pz-peptide, suggesting that Tpr might be involved in collagen degradation. To test this possibility, the coUagenolytic activities of W83 and W83/PM were compared using biologically labeled type I chick embryo [14C]-collagen. Fig. 111.29 shows the coUagenolytic activity of the crude membrane fractions of W83 and W83/PM. Both strains completely hydrolyzed collagen in about 6 h. In the time period between 1 and 4 h, W83/PM showed slightly higher collagen-hydrolyzing activity. SDS-PAGE of the reaction products followed by autoradiography showed that both strains were equally effective in hydrolyzing collagen (data not shown). 2) Effect of temperature and protease inhibitors. The results of the collagenase assay suggested either that Tpr was not involved in collagen degradation, or that coUagenolytic enzymes, other than Tpr, were expressed at higher levels in W83/PM. To further examine the involvement of Tpr in collagen degradation of P. gingivalis, the effects of temperature and protease inhibitors on the coUagenolytic activities of W83 and W83/PM were compared with their effects on Pz-peptidase activity (Fig. 111.28 and Table III.6). The results are shown in Fig. 111.30. In the Pz-peptidase assay, the enzyme retained 75% of the total activity when pretreated at 75°C for 30 min (Fig. 111.28). However, the coUagenolytic activities of both strains were reduced to 7% after pretreatment at 70°C for 30 min. Heating the enzyme to 50°C prior to adding it to the reaction mixture increased coUagenolytic activity. Unlike the Pz-peptidase activity, the coUagenolytic activities of both strains were strongly inhibited by TLCK (5 mM), but not by TPCK (5 mM). These results 99 c OX) — "o u T3 -s OX) T3 ^ 100. 3 W83 W83/PM Clostridial coUagenase Trypsin/TPCK 20 22 TIME (hours) FIGURE m.29. [14C]-labeled collagen-hydrolyzing activities of the crude membrane fractions of P. gingivalis W83 and W83/PM. Clostridial coUagenase and trypsin treated with TPCK were used as positive and negative controls, respectively. 100 $ < CO es 65 0) CJD « "3 U =3 T3 CO o 50°C 60°C 70°C 80°C TLCK TPCK APMA Treatment FIGURE 111.30. Effect of temperature and inhibitors on the collagenolytic activity of P. gingivalis W83 and W83/PM. The crude membrane fractions of both strains were preincubated for 30 min at the given temperature. Each protease inhibitor was added to the reaction mixture at a final concentration of 5 mM. 101 suggested that Tpr was not directly involved in collagen degradation by P. gingivalis. Further studies will be required to determine whether Tpr has a role in the hydrolysis of small peptides released by collagenolytic enzymes. 8.2. Role of Tpr in growth or nutrient uptake of P. gingivalis. 1) Growth of P. gingivalis W83 and W83/PM. Growth studies comparing W83 and W83/PM were done in TYE broth in which the trypticase peptone was reduced from 1.7% (normal TYE) to 0.5% (0.5TYE) (Fig. 111.31 and Table 8). In 0.5TYE, the maximum extent of growth was approximately 50% of that seen in normal TYE. Addition of BSA (final concentration, 0.2%) resulted in an increase in the final optical density, but gelatin (0.2%) had only a marginal effect. No significant differences between W83 and W83/PM were observed. These results suggest that proteases, other than Tpr, were induced when W83/PM was grown in peptide-limited or protein-supplemented media, and that these proteases could be responsible for enhancing the growth of P. gingivalis strains in protein-supplemented media. Generation times for W83 and W83/PM grown in TYE broth were similar, whereas the mutant had a longer generation time in BHI broth (see section 5.5 and Fig. 111.21). 2) Induction of Pz-peptidase activities under growth-limiting conditions. Fig. III. 32 shows Pz-peptidase activities (panel A) and trypsin-like enzyme activities (panel B) in different cellular fractions of W83 and W83/PM cultured for 40 h in TYE, 0.5TYE, 0.5TYE/Gelatin and 0.5TYE/BSA broth. All fractions of W83/PM produced very little Pz-peptidase activity. The Pz-peptidase activity of W83 was detected in crude membrane fractions in the cells from the four different growth media. When W83 was observed in 0.5TYE or 0.5TYE/Gelatin, Pz-peptidase activity of whole cell extracts was slightly higher, compared with that of the same strain grown in TYE or 0.5TYE/BSA. However, much higher activities were observed for the membrane-associated Pz-peptidase activities of W83 grown in 0.5TYE and 0.5TYE/Gelatin. It is noteworthy that, in contrast to BSA, gelatin showed little effect on the growth of W83 when added into 0.5TYE. The results 102 (A) P. gingivalis W83 FIGURE m.31. Growth of P. gingivalis W83 (A) and W83/PM (B) in TYE limited medium and protein supplemented medium. 0.5TYE/BSA and 0.5TYE/Gelatin represent 0.5TYE broth supplemented with 0.2% BSA or 0.2% gelatin, respectively. 103 TABLE III.5. Generation time1 and final optical density of P. gingivalis W83 and W83/PM grown in nutrient limited media and the same media supplemented with BSA or gelatin. Media TYE 0.5TYE 0.5TYE/BSA 0.5TYE/Gelatin Generation Time (hours) W83 3.5+0.3 4.1±0.6 4.6±0.8 4.2+0.4 W83/PM 4.1±0.5 4.1±0.7 4.4+0.5 4.2±0.4 Final Optical Density at 660nm W83 0.90 0.44 0.71 0.51 W83/PM 0.80 0.39 0.60 0.45 * The values are mean value of 9 different cultivations with 3 different inoculums. (A) Pz-peptidase activity o i-o. o M a I W83 W83/PM W83 W83/PM Whole cell extract Soluble cell extract W83 W83/PM Crude membrane (B) Trypsin-like activity 2.5-inTYE E3 in0.5TYE - r W83 W83/PM W83 W83/PM W83 W83/PM Whole cell extract Soluble cell extract Crude membrane FIGURE m.32. Pz-peptidase (A) and trypsin-like (B) activities of P. gingivalis W83 and W83/PM. A unit of trypsin-like activity was defined as the ability of the enzymes to produce 1 (imole of pNA per 1 min from B ApNA. 105 suggested that the enzyme activity of the membrane-associated Tpr was induced under culture conditions where growth of the organism was limited. In contrast, the general trypsin-like enzyme activities of both strains were not affected by the different growth media (Fig. in. 32, B). 8.3. Tpr and pathogenicity of P. gingivalis. The association of Tpr with pathogenicity was assessed by comparing the infectivity of W83 and W83/PM in the mouse model. When 1.5 X 1010 cells were injected subcutaneously in the abdominal area of mice (20 mice for each strain), both strains were capable of making the mice sick (less movement, change of appearance, trembling and occasional bleeding in injection area). 3 mice died (one injected with W83 and two with W83/PM), but the remainder recovered 3 days after injection. Unlike previous reports (73, 128, 231, 333, 335), localized abscess formation was not observed. 106 IV. DISCUSSION 1. Molecular Genetics of P. gingivalis. A protease gene (tpr) of P. gingivalis W83 was cloned and expressed in E. coli. Southern hybridization analysis of P. gingivalis chromosomal DNA using the 3.2 kb-//wdin fragment from pYS307 as a probe confirmed that the cloned gene originated from the genomic donor, and suggested that the gene is species-specific (Fig. III.3). The orientation of the insert did not influence expression of the recombinant protein (Fig. III.7), indicating that the cloned DNA fragment of the tpr gene contains a promoter region recognized by E. coli RNA polymerase. The potential promoter region containing the consensus sequence of E. coli promoter region was identified from the nucleotide sequence of the tpr gene (Fig. 1.3) (23). Klimpel and Clark (137) suggested that E. coli RNA polymerase may not be sufficiently similar to P. gingivalis for it to recognize P. gingivalis promoters. However, the promoter sequence associated with the tpr gene was functional, and generated a biologically active protein in E. coli. In recent years, several research groups have reported cloning and expression of P. gingivalis genes in E. coli; the list includes protease and collagenase genes (10, 65, 129,153, 168, 196, 216, 309), fimbrillin genes (53, 341, 358), superoxide dismutase (SOD) genes (38, 197), methylase gene (12), hemolysin genes (127), hemagglutinin gene (224), and genes encoding other surface antigens (1, 2, 97, 120,184, 342). Putative promoter regions (-10 and -35 regions) have been identified for the genes encoding a protease (23), SOD (38), and glutamate dehydrogenase (119) of P. gingivalis, and these showed homology to E. coli consensus promoter sequences. In the case of the sod gene, it was shown that transcription in E. coli was not initiated from the putative P. gingivalis promoter, but from the lacZ' promoter on the cloning vector. The DNA sequences of some P. gingivalis protease genes, such as prtT (216) and prtH (65), do not appear to have -10 and -35 promoter consensus sequences. Promoter sequences functioning in P. gingivalis have not yet been identified. Therefore, it is 107 not clear whether a true P. gingivalis promoter can be recognized by the transcriptional apparatus of E. coli. Expression of the recombinant P. gingivalis protein may be due to the fortuitous presence of a sequence homologous to that of the E. coli promoter. Isogenic mutants are excellent tools with which to identify virulence factors and to characterize the metabolic role of specific enzymes. The ability to generate these mutants from P. gingivalis has been hindered by the lack of well-defined genetic systems in the organism. Despite recent attempts, no indigenous plasmid has been demonstrated in P. gingivalis, nor has the acquisition of antibiotic resistance been reported (240, 354). Thus, no naturally occurring genetic elements have appeared in this organism that could be exploited for genetic analysis. Genetic systems developed for colonic Bacteroides species (for a review, see references 207, 241) have been applied to the molecular biological study of P. gingivalis for the following reasons: (i) P. gingivalis is phylogenetically more closely related to Bacteroides than to E. coli (219, 349), (ii) RNA polymerase(s) from P. gingivalis and E. coli appear to have different subunit structures and to exhibit different DNA template specificity (137), and (iii) no antibiotic resistance gene has been known to function in E. coli and also in Bacteroides species as well as P. gingivalis (207, 241). The genetic systems of Bacteroides species was successfully adapted for mutagenesis and complementation in the present study. We were interested in constructing a strain which was unable to express the protein Tpr. Construction of a Tpr-negative mutant of P. gingivalis W83 was accomplished by using suicide vector-mediated mutagenesis. The suicide plasmid pBY2-IN (Fig. II.2) contains the tpr gene disrupted by replacement of a portion of the gene with the Emr fragment. Since pBY2-IN can be transferred from E. coli to P. gingivalis but does not replicate in P. gingivalis, it serves for delivery of the disrupted gene into P. gingivalis W83 chromosome. A specific protease-deficient mutant P. gingivalis W83/PM was generated by homologous recombination between the disrupted gene of the suicide plasmid and the tpr gene in the P. gingivalis chromosome (Fig. III. 18). Disruption of the tpr gene was confirmed by Southern 108 hybridization using the cloned DNA fragment and Emr fragment as a probe (Fig. 111.19). Loss of Tpr, which has an apparent molecular mass of 80 kDa, was also confirmed by Western blot immunoassay and gelatin-substrate zymography (Fig. 111.20). Hoover et al. (105) first reported the isolation of randomly generated P. gingivalis mutants, including trypsin-like protease-deficient mutants, using a transposable element 7n4351, a transposon originally found on B. fragilis plasmid pBF4 (263). A plasmid R751::*Q4 used in the study is a conjugal broad-host-range IncP plasmid that contains a partial tandem duplication of Tn4351. They demonstrated that R751::*£24 can be transferred from E. coli to P. gingivalis by conjugation, and that Tn4351 can transpose from R751::*Q4 to the P. gingivalis chromosome. Recently, several research groups (93,119, 169) succeeded in generating isogenic mutants of P. gingivalis by insertional mutagenesis using suicide plasmids which is the same technique described in the present study. By characterizing the nonfimbriated mutant, it was suggested that the FimA protein of P. gingivalis plays an important role in the pathogenesis of periodontal disease (93,169). The present study provides evidence that the shuttle vector plasmids constructed for Bacteroides species can be used to express foreign genes in P. gingivalis strains, and demonstrated that the genetic system of Bacteroides species can be used to genetically manipulate P. gingivalis in examining putative virulence determinants that may participate in the induction or exaggeration of periodontal disease. To complement the Tpr-deficient mutant, the recombinant shuttle plasmid pBY3 was constructed by subcloning the tpr gene into the E. coli-Bacteroides shuttle vector pNJR12 (Fig. II.2). Introduction of pBY3 restored the ability of the isogenic mutant to express Tpr. The complemented strain showed 2 times higher Pz-peptidase activity than the parent strain, and produced a 80-kDa gelatinolytic band on a zymogram. Further studies are required to observe how increased level of the Tpr expression affects P. gingivalis. These studies would provide useful information to help determine the role of Tpr. 109 Plasmids originally developed from Bacteroides species have been successfully introduced from E. coli to P. gingivalis via conjugation (54, 105, 171, 223). There is a difference of opinion about the best conditions necessary to encourage conjugal transfer of plasmids into P. gingivalis. Maley et al. (171) suggested that long anaerobic incubation is important, whereas Hoover et al. (105) suggested that aerobic incubation for 4 to 24 h prior to anaerobic incubation improves the frequency of transfer. Dyer et al. (54) reported that the frequency of transfer of pVAL-1 into P. gingivalis was 10- to 100-fold higher when the mating mixture was incubated under aerobic conditions. We initially found that aerobic incubation for 2 to 4 h followed by anaerobic incubation for 48 to 72 h was the method of choice for the transfer of the suicide plasmid pB Y2-IN. Clearly, it seemed to be important to limit exposure to oxygen to a time which does not lead to death of P. gingivalis. However, in the complementation study, no transconjugant with the shuttle plasmid pBY3 could be isolated by this method. Modification of the method was required to increase the conjugal efficiency. Incubation time under aerobic conditions was increased from 2 h to overnight, in order to increase mobilizing ability of the donor E. coli. After anaerobic incubation for 24 h, the conjugation mixture on the filter was suspended in fresh liquid medium, and incubated anaerobically overnight, in order to increase viability of the recipient P. gingivalis. 5 transconjugants of the recombinant plasmid pBY3 were isolated by this procedure. The shuttle vector pNJR12 showed approximately 100 times higher conjugal efficiency than pBY3. The plasmids, pBY3 and pNJR12, were isolated from P. gingivalis transconjugants, showing that the foreign plasmid DNA can exist as autonomous replicating form in P. gingivalis. In the study by Dyer et al. (54), the autonomous plasmid pVAL-1 could not be detected reproducibly by agarose gel electrophoresis of lysates of P. gingivalis transconjugants. However, the presence of a replicating plasmid was demonstrated by Southern blot analysis, indicating that pVAL-1 existed as a low-copy autonomous plasmid in P. gingivalis. Maley et al. (171) also reported the successful isolation of plasmid from P. 110 gingivalis transconjugants, and showed that plasmid DNA isolated from one of the pNJR12 transconjugants of P. gingivalis W83 had an additional 1.5-kb inserted DNA. In the recent study, the additional DNA was characterized as an insertional sequence (IS 1126) from P. gingivalis W83 which is the only IS element so far isolated from P. gingivalis (170). The intact plasmid pBY3, in the present study, could be isolated from all the transconjugants, and was detected by agarose gel electrophoresis without any major modification. Instability of plasmids in P. gingivalis was reported by several research groups (54,171, 353). Maley et al. (171) demonstrated that, unlike pNJR12 transconjugants, pNJR5 transconjugants were difficult to maintain on selective media, indicating that one plasmid is less stable in P. gingivalis than the other.. Both plasmids are IncQ RSF1010 based E. coli-Bacteroides shuttle vector (262). The plasmid pNJR12 which appeared to be stable in P. gingivalis was chosen as a shuttle vector in the complementation study. However, in the present study, instability of the shuttle plasmids, pNJR12 and the recombinant plasmid pBY3, was observed. A high percentage (40 to 60%) of the transconjugants lost the ability to grow in the selection medium containing Em and Tc, after cultivation without either antibiotic. Even the transconjugants stopped growing in the selection medium after several subcultures in the medium. 2. Characterization of the tpr Gene and its Gene Product. The cloned protease gene was localized to the 1.9-kb HincH-HindUl fragment (Fig. HI.7). The potential promoter region and transcription direction were also determined (Fig. HI.7) by Western immunoblot analysis of the subclones (Fig. III.9). The location and potential transcription direction of the cloned gene were identical to those of tpr (Fig. 1.3) cloned by Bourgeau et al. (23). In addition, both studies showed similarities in the effect of protease inhibitors on the recombinant enzyme. Therefore, it was believed that both genes were identical. In the present study, the molecular mass of the cloned protease was determined to be approximately 90 kDa for the nondenatured active protease and 50 kDa for the denatured protein in Western blot immunoassay (Fig. III.5), suggesting that the cloned protease of 111 90 kDa contains a 50-kDa subunit. The in vitro translation study (Fig. 111.6) and a high-level expression study (Fig. III. 14) provided additional evidence showing that the gene encodes the 50-kDa protein. According to the study by Bourgeau et al. (23), the molecular mass of the tpr gene product was 64 kDa on BSA-substrate zymogram. However, the molecular mass of the same gene product had been previously determined to be 82 kDa using the same assay (157). This is a similar molecular mass to that of the proteolytic band (90 kDa) detected in the present study. We have been unable to detect any proteolytic band with a molecular mass 64 kDa on the zymogram, nor have we been able to detect protein or specific antigen in this region. Bourgeau et al. (23) also reported that the molecular mass of the cloned protease was 62.5 kDa when calculated from the translation product deduced from the nucleotide sequence of the tpr gene. However, using two different computer sequence analysis programs, we determined the molecular mass of the gene product to be 55 kDa which is comparable with that of the denatured protein (50 kDa) in SDS-PAGE analysis . A 55-kDa protein which did not exist in the isogenic mutant was identified in 2D-PAGE analysis (Fig. 111.22), and was believed to be the tpr gene product in P. gingivalis W83. The molecular mass of the protein was identical to that calculated from the translation product of the tpr gene, and was similar to that of the denatured recombinant Tpr (50 kDa). However, Western immunoblot analysis, using antibody purified against the active 90-kDa recombinant protein or anti-rTpr (Fig. in. 15 & in. 16), showed that the molecular mass of the native Tpr in P. gingivalis W83 (80 kDa) was smaller than that of the active recombinant protease (90 kDa). These findings showed that the active protease is composed of the tpr gene product with a molecular mass of approximately 55 kDa, and suggested that the conformation of the mature Tpr may be different in E. coli and P. gingivalis. It was also suggested that the active protease Tpr may exist as a dimer of the 55-kDa tpr gene product. 2D-PAGE analysis of P. gingivalis proteins (Fig. 111.22) showed an additional protein with a molecular mass of approximately 27 kDa in P. gingivalis W83 which was not present in W83/PM. This may have been a degradation product of the 55-kDa protein or another gene 112 product whose expression depends on the Tpr expression. It still remains to be determined whether the protein is the degradation product of Tpr or whether expression of the protein is regulated by Tpr. During screening of pro tease-positive clones, it was found that the positive clone could produce a proteolytic zone on skim-milk plate only when growth was deep within the agar or when incubated under anaerobic conditions, suggesting that the protease was only active under the reducing conditions. This was consistent with an observation by Bourgeau et al. (23). They showed that, while the clone produced a small clear zone after growth for 1 week on skim-milk plate in the presence of oxygen, a marked and clearly visible zone could be seen after 3 to 4 days under anaerobic conditions. These results were also similar to those of Arnott et al. (10). In their study, preliminary attempts to screen the library under aerobic conditions failed to detect any expression of cloned protease. However, repeating the screening procedure anaerobically identified three proteolytic recombinants. The recombinant Tpr with an approximate molecular mass of 90 kDa was only active in the presence of a reducing agent, cysteine, in BSA-substrate zymography (Fig. III.4). Pz-peptidase activity of the P. gingivalis Tpr was also stimulated by reducing agents such as DTT, (5ME and cysteine (Fig. 111.25). These results indicated that Tpr is a cysteine (thiol-dependent) protease. Additional evidence was provided by the amino-acid sequence analysis of Tpr. It was found that Tpr contains a consensus sequence that is highly conserved (60 to 65%) to thiol-dependent (cysteine) proteases from a wide range of species (23). The sequence is as follows; ^HAYTVLGYTVSNGAYYLIIRNPWG430 , where the underlined sequences indicate the identical amino acids in the active site of the cysteine proteases. It had been also suggested that three dimensional structure of the proteins tend to be more highly conserved, despite the fact that there was little overall identity between the related proteins from a wide range of species. 113 Additional related regions were identified in three-different regions on the Tpr protein by further analysis of the sequence (Fig. III.ll). These were highly related (58 to 87%) to the conserved regions of the eucaryotic calcium activated neutral proteases (CANP, calpains). These include one location that overlaps with the consensus sequence described above. Calpain (CANP; EC 3.4.22.17) is an intracellular nonlysosomal cysteine protease, the activity of which is regulated by calcium ions (for a review, see references 44, 293). The calpains exist ubiquitously in mammalian tissues, suggesting that they are responsible for essential and basic functions of cells. They consist of two distinct subunits, a large subunit (approximately 80 kDa) and a small subunit (approximately 30 kDa). The large subunit is responsible for protease activity and Ca2+ dependency, and can be structurally divided into 4 domains. The second domain (II) is a cysteine protease domain, and the fourth domain (IV) is a Ca2+-binding domain. Although the functions of the first and third domains are not clear, the domain structure clearly indicates that the calpain large subunit was generated by fusion of ancestral genes for cysteine protease and calmodulin (209). The small subunit is considered to be a regulatory subunit and contains a calcium binding domain similar to the domain IV of the large subunit and Gly-rich hydrophobic domain. The regions conserved in Tpr were located in domain II, the cysteine protease domain of the various calpains. The sequence of Tpr could be also aligned with those of domain II without significant deletions or insertions, showing 22% of identity and 53% of similarity (Fig. in. 12). Especially, the sequences of Tpr and of the calpains showed high-level similarity in the region where tentative active site residues (amino acids, C, H and N) of the proteases are located, suggesting that those residues of Tpr may also function as active site residues. Sequence analysis of Tpr reconfirmed the suggestion that the enzyme is a thiol-dependent protease of P. gingivalis. Nucleotide sequences of several protease genes from P. gingivalis have been reported; they include prtT coding for trypsin-like protease, (216), prtC for collagenase (129, 309), and prtH for a protease hydrolyzing human C3 complement protein (65). No significant 114 homology was seen in the deduced amino acid sequences between tpr and the other protease genes. Although the proteolytic activity of the recombinant Tpr was detected by zymography, the activity could not be quantified by enzyme assay using normal and synthetic protease substrates. This might have been due to low-level expression of the recombinant protease. Therefore, a T7 RNA polymerase/promoter system (295, 307) was adapted to express high levels of the recombinant protein. Most of the high-level expressed product formed inclusion bodies. The expression level of the recombinant protein after induction was not as high as expected. It might be due to the auto-degradation of the protein. We were unable to generate antiserum raised against P. gingivalis W83 whole cells reactive with the recombinant protein (rTpr). Therefore, antiserum previously raised against P. gingivalis W50 was shown to be reactive with the protein, and was used to detect rTpr in Western immunoblot analysis. Anti-enzyme antibodies protect the host by inactivating the enzyme and this might reduce the effectiveness of the proteases. However, some proteases have been reported to be insensitive to antibody. Frandsen et al. (66) reported that it was difficult to raise neutralizing antibody against a variety of Porphyromonas and Capnocytophaga proteases. Grenier et al. (89) found that the glycylprolyl protease of P. gingivalis (90) was insensitive to antibody raised against the purified active protease, and suggested that this insensitivity would make the enzyme a particularly effective pathogenic determinant. Antiserum specifically reactive with Tpr was required for the study. Therefore, the inclusion bodies containing rTpr were used as antigen to generate antiserum (anti-rTpr). A single protein of an apparent molecular mass of 80 kDa reacted with anti-rTpr (Fig. III. 16), when nonreduced-nondenatured sample of P. gingivalis W83 was used for Western immunoblot. Anti-rTpr did not react with any denatured P. gingivalis protein. Several different P. gingivalis strains were subjected to Western immunoblot analysis using anti-rTpr to examine the expression of Tpr in those strains (Fig. III. 17). Unlike the 115 other strains, P. gingivalis ATCC 33277 did not produce a reactive band, suggesting that ATCC 33277 does not express Tpr protein at levels detectable by Western immunoblot assay. Gelatin-substrate zymography also showed that ATCC 33277 does not produce a distinct proteolytic band of 80 kDa. In Southern hybridization study (Fig. III.3), this strain showed different hybridized pattern from the other P. gingivalis strains. The tpr gene and/or the regulatory region of the gene in ATCC 33277 may be modified to reduce or eliminate gene expression. Previously, Grenier et al. (84) reported that a total of eight distinct bands of proteolytic activities of P. gingivalis could be detected on BSA-substrate zymogram, and that identical proteolytic bands were found in all four strains of P. gingivalis studied including ATCC 33277 and W83. The 80-kDa protease band (P4) was included in the eight distinct bands. In their study, all the proteolytic activity of ATCC 33277 was lost after the samples were heated at 75°C for 30 min. Our observation that Tpr retained 75% of its activity after the same treatment (Fig. 111.28) supports the suggestion that ATCC 33277 does not express Tpr protein. Birkedal-Hansen et al. (21) also reported that ATCC 33277 produced a gelatinolytic band of approximately 90 kDa on the zymogram, which was believed to be responsible for collagenolytic activity of the organism. However, in the present study, we were unable to detect a 90-kDa gelatinolytic band produced by ATCC 33277, but able to detect a smear band covering the region of 75 to 150 kDa. This may be responsible for the band observed by Birkedal-Hansen et al. (21). In Pz-peptidase assay, ATCC 33277 showed low-level activity similar to that of the isogenic mutant W83/PM, indicating the lack of the Tpr expression (data not shown). This result is apparently contradictory to that of Ng and Fung (199), in which ATCC 33277 showed higher activity than W83. The difference may be due to the enzyme assay used. In their study, P. gingivalis whole cells were used, and the activity appeared not to be thiol-dependent. When whole cells were used in our assay, there were significant variations in the activities, making interpretation difficult. 3. Characterization of Tpr and Determination of its Role. 116 By comparing the parent strain W83 and the isogenic mutant W83/PM, it was possible to determine the enzymatic function and characteristics of Tpr. Although it was shown that W83/PM does not produce a proteolytic band of 80 kDa on zymogram, no significant difference between W83 and W83/PM was found in the hydrolyzing activities against several synthetic substrates (BApNA, SAApNA, and Azocoll). Zymography studies showed that the enzyme is much more active against gelatin than against BSA. Gelatin is known to be a denatured form of collagen. Therefore, a bacterial collagenase substrate, Pz-peptide, was chosen as a substrate for enzyme assay. A chromogenic Pz-peptide (Pz-Pro-Leu-Gly-Pro-D-Arg), which contains an amino-acid sequence based on the -Gly-Pro-Xaa- tripeptide repeating pattern of the helical region of collagen, was designed for the collagenase of the bacterium Clostridium histolyticum by Wunsch and Heidrich in 1963 (350). Although Pz-peptide was designed for bacterial collagenase, mammalian collagenase has been shown to be unable to hydrolyze the synthetic substrate (96). When the substrate is cleaved at the bond between Leu and Gly, the hydrophobic Pz-Pro-Leu product is formed, and this can be extracted into ethyl acetate to be quantified spectrophotometrically. The isogenic mutant W83/PM showed significantly reduced capability of hydrolyzing Pz-peptide (Table III.l), suggesting that Tpr is the protease of P. gingivalis W83 capable of hydrolyzing Pz-peptide. Some level of Pz-peptide hydrolyzing activity of W83/PM might be due to other P. gingivalis proteases capable of hydrolyzing collagen (17, 21,154, 285, 321) or due to non-specific reaction by the other proteases. A comparable level of the activity could be detected with purified trypsin (data not shown). However, the whole cell extract of the E. coli recombinant clone could not hydrolyze the substrate. This might be due to low-level expression, and/or due to the instability of the recombinant protein in the clone. Pz-peptidase activities of P. gingivalis strains have been reported by several research groups (118, 199). Ng and Fung (199) suggested that the Pz-peptidase activity of P. gingivalis is associated with the cell membrane, which is consistent with the result of the present study. It has been suggested that the Pz-peptidase activities of P. gingivalis are 117 related to the coUagenase activity, because Pz-peptide is known to be a bacterial coUagenase substrate. Hino et al. (98) showed high activity of Pz-peptidase in inflamed gingiva and suggested implication of the enzyme in this pathological condition, probably in collagen metabolism. Recently, Sojar et al. (290) reported purification of a membrane-associated protease from P. gingivalis ATCC 33277 capable of hydrolyzing Pz-peptide as well as salt-solubilized collagen. The native protein with a molecular mass of 120 kDa was dissociated into two polypeptides with molecular mass of 60 kDa and 50 kDa when heated at the temperature above 50°C. This enzyme is believed to be different from Tpr, since its activity was not enhanced by reducing agents. Enzyme assay of soluble cell extract and crude membrane fraction of P. gingivalis W83 showed that Pz-peptidase activity is mainly associated with the cell envelope of the organism (Table III. 1), suggesting that Tpr is a membrane-associated protease. This was further supported by 2D-PAGE analysis (Fig. flT.22) and Western immunoblot analysis of the cellular fractions (Fig. 111.23). Location of the protease on cell envelope may be related to its potential roles, possibly degradation of collagenous substrate or uptake of peptide nutrients. Although Tpr was shown to be associated with the cell envelope of P. gingivalis, no potential signal sequence could be identified in the derived amino-acid sequence of the tpr gene. Two potential membrane associated helical domains were identified. These include a major hydrophobic region of approximately 30 residues in the middle of the sequence. This hydrophobic domain may play a role in localizing the molecule to the cell envelope of P. gingivalis. We tried to determine the function or role of P. gingivalis Tpr in collagen degradation, nutrient uptake and pathogenesis. Primarily, Tpr had been recognized as a protease involved in collagen hydrolysis, because of its ability to hydrolyze gelatin and Pz-peptide. However, the crude membrane fractions of both strains completely hydrolyzed collagen in 6 h (Fig. m.29). No significant difference was found in [14C]-labeled collagen hydrolyzing capabilities between W83 and W83/PM. There is a possibility that P. gingivalis W83/PM is able to 118 induce the expression of coUagenolytic enzyme(s) other than Tpr, that are capable of hydrolyzing collagen at rates comparable to that of the parent strain P. gingivalis W83. The effects of protease inhibitors and temperature on coUagenolytic activity of the membrane fractions of W83 and W83/PM were compared with those to the Pz-peptidase activity of W83. Unlike the Pz-peptidase activity, the coUagenolytic activities of both strains were significantly reduced when pretreated at 70°C for 30 min, and were strongly inhibited by TLCK but not by TPCK (Fig. 111.30). These results suggest that Tpr is not a coUagenolytic enzyme of P. gingivalis. In the study by Birkedal-Hansen et al. (21), P. gingivalis ATCC 33277 incapable of expressing Tpr showed slightly higher coUagenolytic activity than W83. The prtC gene was characterized by Kato et al. (129) to encode a novel collagenase which degrades soluble and reconstituted fibrillar type I collagen, but not gelatin or Pz-peptide. These results suggest that proteases other than Tpr are responsible for the coUagenolytic activity of P. gingivalis. Pz-peptidase which does not hydrolyze collagen is widely distributed in mammalian tissues (for reviews, see references 14, 320). There have been a number of speculations about the function of the mammalian Pz-peptidases. It has been variously named mammalian Pz-peptidase (EC 3.4.99.31), soluble metalloendopeptidase or endopeptidase 24.15 (thimet oligopeptidase, EC 3.4.24.15), endo-oligopeptidase A (EC 3.4.22.19). Because of the capability of hydrolyzing a bacterial collagenase substrate Pz-peptide, Pz-peptidase has been thought to be linked to the degradation of collagen. Indeed, increases in the enzyme activity are found in tissues with high remodeling activity and in biological situations in which collagen degradation is accelerated, suggesting that Pz-peptidase plays a part in the late stages of the degradation of collagen (189). However, there is no direct evidence for an involvement of the enzyme in extracellular matrix degradation. It was recently proposed that the multicatalytic proteinase is responsible for the first step in the cytoplasmic protein degradation pathway, and that Pz-peptidase together with other endo- and exopeptidases is involved in the degradation of the smaller peptides. Thus, there is still a possibility that Tpr may be involved in late stage of the degradation of collagen or proteins containing a Pz-119 peptide like sequence, in order to produce small peptides and/or amino acids which can be used as energy source for the growth of P. gingivalis. The collagenase assays used in the present study did not provide any evidence supporting this suggestion. Further studies are required. P. gingivalis, as an asaccharolytic microorganism, requires amino acids or peptides as energy sources. The proteolytic enzymes of the organism are believed to be involved in degradation of proteins in order to provide peptide nutrients for bacterial growth (75, 249, 260, 340). The involvement of Tpr in nutrient uptake of P. gingivalis was examined by comparing growth patterns between W83 and W83/PM in peptide-nutrient limited medium (0.5TYE) and protein supplemented media. Both strains showed limited growth in 0.5TYE. The final optical density when cells were grown in 0.5TYE was only half of that in normal TYE. Growth of both strains was enhanced by supplementing proteins such as BSA and gelatin (0.5TYE/BSA and 0.5TYE/Gelatin), compared with growth in 0.5TYE (Fig. 111.31, Table III.5). However, the effect of gelatin on the growth was very little, compared with that of BSA. The result suggests that Tpr may not be responsible for nutrient uptake of P. gingivalis. Increased level of the Pz-peptidase activity associated with the crude membrane fraction of P. gingivalis W83 was detected when the organism was grown in 0.5TYE and 0.5TYE/Gelatin (Fig. 111.32). This suggested that the Tpr expression is regulated by the growth conditions of P. gingivalis where peptide nutrients preferred by P. gingivalis are limited. However, general trypsin-like enzyme activity of P. gingivalis W83 was not affected by the different growth media. This is similar to the observation by Robertson et al. (228). In their study, the collagenolytic activity of P. gingivalis appeared to be enhanced when grown in a peptide-depleted medium Marsh et al. (176) also reported that levels of proteolytic enzyme activities of P. gingivalis are regulated by hemin in growth media. Trypsin-like activity was 3.5-fold higher in the hemin-excess grown cells. However, collagenolytic activity was 3-fold higher in hemin-limited cultures. These findings suggested that 120 expression of P. gingivalis proteases is regulated, dependent on the environment. Further studies are required to determine the regulation of the Tpr expression in different environments and the effect of the enzyme in physiology of this organism. In-vivo studies of pathogenicity using a mouse model has shown heterogeneity in virulence between some P. gingivalis strains (73, 128, 231, 333, 335). P. gingivalis W83 is classified as a virulent strain, but P. gingivalis ATCC 33277 appears to be avirulent (74, 88). Western immunoblot and zymography analyses showed that ATCC 33277 does not express Tpr, at least at a detectable level, unlike virulent P. gingivalis strains used in the present study (Fig. III. 17). This result suggested that Tpr might be involved in in vivo pathogenesis of P. gingivalis. However, virulence studies using the mouse model showed no significant difference in pathogenesis between W83 and W83/PM. We experienced some difficulties in the virulence test. Unlike previous studies, major changes such as abscess formation could not be observed in the infected animals. Therefore, virulence by P. gingivalis strains could have not been quantified by the method used in the present study, even if there had been differences between W83 and W83/PM in pathogenicity. A different method or animal model may be required to determine the involvement of Tpr in the pathogenesis of P. gingivalis in future. In conclusion, this study has identified and characterized a protease gene (tpr) of the oral pathogen P. gingivalis and its product Tpr was shown to be a thiol-dependent Pz-peptidase whose activity is increased under growth conditions limiting in peptide/amino acid nutrients. Animal infectivity studies using the mouse model suggested that Tpr may not be involved in the pathogenicity of P. gingivalis. The role of Tpr in pathogenicity of P. gingivalis still remains to be determined. 121 V. REFERENCES 1. Abiko, Y., M. Hayakawa, H. Aoki, T. Kikuchi, H. Shimatake, and H. Takiguchi. 1990. Cloning of a Bacteroides gingivalis outer membrane protein gene in Escherichia coli. Arch. Oral Biol. 35:689-695. 2. Abiko, Y., M. Hayakawa, H. Aoki, and H. Takiguchi. 1988. Gene cloning and expression of a Bacteroides gingivalis-specific protein antigen in Escherichia coli. Adv. Dent. Res. 2:310-314. 3. Abiko, Y., M. Hayakawa, S. Murai, and H. Takiguchi. 1985. Glycylprolyl dipeptidylaminopeptidase from Bacteroides gingivalis. J. Dent. Res. 64:106-111. 4. Amano, A,, T. Ishimoto, H. Tamagawa, and S. Shizukuishi. 1992. Role of superoxide dismutase in resistance of Porphyromonas gingivalis to killing by polymorphonuclear leukocytes. Infect. Immun. 60:712-714. 5. Amano, A., S. Shizukuish, A. Tsunamitsu, K. Maekawa, and S. Tsunasawa. 1990. The primary structure of superoxide dismutase purified from anaerobically maintained Bacteroides gingivalis. FEBS Lett. 272:217-220. 6. Amano, A., S. Shizukuishi, H. Tamagawa, K. Iwakura, S. Tsunasawa, and A. Tsunemitsu. 1990. Characterization of superoxide dismutases purified from either anaerobically maintained or aerated Bacteroides gingivalis. J. Bacteriol. 172:1457-1463. 7. Amano, A., H. Tamagawa, M. Takagaki, Y. Murakami, S. Shizukuishi, and A. Tsunemitsu. 1988. Relationship between enzyme activities involved in oxygen metabolism and oxygen tolerance in black-pigmented Bacteroides. J. Dent. Res. 67:1196-1199. 8. Ando, K. 1980. Collagenase, dipeptidylpeptidase IV, and cathepsin D activities in gingival fluid and whole saliva from patients with periodontal disease. J. Jpn. Assoc. Periodont. 22:387-402. 9. Aoki, K., S. Imajoh, S. Ohno, Y. Emori, M. Koike, G. Kosaki, and K. Suzuki. 1986. Complete sequence of large subunits of low-Ca2+-requiring from of human Ca2+-activated neutral protease (uCANP) deduced from its cDNA sequence. FEBS Lett. 205:313-317. 10. Arnott, M. A., G. Rigg, H. Shah, D. Williams, A. Wallace, and I. S. Roberts. 1990. Cloning and expression of a Porphyromonas (Bacteroides) gingivalis protease gene in Escherichia coli. Arch. Oral Biol. 35:97S-99S. 11. Ausubel, F. A., R. Brent, R. E. Kingston, D. D. Moore, J. G. Seidman, J. A. Smith, and K. Struhl. 1993. Current Protocols in Molecular Biology. Greene Publishing and Wiley-Interscience, New York. 12. Banas, J. A., J. J. Ferretti, and A. Progulske-Fox. 1991. Identification and sequence analysis of a methylase gene in Porphyromonas gingivalis. Nucleic Acids Res. 19:4189-4192. 122 13. Banda, M. J., Z. Werb, and J. H. McKerrow. 1987. Elastin degradation. Methods Enzymol. 144:288-305. 14. Barret, A. J. 1990. A new look at PZ-peptidase. Biol. Chem. Hoppe-Seyler. 371S: 311-320. 15. Barua, P. K., M. E. Neiders, A. Topolnycky, J. J. Zambon, and H. Birkedal-Hansen. 1989. Purification of an 80,000-Mr glycylprolyl peptidase from Bacteroides gingivalis. Infect. Immun. 57:2522-2528. 16. Beaman, L., and B. L. Beaman. 1984. The role of oxygen and its derivatives in microbial pathogenesis and host defense. Ann. Rev. Microbiol. 38:27-48. 17. Bedi, G. S., and T. Williams. 1994. Purification and characterization of a collagen-degrading protease fromPorphyromonas gingivalis. J. Biol. Chem. 269:599-606. 18. Bejarano, P. A., J. P. M. Langeveld, B. G. Hudson, and M. E. Noelken. 1989. Degradation of basement membranes by Pseudomonas aeruginosa elastase. Infect. Immun. 57:3783-3787. 19. Bergey, D. H., F. C. Harrison, R. S. Breed, B. W. Hammer, and F. M. Huntoon. 1930. Bergey's Manual of Determinative Bacteriology, 3rd ed. The Williams and Wilkins Co., Baltimore. 20. Berglundl, T., J. Linde, I. Ericsson, and B. Liljenberg. 1992. Enhanced gingivitis in deciduous and permanent dentition. J. Clin. Periodontol. 19:134-142. 21. Birkedal-Hansen, H., R. E. Taylor, J. J. Zambon, P. K. Barwa, and M. E. Neiders. 1988. Characterization of collagenolytic activity from strains of Bacteroides gingivalis. J. Periodontal Res. 23:258-264. 22. Birkedal-Hansen, H., B. R. Wells, H. Y. Lin, P. W. Caufield, and R. E. Taylor. 1984. Activation of keratinocyte-mediated collagen (type I) breakdown by suspected human periodontopathogen. Evidence of a novel mechanism of connective tissue breakdown. J. Periodontal Res. 19:645-650. 23. Bourgeau, G., H. Lapointe, P. Peloquin, and D. Mayrand. 1992. Cloning, expression, and sequencing of a protease gene (tpr) from Porphyromonas gingivalis W83 in Escherichia coli. Infect Immun. 60:3186-3192. 24. Bourgeau, G., and D. Mayrand. 1990. Aggregation of Actinomyces strains by extracellular vesicles produced by Bacteroides gingivalis. Can. J. Microbiol. 36:362-365. 25. Boyd, J., and B. C. McBride. 1984. Fractionation of hemagglutinating and bacterial binding adhesins of Bacteroides gingivalis. Infect. Immun. 45:403-409. 26. Bramanti, T. E., G. G. Wong, S. T. Weintraub, and S. C. Holt. 1989. Chemical characterization and biologic properties of lipopolysaccharide from Bacteroides gingivalis W50, W83 and ATCC 33277. Oral Microbiol. Immunol. 4:183-192. 27. Bricker, J., M. H. Mulks, A. G. Plaut, E. R. Moxon, and A. Wright. 1983. IgAl proteases of Haemophilus influenzae : Cloning and characterization in Escherichia coli K-12. Proc. Natl. Acad. Sci. USA. 80:2681-2685. 123 28. Brocklehurst, K. 1982. Two-protonic state electophiles as probes of enzyme mechanism. Methods Enzymol. 87:427-469. 29. Brocklehurst, K., and G. Little. 1973. Reactions of papain and of low molecular weight thiols with some aromatic disulphides 2,2'-dipyridyl disulphide as a convenient active site titrant for papain even in the presence of other thiols. Biochem. J. 133:67-80. 30. Brook, I. 1986. Encapsulated anaerobic bacteria in synergistic infections. Microbiol. Rev. 50:452-457. 31. Carlsson, J., B. F. Herrmann, J. F. Hofling, and G. K. Sundqvist. 1984. Degradation of the human proteinase inhibitors alpha-1-antitrypsin and alpha-2-macroglobulin by Bacteroides gingivalis. Infect. Immun. 43:644-648. 32. Carlsson, J., J. F. Hofling, and G. K. Sundqvist. 1984. Degradation of albumin, haemopexin, haptoglobin and transferrin, by black-pigmented Bacteroides species. J. Med. Microbiol. 18:39-46. 33. Chen, Z., J. Potempa, A. Polanowski, S. Renvert, M. Wikstrom, and J. Travis. 1991. Stimulation of proteinase and amidase activities in Porphyromonas {Bacteroides) gingivalis by amino acids and dipeptides. Infect. Immun. 59:2846-2850. 34. Chen, Z., J. Potempa, A. Polanowski, M. Wikstrom, and J. Travis. 1992. Purification and characterization of a 50-kDa cysteine proteinase (gingipain) from Porphyromonas gingivalis. J. Biol. Chem. 267:18896-18901. 35. Childs, W. C , and R. J. Genco. 1988. Use of percoll density gradients for studing the attachment of bacteria to oral epithelial cells. J. Dent. Res. 67:826-830. 36. Childs, W. C , III, and R. J. Gibbons. 1990. Selective modulation of bacterial attachment to oral epithelial cells by enzyme activities associated with poor oral hygiene. J. Periodontal Res. 25:172-178. 37. Choi, J.-L, T. Nakagawa, S. Yamada, I. Takazoe, and K. Okuda. 1990. Clinical microbiolgical and immunological studies or recurrent periodontal disease. J. Clin. Periodontol. 17:426-434. 38. Choi, J.-L, N. Takahashi, T. Kato, and H. K. Kuramitsu. 1991. Isolation, expression, and nucleotide sequence of the sod gene from Porphyromonas gingivalis. Infect. Immun. 59:1564-1566. 39. Christersson, L. A., J. J. Zambon, R. G. Dunford, S. G. Grossi, and R. J. Genco. 1989. Specific subgingival bacteria and diagnosis of gingivitis and periodontitis. J. Dent. Res. 68:1633-1639. 40. Chu, L., T. E. Bramanti, J. L. Ebersole, and S. C. Holt. 1991. Hemolytic activity in the periodontopathogen Porphyromonas gingivalis: Kinetics of enzyme release and localization. Infect. Immun. 59:1932-1940. 41. Cimasoni, G., M. Song, and B. C. McBride. 1987. Effect of crevicular fluid and lysosomal enzymes on the adherence of streptococci and bacteroides to hydroxyapatite. Infect. Immun. 55:1484-1489. 124 42. Courant, P. R., and R. J. Gibbons. 1967. Biochemical and immunological heterogeneity of Bacteroides melaninogenicus. Arch. Oral Biol. 12:1605-1613. 43. Coykendall, A. L., F. S. Kaczmarek, and J. Slots. 1980. Genetic heterogeneity in Bacteroides asaccharolyticus (Holdeman and Moore 1970) Finegold and Barnes 1977 (Approved Lists, 1980) and proposal of Bacteroides gingivalis sp. nov. and Bacteroides macacae (Slots and Genco) comb. nov. Int. J. Syst. Bacteriol. 30:559-564. 44. Croall, D. E., and G. N. Dermartino. 1991. Calcium-activated neutral protease (calpain) system: Structure, function, and regulation. Physiol. Rev. 71:813-847. 45. Curtis, M. A., M. Ramakrishnan, and J. M. Slaney. 1993. Characterization of the trypsin-like enzymes of Porphyromonas gingivalis W83 using a radiolabeled active-site-directed inhibitor. J. Gen. Microbiol. 139:949-955. 46. Cutler, C. W., R. R. Arnold, and H. A. Schenkein. 1993. Inhibition of C3 and IgG proteolysis enhances phagocytosis of Porphyromonas gingivalis. J. Immunol. 151:7016-7029. 47. Davison, J., M. Heusterspreute, N. Chevalier, V. Ha-Thi, and F. Brunei. 1987. Vectors with restriction site banks; V. pJRD215, a wide-host-range cosmid vector with multiple cloning sites. Gene. 51:275-280. 48. de Nardin, A. M., H. T. Sojar, S. G. Grossi, L. A. Christersson, and R. J. Genco. 1991. Humarol immunity of older adults with periodontal disease to Porphyromonas gingivalis. Infect. Immun. 59:4363-4370. 49. Delaney, S. J., D. C. Hayward, F. Barleben, K. F. Fischbach, and G. L. Gabor Miklos. 1991. Molecular cloning and analysis of small optic lobes, a structural brain gene of Drosophila melanogaster. Proc. Natl. Acad. Sci. USA. 88:7214-7218. 50. DeLuca, C. I., P. L. Davies, J. A. Samis, and J. S. Elce. 1993. Molecular cloning and bacterial expression of cDNA for rat calpain II 80kDa subunit. BBA. 1216:81-93. 51. DesLauriers, M., and C. Mouton. 1992. Epitope mapping of hemagglutination adhesin HA-Ag2 of Bacteroides {Porphyromonas) gingivalis. Infect. Immun. 60:2791-2799. 52. Di Murro, C , R. Nishini, M. Cattabriga, A. Simonetti-D'Arca, S. LeMole, M. Paolantanio, L. Sebastiani, and R. D'Amelio. 1987. Rapidly progressive periodontitis in neutrophil chemotaxis inhibitory factors associated with the presence of Bacteroides gingivalis in crevicular fluid. J. Periodontal. 58:868-872. 53. Dickinson, D. P., M. A. Kubiniec, F. Yoshimura, and R. J. Genco. 1988. Molecular cloning and sequencing of the gene encoding the fimbrial subunit protein of Bacteroides gingivalis. J. Bacteriol. 170:1658-1665. 54. Dyer, D. W., G. Bilalis, J. H. Michel, and R. Malek. 1992. Conjugal transfer of plasmid and transposon DNA from Escherichia coli into Porphyromonas gingivalis. Biochem. Biophys. Res. Commun. 186:1012-1019. 55. Dzink, J. L., S. S. Socransky, and A. D. Haffajee. 1988. The predominant cultivable microbiota of active and inactive lesions of destructive periodontal diseases. J. Clin. Periodontol. 15:316-323. 125 56. Ebersole, J. L., D. E. Frey, M. A. Taubman, A. D. Haffajee, and S. S. Socransky. 1987. Dynamics of systemic antibody responses in periodontal disease. J. Periodontal Res. 22:184-186. 57. Ebersole, J. L., M. A. Taubman, and D. J. Smith. 1985. Gingival crevicular fluid antibody to oral microorganisms. II. Distribution and specificity to local antibody responses. J. Periodontal Res. 20:3149-356. 58. Ebersole, J. L., M. A. Taubman, D. J. Smith, and D. E. Frey. 1986. Human immune responses to oral microorganisms; patterns of systemic antibody levels to Bacteroides species. Infect. Immun. 51:507-513. 59. Ebersole, J. L., M. A. Taubman, D. J. Smith, and J. M. Goodson. 1984. Gingival crevicular fluid antibody to oral microorganisms. I. Method of collection and analysis of antibody. J. Periodontal Res. 19:124-132. 60. Ellen, R. P., and D. A. Grove. 1989. Bacteroides gingivalis vesicles bind to and aggregate Actinomyces viscosus. Infect. Immun. 57:1618-1620. 61. Endo, J., M. Otsuka, E. Ohara, M. Sato, and R. Nakamura. 1989. Cleavage action of a trypsin-like protease from Bacteroides gingivalis 381 on reduced egg-white lysozyme. Arch. Oral Biol. 34:911-916. 62. Finegold, S. M., and E. M. Barnes. 1977. Report of the ICSB Taxonomic Subcommittee on Gram-Negative Anaerobic Rods. Proposal that the saccharolytic and asaccharolytic strains at present classified in the species Bacteroides melaninogenicus (Oliver and Wherry) be classified in two species as Bacteroides melaninogenicus and Bacteroides asaccharolyticus. Int. J. Syst. Bacterid. 27:388-391. 63. Finkelstein, R. A., M. Boesman-Finkelstein, Y. Chang, and C. C. Hase. 1992. Vibrio cholerae hemagglutinin/protease, colonial variation, virulence, and detachment. Infect. Immun. 60:472-478. 64. Fishburn, C. S., J. M. Slaney, R. J. Carman, and M. A. Curtis. 1991. Degradation of plasma proteins by the trypsin-like enzyme of Porphyromonas gingivalis and inhibition of protease activity by a serine protease inhibitor of human plasma. Oral Microbiol. Immunol. 6:209-215. 65. Fletcher, H. M., H. A. Schenkein, and F. L. Macrina. 1994. Cloning and characterization of a new protease gene (prtH) from Porphyromonas gingivalis. Infect. Immun. 62:4279-4286. 66. Frandsen, E. V. G., J. Reinholdt, and M. Kilian. 1987. Enzymatic and antigenic characterization of immunoglobulin Al proteases from Bacteroides and Capnocytophaga spp. Infect. Immun. 55:631-638. 67. From, S. H., and S. D. Schultz-Hault. 1963. Comparative histological and microchemical evaluations of the collagen of human gingiva. J. Periodontal. 34:216-222. 68. Fujimura, S., and T. Nakamura. 1987. Isolation and characterization of a protease from Bacteroides gingivalis. Infect. Immun. 55:716-720. 126 69. Fujimura, S., and T. Nakamura. 1989. Multiple forms of proteases of Bacteroides gingivalis and their cellular location. Oral Microbiol. Immunol. 4:227-229. 70. Fujimura, S., Y. Shibata, and T. Nakamura. 1992. Comparative studies of three proteases of Porphyromonas gingivalis. Oral Microbiol. Immunol. 7:212-217. 71. Fujimura, S., Y. Shibata, and T. Nakamura. 1993. Purification and partial characterization of a lysine-specific protease of Porphyromonas gingivalis. FEMS Microbiol. Lett. 113:133-138. 72. Fullmer, H. M., W. A. Gibson, G. S. Lazrus, H. A. Bladen, and K. A. Whedon. 1969. The origin of collagenase in periodontal tissues. J. Dent. Res. 48:646-651. 73. Genco, C. A., C. W. Cutler, D. Kapczynski, K. Maloney, and R. R. Arnold. 1991. A novel mouse model to study the virulence of and host response to Porphyromonas (Bacteroides) gingivalis. Infect. Immun. 59:1255-1263. 74. Genco, C. A., D. R. Kapczynski, C. W. Cutler, R. J. Arko, and R. R. Arnold. 1992. Influence of immunization on Porphyromonas gingivalis colonization and invasion in the mouse chamber model. Infect. Immun. 60:1447-1454. 75. Gharbia, S. E., and H. N. Shah. 1991. Utilization of aspartate, glutamate, and their corresponding peptides by Fusobacterium nucleatum subspecies and Porphyromonas gingivalis. Curr. Microbiol. 22:159-163. 76. Gibbons, R. J. 1989. Bacterial adhesion to oral tissue: a model for infectious disease. J. Dent. Res. 68:750-760. 77. Gibbons, R. J., and J. B. MacDonald. 1960. Degradation of collagenous substrates by Bacteroides melaninogenicus. J. Bacterid. 81:614-621. 78. Gibbons, R. J., and J. B. MacDonald. 1960. Hemin and vitamin K compounds as required factors for the cultivation of certain strains of Bacteroides melaninogenicus. J. Bacteriol. 80:164-170. 79. Grenier, D. 1992. Demonstration of a bimodal coaggregation reaction between Porphyromonas gingivalis and Treponema denticola. Oral Microbiol. Immunol. 7:280-284. 80. Grenier, D. 1992. Further evidence for a possible role of trypsin-like activity in the adherence of Porphyromonas gingivalis. Can. J. Microbiol. 38:1189-1192. 81. Grenier, D. 1992. Inactivation of human serum bactericidal activity by a trypsinlike protease isolated from Porphyromonas gingivalis. Infect. Immun. 60:1854-1857. 82. Grenier, D. 1992. Nutritional interactions between two suspected periodontopathogens, Treponema denticola and Porphyromonas gingivalis. Infect. Immun. 60:5298-5301. 83. Grenier, D., and M. Belanger. 1991. Protective effect of Porphyromonas gingivalis outer membrane vesicles against bactericidal activity of human serum. Infect. Immun. 59:3004-3008. 127 84. Grenier, D., G. Chao, and B. C. McBride. 1989. Characterization of sodium dodecyl sulfate-stable Bacteroides gingivalis proteases by polyacrylamide gel electrophoresis. Infect Immun. 57:95-99. 85. Grenier, D., and D. Mayrand. 1983. Etudes d'infectins mixtes anaerobies comportant Bacteroides gingivalis. Can. J. Microbiol. 29:612-618. 86. Grenier, D., and D. Mayrand. 1985. Cytotoxic effects of culture supernatants of oral bacteria and various organic acids on vero cells. Can. J. Microbiol. 31:302-304. 87. Grenier, D., and D. Mayrand. 1987. Functional characterization of extracellular vesicles produced by Bacteroides gingivalis. Infect. Immun. 55:111-117. 88. Grenier, D., and D. Mayrand. 1987. Selected characteristics of pathogenic and nonpathogenic strains of Bacteroides gingivalis. J. Clin. Microbiol. 25:738-740. 89. Grenier, D., D. Mayrand, and B. C. McBride. 1989. Further studies on the degradation of immunoglobulins by black-pigmented Bacteroides. Oral Microbiol. Immunol. 4:12-18. 90. Grenier, D., and B. C. McBride. 1987. Isolation of a membrane-associated Bacteroides gingivalis glycylprolyl protease. Infect. Immun. 55:3131-3136. 91. Grenier, D., and B. C. McBride. 1989. Surface location of a Bacteroides gingivalis glycylprolyl protease. Infect. Immun. 57:3265-3269. 92. Grenier, D., and B. C. McBride. 1991. Preliminary studies on the influence of in vivo growth on selected characteristics of Porphyromonas gingivalis W83. Microbial Ecol. Health Dis. 4:105-111. 93. Hamada, N., K. Watanabe, C. Sasakawa, M. Yoshikawa, F. Yoshimura, and T. Umemoto. 1994. Construction and characterization of a fimA mutant of Porphyromonas gingivalis. Infect. Immun. 62:1696-1704. 94. Hanazawa, S., Y. Murakami, A. Takeshita, H. Kitami, K. Ohta, S. Amano, and S. Kitano. 1992. Porphyromonas gingivalis fimbriae induce expression of the neutrophil chemotactic factor KC gene of mouse peritoneal macrophages: Role of protein kinase C. Infect. Immun. 60:1544-1549. 95. Hanazawa, S., K. Nakada, Y. Ohmori, T. Miyoshi, S. Amano, and S. Kitano. 1985. Functional role of interleukin-1 in periodontal disease: induction of interleukin-1 production by Bacteroides gingivalis lipopolysaccharide in peritoneal macrophages from C3H/HeN and C3H/HeJ mice. Infect. Immun. 50:262-270. 96. Harper, E., and J. Gross. 1970. Separation of collagenase and peptidase activities of todpole tissues in culture. BBA. 198:286-292. 97. Hayakawa, M., Y. Abiko, T. Ito, H. Sasahara, H. Yamano, and H. Takiguchi. 1992. Gene cloning of Porphyromonas gingivalis specific antigens recognized by serum of adult periodontitis patient. Int. J. Biochem. 24:945-950. 98. Hino, M., G. Nakano, M. Harada, and T. Nagatsu. 1975. Distribution of PZ-peptidase and glycylprolyl p-naphthylamidase activities in oral tissues. Arch. Oral Biol. 20:19-22. 128 99. Hinode, D., H. Hayashi, and R. Nakamura. 1991. Purification and characterization,of three types of proteases from culture supernatants of Porphyromonas gingivalis. Infect. Immun. 59:3060-3068. 100. Hinode, D., A. Nagata, S. Ichimiya, H. Hayashi, M. Morioka, and R. Nakamura. 1992. Generation of plasma kinin by three types of protease isolated from Porphyromonas gingivalis 381. Arch. Oral Biol. 37:859-861. 101. Hofstad, T. 1970. Biological activities of endotoxins from Bacteroides melaninogenicus. Arch. Oral Biol. 15:343-348. 102. Hofstad, T. 1974. The distribution of heptose and 2-keto-3-deoxy-octane in Bacteroiaceae. J. Gen. Microbiol. 85:314-320. 103. Holdeman, L. V., and W. E. C. Moore. 1970. Outline of Clinical Methods in Anaerobic Bacteriology, 2nd ed. Virginia Plytechnic Institute and State University Anaerobe Laboratory, Blacksburg. 104. Holt, S. C , J. Ebersole, J. Felton, M. Brunsvold, and K. S. Kornman. 1988. Implantation of Bacteroides gingivalis in nonhuman primates initiates progression of periodontitis. Science. 239:55-57. 105. Hoover, C. I., E. Abarbarchuk, C. Y. Ng, and J. R. Felton. 1992. Transposition of Tn4351 in Porphyromonas gingivalis. Plasmid. 27:246-250. 106. Hoover, C. I., C. Y. Ng, and J. R. Felton. 1992. Correlation of haemagglutination activity with trypsin-like protease activity of Porphyromonas gingivalis. Arch. Oral Biol. 37:515-520. 107. Hopps, R. M., and H. J. Sismey-Durrant. 1991. Mechanisms of alveolar bone loss in periodontal disease, p. 307-. In S. Hamada, S. C. Holt, and J. R. McGhee (ed.), Periodontal Disease: Pathogens and Host Immune Responses. Quintessence, Tokyo. 108. Hotez, P., J. Haggerty, J. Hawdon, L. Milstone, H. R. Gamble, G. Schad, and F. Richards. 1990. Metalloproteases of infective Ancylostoma hookworm larvae and their possible functions in tissue invasion and ecdysis. Infect. Immun. 58:3883-3892. 109. Hunt, D. E., J. V. Jones, and V. R. Dowell Jr. 1986. Selective medium for the isolation of Bacteroides gingivalis. J. Clin. Microbiol. 23:441-445. 110. lino, Y., and R. M. Hopps. 1984. The bone resorbing activities in tissue culture of lipopolysaccharides from the bacteria Actinobacillus actinomycetemcomitans, Bacteroides gingivalis and Capnocytophaga ochracea isolated from human mouths. Arch. Oral Biol. 29:59-63. 111. Imajoh, S., K. Aoki, S. Ohmo, Y. Emori, H. Kawasaki, H. Sugihar, and K. Suzuki. 1988. Molecular cloning of the cDNA for the large subunit of the high-Ca2+-requiring form of human Ca2+-activated neutral protease. Biochem. 27:8122-8128. 112. Imamura, T., R. N. Pike, J. Potempa, and J. Travis. 1994. Pathogenesis of periodontitis: a major arginine-specific cysteine proteinase from Porphyromonas gingivalis induces vascular permeability enhancement through activation of the kallikrein/kinin pathway. J. Clin. Invest. 94:361-367. 129 113. Inoshita, E., A. Amano, T. Hanioka, H. Tamagawa, S. Shizukuishi, and A. Tsunemitsu. 1986. Isolation and some properties of exogemagglutinin from the culture medium of Bacteroides gingivalis. Infect. Immun. 52:421-427. 114. Inoshita, E., K. Iwakura, A. Amano, H. Tamagawa, and S. Shizukuishi. 1991. Effect of transferrin on the growth of Porphyromonas gingivalis. J. Dent. Res. 70:1258-1261. 115. Ismaiel, M. O., J. Greenman, K. Morgan, M. G. Glover, A. S. Rees, and C. Scully. 1989. Periodontitis in sheep: a model for human periodontal disease. J. Periodontal. 60:279-284. 116. Isogai, E., K. Hirose, N. Fujii, and H. Isogai. 1992. Three types of binding by Porphyromonas gingivalis and oral bacteria to fibronectin, buccal epithelial cells, and erythrocytes. Arch. Oral Biol. 8:667-670. 117. Isogai, H., E. Isogai, F. Yohimura, T. Suzuki, W. Kagota, and K. Takano. 1988. Specific inhibition of an oral strain of Bacteroides gingivalis 381 to epithelial cells by monoclonal antibodies against the bacterial fimbriae. Arch. Oral Biol. 33:479-485. 118. Jin, K.-C, P. K. Barua, J. J. Zambon, and M. E. Neiders. 1989. Proteolytic activity in black-pigmented Bacteroides species. J. Endodont. 15:463-467. 119. Joe, A., C. S. Murray, and B. C. McBride. 1994. Nucleotide sequence of a Porphyromonas gingivalis gene encoding a surface-associated glutamate dehydrogenase and construction of a glutamate dehydrogenase-deficient isogenic mutant. Infect. Immun. 62:1358-1368. 120. Joe, A., A. Yamamoto, and B. C. McBride. 1993. Characterization of recombinant and native forms of a cell surface antigen of Porphyromonas (Bacteroides) gingivalis. Infect. Immun. 61:3294-3303. 121. Johne, B., I. Olsen, and K. Bryn. 1988. Fatty acids and sugars in lipopolysaccharides from Bacteroides intermedius, Bacteroides gingivalis, and Bacteroides loescheii. Oral Microbiol. Immunol. 3:22-27. 122. Johnson, J. L., and L. V. Holdeman. 1983. Bacteroides intermedius comb. nov. and descriptions of Bacteroides corporis, and Bacteroides levii sp. nov. Int. J. Syst. Bacteriol. 33:15-25. 123. Kadowaki, T., M. Yoneda, K. Okamoto, K. Maeda, and K. Yamamoto. 1994. Purification and characterization of a novel arginine-specific cysteine proteinase (argingipain) involved in the pathogenesis of periodontal disease from the culture supernatant of Porphyromonas gingivalis. J. Biol. Chem. 269:21371-21378. 124. Kagan, J. M. 1980. Local immunity to Bacteroides gingivalis in periodontal disease. J. Dent. Res. 59:1750-1756. 125. Kaminishi, H., T. Cho, T. Itoh, A. Iwata, K. Kawasaki, Y. Hagihara, and H. Maeda. 1993. Vascular permeability enhancing activity of Porphyromonas gingivalis protease in guinea pigs. FEMS Microbiol. Lett. 114:109-114. 126. Kapur, V., M. W. Majesky, L.-L. Li, R. A. Black, and J. M. Musser. 1993. Cleavage of interleukin 1(3 (TL-ip) precursor to produce active IL-ip by a conserved 130 extracellular cysteine protease from Streptococcus pyogenes. Proc. Natl. Acad. Sci. USA. 90:7676-7680. 127. Karunakaran, T., and S. C. Holt. 1993. Cloning of two distinct hemolysin genes from Porphyromonas (Bacteroides) gingivalis in Escherichia coli. Microb. Pathog. 15:37-49. 128. Kastelein, P., T. J. M. van Steenbergen, J. M. Bras, and J. de Graaff. 1981. An experimentally induced phlegmonous abscess by a strain of Bacteroides gingivalis in guinea pigs and mice. Antonie van Leeuwenhoek. 47:1-9. 129. Kato, T., N. Takahashi, and H. K. Kuramitsu. 1992. Sequence analysis and characterization of the Porphyromonas gingivalis prtC gene, which expresses a novel collagenase activity. J. Bacteriol. 174:3889-3895. 130. Kawata, Y., S. Hanazawa, S. Amano, Y. Murakami, T. Matsumoto, K. Nishida, and S. Kitano. 1994. Porphyromonas gingivalis fimbriae stimulate bone resorption in vitro. Infect. Immun. 62:3012-3016. 131. Kay, H. M., A. J. Birss, and J. W. Smalley. 1989. Glycylprolyl dipeptidase activity of Bacteroides gingivalis W50 and the avirulent variant W50/BEI. FEMS Microbiol. Lett. 57:93-96. 132. Kay, H. M., A. J. Birss, and J. W. Smalley. 1990. Haemagglutinating and haemolytic activity of the extracellular vesicles of Bacteroides gingivalis W50. Oral Microbiol. Immunol. 5:269-274. 133. Kiel, R. A., K. S. Kornman, and P. B. Robertson. 1983. Clinical and microbiological effects of localized ligature-induced periodontitis on non-ligated sites in the cynomolgus monkey. J. Periodontal Res. 18:200-211. 134. Kiley, P. A., and S. C. Holt. 1980. Characterization of the lipopolysaccharide from Actinobacillus actinomycetemcomitans. Infect. Immun. 30:862-873. 135. Kilian, M. 1981. Degradation of immunoglobulins Al, A2, and G by suspected principal periodontal pathogens. Infect. Immun. 34:757-765. 136. Kinder, S. A., and S. C. Holt. 1989. Characterization of coaggregation between Bacteroides gingivalis T22 and Fusobacterium nucleatum T18. Infect. Immun. 57:3425-3433. 137. Klimpel, K. W., and V. L. Clark. 1990. The RNA polymerase of Porphyromonas gingivalis and Fusobacterium nucleatum are unrelated to the RNA polymerase of Escherichia coli. J. Dent. Res. 69:1567-1572. 138. Kolenbrander, P. E., and R. N. Andersen. 1989. Inhibition of coaggregation between Fusobacterium nucleatum and Porphyromonas {Bacteroides) gingivalis by lactose and related sugars. Infect. Immun. 57:3204-3209. 139. Koomey, J. M., R. E. Gill, and S. Falkow. 1982. Genetic and biochemical analysis of gonococcal IgAl protease: Cloning in Escherichia coli and construction of mutants of gonococci that fail to produce the activity. Proc. Natl Acad. Sc. USA. 79:7881-7885. 131 140. Kumada, H., K. Watanabe, T. Umemoto, Y. Haishima, S. Kondo, and K. Hisatsune. 1988. Occurrence of O-phosphorylated 2-keto-3-deoxyoctonate in lipopolysaccharide of Bacteroides gingivalis. FEMS Microbiol. Lett. 51:77-80. 141. Laemmli, U. K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature (London). 277:680-685. 142. Lala, A., A. Amano, H. T. Sojar, S. J. Radel, and E. De Nardin. 1994. Porphyromonas gingivalis trypsin-like protease: A possible natural ligand for the neutrophil formyl peptide receptor. Biochem. Biophys. Res. Commun. 199:1489-1496. 143. Lambe, D. W., Jr., K. P. Ferguson, and W. R. Mayberry. 1982. Characterization of Bacteroides gingivalis by direct fluorescent antibody staining and cellular fatty acid profiles. Can. J. Microbiol. 28:367-374. 144. Lantz, M. S., R. D. Allen, P. Bounelis, L. M. Switalski, and M. Hook. 1990. Bacteroides gingivalis and Bacteroides intermedins recognize different sites on human fibrinogen. J. Bacteriol. 172:716-726. 145. Lantz, M. S., R. D. Allen, P. Ciborowski, and S. C. Holt. 1993. Purification and immunolocalization of a cysteine protease from Porphyromonas gingivalis. J. Periodontal Res. 28:467-469. 146. Lantz, M. S., R. D. Allen, L. W. Duck, J. L. Blume, L. M. Switalski, and M. Hook. 1991. Identification of Porphyromonas gingivalis components that mediate its interactions with fibronectin. J. Bacteriol. 173:4263-4270. 147. Lantz, M. S., R. D. Allen, T. A. Vail, L. M. Switalski, and M. Hook. 1991. Specific cell components of Bacteroides gingivalis mediate binding and degradation of human fibrinogen. J. Bacteriol. 173:495-504. 148. Lantz, M. S., R. W. Rowland, L. M. Switalski, and M. Hook. 1986. Interactions of Bacteroides gingivalis with fibrinogen. Infect. Immun. 54:654-658. 149. Larjava, H., V.-J. Uitto, E. Eerola, and M. Haapasalo. 1987. Inhibition of gingival fibroblast growth by Bacteroides gingivalis. Infect. Immun. 55:201-205. 150. Larjava, H., V.-J. Uitto, M. Haapasalo, J. Heino, and M. Vuento. 1987. Fibronectin fragmentation induced by dental plaque and Bacteroides gingivalis. Scand. J. Dent. Res. 95:308-314. 151. Laughon, B. E., S. A. Syed, and W. J. Loesche. 1982. API ZYM system for identification of Bacteroides spp., Capnocytophaga spp., and spirochetes of oral origin. J. Clin. Microbiol. 15:97-102. 152. Laughon, B. E., S. A. Syed, and W. J. Loesche. 1982. Rapid identification of Bacteroides gingivalis. J. Clin. Microbiol. 15:345-346. 153. Lawson, D. A., R. Haas, J. Meyle, and T. F. Meyer. 1990. Molecular analysis of periodontal pathogens. Arch. Oral Biol. 35:101S-105S. 154. Lawson, D. A., and T. F. Meyer. 1992. Biochemical characterization of Porphyromonas (Bacteroides) gingivalis collagenase. Infect. Immun. 60:1524-1529. 132 155. Lee, J.-Y., H. T. Sojar, G. S. Bedi, and R. J. Genco. 1992. Synthetic peptides analogous to the fimbrillin sequence inhibit adherence of Porphyromonas gingivalis. Infect Immun. 60:1662-1670. 156. Lee, J.-Y., H. T. Sorja, G. S. Bedi, and R. J. Genco. 1991. Porphyromonas (Bacteroides) gingivalis fimbrillin: size, amino-terminal sequence, and antigenic heterogeneity. Infect. Immun. 59:383-389. 157. Lupine, G., and A. Progulske-Fox. 1993. Molecular biology, p. 293-319. In H. N. Shah, D. Mayrand, and R. Genco (ed.), Biology of the species Porphyromonas gingivalis. CRC Press, Inc., Boca Raton, Fl. 158. Lev, M., and A. F. Milford. 1971. Vitamin K stimulation of sphingolipid synthesis. Biochem. Biophys. Res. Commun. 45:358-362. 159. Li, J., R. P. Ellen, C. I. Hoover, and J. R. Felton. 1991. Association of proteases of Porphyromonas (Bacteroides) gingivalis with its adhesion to Actinomyces viscosus. J. Dent. Res. 70:82-86. 160. Listgarten, M. A. 1987. Nature of periodontal diseases: Pathogenic mechanisms. J. Periodontal Res. 22:172-178. 161. Loesche, W. J. 1976. Chemotherapy of dental plaque infections. Oral Sci. Rev. 9:65-107. 162. Loesche, W. J., K. U. Paunio, M. P. Woolfolk, and R. N. Hockett. 1974. Collagenolytic activity of dental plaque associated with periodontal pathology. Infect. Immun. 9:329-336. 163. Loesche, W. J., S. A. Syed, E. C. Morrison, B. Langhon, and N. S. Grossman. 1981. Treatment of periodontal infections due to anaerobic bacteria with short term treatment with metronidazole. J. Clin. Periodontol. 8:29-44. 164. Loesche, W. J., S. A. Syed, E. Schmidt, and E. C. Morrison. 1985. Bacterial profiles of subgingival plaques in periodontitis. J. Periodontol. 56:447-456. 165. Loomer, P. M., B. Sigusch, B. Sukhu, R. P. Ellen, and H. C. Tenenbaum. 1994. Direct effects of metabolic products and sonicated extracts of Porphyromonas gingivalis 2561 on osterogenesis in vitro. Infect. Immun. 62:1289-1297. 166. MacDonald, J. B., and R. J. Gibbons. 1962. The relationship of indigenous bacteria to periodontal disease. J. Dent. Res. 41:320-326. 167. MacDonald, J. B., S. S. Socransky, and R. J. Gibbons. 1963. Aspects of the pathogenesis of mixed anaerobic infections of mucous membranes. J. Dent. Res. 42:529-544. 168. Madden, T. E., T. M. Thompson, and V. L. Clark. 1992. Expression of Porphyromonas gingivalis proteolytic activity in Escherichia coli. Oral Microbiol. Immunol. 7:349-356. 169. Malek, R., J. G. Fisher, A. Caleca, M. Stinson, C. J. van Oss, J.-Y. Lee, M.-I. Cho, R. J. Genco, R. T. Evans, and D. W. Dyer. 1994. Inactivation of the Porphyromonas 133 gingivalis fimA gene blocks periodontal damage in gnotobiotic rats. J. Bacterid. 176:1052-1059. 170. Maley, J., and I. S. Roberts. 1994. Characterization of IS1126 from Porphyromonas gingivalis W83: a new member of the IS4 family of insertion sequence elements. FEMS Microbiol. Lett. 123:219-224. 171. Maley, J., N. B. Shoemaker, and I. S. Roberts. 1992. The introduction of colonic -Bacteroides shuttle plasmids into Porphyromonas gingivalis: Identification of a putative P. gingivalis insertion-sequence element. FEMS Microbiol. Lett 93:75-82. 172. Mansheim, B. J., and D. L. Kasper. 1977. Purification and immunochemical characterization of the outer membrane complex of Bacteroides melaninogenicus subspecies asaccharolyticus. J. Infect. Dis. 135:787-799. 173. Mansheim, B. J., A. B. Onderdonk, and D. L. Kasper. 1978. Immunochemical and biological studies of the lipopolysaccharide of Bacteroides melaninogenicus subspecies asaccharolyticus. J. Immunol. 120:72-78. 174. Markaryan, A., I. Morozova, H. Yu, and P. E. Kolattukudy. 1994. Purification and characterization of an elastic metalloprotease from Aspergillus fumigatus and immunoelectron microscopic evidence of secretion of this enzyme by the fungus invading the murine lung. Infect. Immun. 62:2149-2157. 175. Markkanen, H., S. M. Syrjanen, and P. Alakuijala. 1986. Salivary IgA, lysozyme and (^microglobulin in periodontal disease. Scand. J. Dent. Res. 94:115-120. 176. Marsh, P. D., A. S. McKee, and A. S. McDermid. 1988. Effect of haemin on enzyme activity and cytotoxin production by Bacteroides gingivalis W50. FEMS Microbiol. Lett 55:87-92. 177. Matsudaira, P. 1990. Limited N-terminal sequence analysis. Methods Enzymol. 182:602-613. 178. Mayrand, D. 1989. Biological activities of outer membrane vesicles. Can. J. Microbiol. 35:607-613. 179. Mayrand, D., G. Bourgeau, D. Grenier, and J. M. Lacroix. 1984. Properties of oral asaccharolytic bluck-pigmented Bacteroides. Can. J. Microbiol. 30:1133-1136. 180. Mayrand, D., and D. Grenier. 1985. Detection of collagenase activity in oral bacteria. Can. J. Microbiol. 31:134-138. 181. Mayrand, D., and S. C. Holt. 1988. Biology of asaccharolytic black-pigmented Bacteroides species. Microbiol. Rev. 52:134-152. 182. Mayrand, D., and B. C. McBride. 1980. Ecological relationships of bacteria involved in a simple, mixed anaerobic infection. Infect. Immun. 27:44-55. 183. Mayrand, D., B. C. McBride, T. Edwards, and S. Jensen. 1980. Characterization of Bacteroides asaccharolyticus and B. melaninogenicus oral isolates. Can. J. Microbiol. 26:1178-1183. 134 184. McBride, B. C , A. Joe, and U. Singh. 1990. Cloning of Bacteroides gingivalis surface antigens involved in adherence. Arch. Oral Biol. 35:59S-68S. 185. Mead, D. A., E. Szczesna-Skorupa, and B. Kemper. 1986. Single-stranded DNA 'blue' T7 promoter plasmids: a versatile tandem promoter system for cloning and protein engineering. Protein Engineering, h 67-74. 186. Millar, S. J., E. G. Goldstein, M. J. Levine, and E. Hausmann. 1986. Modulation of bone metabolism by two chemically distinct lipopolysaccharide fractions from Bacteroides gingivalis. Infect. Immun. 51:302-306. 187. Minhas, T., and J. Greenman. 1989. Production of cell-bound and vesicle-associated trypsin-like protease, alkaline phosphatase and iV-acetyl-/?-glucosaminidase by Bacteroides gingivalis strain W50. J. Gen. Microbiol. 135:557-564. 188. Miyauchi, T., M. Hayakawa, and Y. Abiko. 1989. Purification and characterization of glycylprolyl aminopeptidase from Bacteroides gingivalis. Oral Microbiol. Immunol. 4:222-226. 189. Morales, T. I., and J. F. Woessner Jr. 1977. PZ-peptidase from chick embryos. J. Biol. Chem. 252:4855-4860. 190. Mouton, C., D. Bouchard, M. DesLaurieres, and L. LaMoude. 1989. Immunochemical identification and preliminary characterization of a nonfimbrial hemagglutinating adhesin of Bacteroides gingivalis. Infect. Immun. 57:566-573. 191. Mouton, C , D. ni Eidhin, M. DesLauriers, and L. Lamy. 1991. Hemagglutinating adhesin HA-Ag2 of Bacteroides gingivalis is distinct from fimbrillin. Oral Microbiol. Immunol. 6:6-11. 192. Mouton, C. T., P. G. Hammond, J. Slots, and R. J. Genco. 1981. Serum antibodies to oral Bacteroides asaccharolyticus {Bacteroides gingivalis): relationships to age and periodontal disease. Infect. Immun. 31:182-192. 193. Nair, B. C , W. R. Mayberry, R. Dziak, P. B. Chen, M. J. Levine, and E. Hausmann. 1983. Biological effects of a purified lipopolysaccharide from Bacteroides gingivalis. J. Periodontal Res. 18:40-49. 194. Naito, Y., and R. J. Gibbons. 1988. Attachment of Bacteroides gingivalis to collagenous substrata. J. Dent. Res. 67:1075-1080. 195. Naito, Y., H. Tohda, K. Okuda, and I. Takazoe. 1993. Adherence and hydrophobicity of invasive and noninvasive strains of Porphyromonas gingivalis. Oral Microbiol. Immunol. 8:195-202. 196. Nakamura, S., A. Takeuchi, Y. Masamoto, Y. Abiko, M. Hayakawa, and H. Takiguchi. 1992. Cloning of the gene encoding a glycylprolyl aminopeptidase from Porphyromonas gingivalis. Arch. Oral Biol. 37:807-812. 197. Nakayama, K. 1990. The superoxide dismutase-encoding gene of the obligately anaerobic bacterium Bacteroides gingivalis. Gene. 96:149-150. 135 198. Neiders, M. E., P. B. Chen, H. Suido, H. S. Reynolds, J. J. Zambon, M. Shlossman, and R. J. Genco. 1989. Heterogeneity of virulence among strains of Bacteroides gingivalis. J. Periodontal Res. 24:192-198. 199. Ng, S. K., and S. S. Fung. 1984. PZ-peptidase activities of some black-pigmented Bacteroides species. Can. J. Microbiol. 30:1305-1308. 200. Ng, W., and J. Tonzetich. 1983. Effects of H2S on permeability of oral mucosa. J. Dent. Res. 62:275, Abst. No. 953. 201. Nilsson, T., J. Carlsson, and G. Sundqvist. 1985. Inactivation of key factors of the plasma proteinase cascade systems by Bacteroides gingivalis. Infect. Immun. 50:467-471. 202. Nishikata, M., and F. Yoshimura. 1991. Characterization of Porphyromonas (Bacteroides) gingivalis hemagglutinin as a protease. Biochem. Biophys. Res. Commun. 178:336-342. 203. Nishikata, M., F. Yoshimura, and Y. Nodasaka. 1989. Possibility of Bacteroides gingivalis hemagglutinin possessing protease activity revealed by inhibition studies. Microbiol. Immunol. 33:75-80. 204. Norqvist, A., B. Norrman, and H. Wolf-Watz. 1990. Identification and characterization of a zinc metalloprotease associated with invasion by the fish pathogen Vibrio anguillarum. Infect. Immun. 58:3731-3736. 205. O'Farell, P. H. 1975. High resolution two-dimensional electrophoresis of proteins. J. Biol. Chem. 250:4007-4021. 206. Oakley, B. R., D. T. Kirsch, and N. T. Morris. 1980. A simplified silver stain for detecting proteins in polyacrylamide gels. Anal. Biochem. 105:361-363. 207. Odelson, D. A., J. L. Rasmussen, C. J. Smith, and F. L. Macrina. 1987. Extrachromosomal systems and gene transmission in anaerobic bacteria. Plasmid. 17:87-109. 208. Ogawa, T., and S. Hamada. 1994. Hemagglutinating and chemotactic properties of synthetic peptide segments of fimbria! protein from Porphyromonas gingivalis. Infect. Immun. 62:3305-3310. 209. Ohno, S., Y. Emori, S. Imajoh, H. Kawasaki, M. Kisaragi, and K. Suzuki. 1984. Evolutionary origin of a calcium-dependent protease by fusion of genes for a thiol protease and a calcium-binding protein? Nature. 312:566-570. 210. Okuda, K., and T. Kato. 1987. Hemagglutinating activity of lipopolysaccharides from subgingival plaque. Infect. Immun. 55:3192-3199. 211. Okuda, K., T. Kato, Y. Naito, I. Takazoe, Y. Kikuchi, T. Nakamura, T. Kiyoshiga, and S. Sasaki. 1988. Protective efficacy of active and passive immunizations against experimental infections with Bacteroides gingivalis. J. Dent. Res. 67:807-811. 212. Okuda, K., J. Slots, and R. J. Genco. 1981. Bacteroides gingivalis, Bacteroides asaccharolyticus, and Bacteroides melaninogenicus subspecies: Cell surface 136 morphology and adherence to erythrocytes and human buccal epithelial cells. Curr. Microbiol. 6:7-12. 213. Okuda, K., and J. Takazoe. 1974. Hemagglutinating activity of Bacteroides melaninogenicus. Arch. Oral Biol. 19:415-416. 214. Okuda, K., A. Yamamoto, Y. Naito, I. Takazoe, J. Slots, and R. J. Genco. 1986. Purification and properties of hemagglutinin from culture supernatant of Bacteroides gingivalis. Infect. Immun. 54:659-665. 215. Oliver, W. W., and W. B. Wherry. 1921. Notes on some bacterial parasites of the human mucous membranes. J. Infect. Dis. 28:341-345. 216. Otogoto, J.-L, and H. K. Kuramitsu. 1993. Isolation and characterization of the Porphyromonas gingivalis prtT gene, coding for protease activity. Infect. Immun. 61:117-123. 217. Otsuka, M., J. Endo, D. Hinode, A. Nagata, R. Maehara, M. Sato, and R. Nakamura. 1987. Isolation and characterization of protease from culture supernatant of Bacteroides gingivalis. J. Periodontal Res. 22:491-498. 218. Page, R. C , and H. E. Schroeder. 1973. Biochemical aspects of the connective tissue alterations in inflammatory gingival and periodontal disease. Int. Dent. J. 23:455-469. 219. Paster, B. J., G. J. Fraser, F. E. Dewhirst, and I. Olsen. 1991. Phylogeny of Bacteroides and related bacteria. J. Dent. Res. 70:318, Abst. No. 425. 220. Pekovic, D. D., and E. D. Fillery. 1984. Identification of bacteria in immunopathological mechanisms of human periodontal diseases. J. Periodontal Res. 19:329-351. 221. Pike, R., W. McGraw, J. Potempa, and J. Travis. 1994. Lysine- and arginine-specific proteinases from Porphyromonas gingivalis. J. Biol. Chem. 269:406-411. 222. Pratt, J. M., G. J. Boulnois, V. Darby, E. Orr, E. Wahle, and I. B. Holland. 1981. Identification of gene products programmed by restriction endonuclease DNA fragments using an E. coli in vitro system. Nucleic Acids Res. 9:4459-4474. 223. Progulske-Fox, A., A. Oberste, C. Drummond, and W. P. McArthur. 1989. Transfer of plasmid pE5-2 from Escherichia coli to Bacteroides gingivalis and B. intermedins. Oral Microbiol. Immunol. 4:132-134. 224. Progulske-Fox, A., S. Tumwasorn, and S. C. Holt. 1989. The expression and function of a Bacteroides gingivalis hemagglutinin gene in Escherichia coli. Oral Microbiol. Immunol. 4:121-131. 225. Putnam, F. W. 1975. Alpha, Beta, Gamma, Omega - The roster of the plasma proteins, p. 57-131. In F. W. Putnam (ed.), The Plasma Proteins. Structure, function and genetic control. Vol. 1. Academic Press, Inc., New York. 226. Renart, J., and I. V. Sandoval. 1984. Western blots. Methods Enzymol. 104:455-460. 137 227. Rizza, V., P. R. Sinclair, D. C. White, and P. R. Courant. 1968. Electron transport system of the protoheme-requiring anaerobe Bacteroides melaninogenicus. J. Bacterid. 96:665-671. 228. Robertson, P. B., M. Lantz, P. T. Marucha, K. S. Kornman, C. L. Trummel, and S. C. Holt. 1982. Collagenolytic activity associated with Bacteroides species and Actinobacillus actinomycetemcomitans. J. Periodontal Res. 17:275-283. 229. Robillard, N. J., F. P. Tally, and M. H. Malamy. 1985. Tn4400, a compound transposon isolated from Bacteroides fragilis functions in Escherichia coli. J. Bacterid. 164:1248-1255. 230. Rodriguez, R. L., and R. C. Tait. 1983. Recombinant DNA Techniques, An Introduction. Addison-Wesley Publishing Company, Inc., Reading, Massachusetts. 231. Roeterink, C. H., T. J. M. van Steenbergen, W. F. B. de Jong, and J. de Graaff. 1984. Histopathological effects in the palate of the rat induced by injection with different black-pigmented Bacteroides strains. J. Periodontal Res. 19:292-302. 232. Rotstein, O. D., P. E. Nasmitsh, and S. Grinstein. 1987. The Bacteroides by-product succinic acid inhibits neutrophil respiratory burst by reducing intracellular pH. Infect. Immun. 55:864-870. 233. Rotstein, O. D., T. L. Pivett, V. D. Fiegel, R. D. Nelson, and R. L. Simmons. 1985. Succinic acid, a metabolic by-product of Bacteroides species, inhibits polymorphonuclear leukocyte function. Infect. Immun. 48:402-408. 234. Roy, T. E., and C. D. Kelly. 1939. Genus VIII. Bacteroides Catellani and Chalmers, p. 556-569. In D. H. Bergey, R. S. Breed, E. G. D. Murray, and A. P. Hitchens (ed.), Bergey's Manual of Determinative Bacteriology, 5th ed. The Williams and Wilkins Co., Baltimore. 235. Saglie, F. R., A. Marfany, and P. Camargo. 1988. Intragingival occurrence of Actinobacillus actinomycetemcomitans and Bacteroides gingivalis in active destructive periodontal lesions. J. Periodontol. 59:259-265. 236. Saglie, F. R., J. Pertuiser, M. T. Rezende, M. Nestor, A. Marfany, and J. Cheung. 1988. In situ correlative immuno-identification of mononuclear infiltrates and invasive bacteria in disease gingiva. J. Periodontol. 59:688-696. 237. Saglie, R., F. A. Carranza, and M. G. Newman. 1985. The presence of bacteria within the oral epithelium in periodontal disease. I. A scanning and transmission electron microscopic study. J. Periodontol. 56:618-624. 238. Sakai, D. K. 1985. Loss of virulence in a protease-deficient mutant of Aeromonas salmonicida. Infect. Immun. 48:146-152. 239. Sakanari, J. A., C. E. Staunton, A. E. Eakin, and C. S. Craik. 1989. Serine proteases from nematode and protozoan parasites: Isolation of sequence homologs using generic molecular probes. Proc. Natl. Acad. Sci. USA. 86:4863-4867. 240. Sako, K., I. Takazoe, and K. Okuda. 1988. Isolation and characterization of plasmid DNA from Bacteroides strains isolated from the oral cavity. Oral Microbiol. Immunol. 3:72-76. 138 241. Salyers, A. A., N. B. Shoemaker, and E. P. Guthrie. 1987. Recent advance in Bacteroides genetics. CRC Crit. Rev. Microbiol. 14:49-71. 242. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular Cloning: a Laboratory Manual (2nd Ed.). Cold Spring Harbor Laboratory, Cold Spring Harbor, N. Y. 243. Sandholm, L. 1986. Proteases and their inhibitors in chronic inflammatory periodontal disease. J. Clin. Periodontal. 13:19-26. 244. Sato, M., M. Otsuka, R. Maehara, J. Endo, and R. Nakamura. 1987. Degradation of immunoglobulin A by protease isolated from the anaerobic periodontopathogenic bacterium, Bacteroides gingivalis. Arch. Oral Biol. 32:235-238. 245. Sawyer, S. J., J. B. MacDonald, and R. J. Gibbons. 1962. Biochemical characteristics of Bacteroides melaninogenicus. A study of thirty-one strains. Arch. Oral Biol. 7:685-691. 246. Schenkein, H. A. 1988. The effect of periodontal proteolytic Bacteroides species on proteins of the human complement system. J. Periodontal Res. 23:187-192. 247. Schenkein, H. A. 1991. Complement factor D-like activity of Porphyromonas gingivalis W83. Oral Microbiol. Immunol. 6:216-220. 248. Scott, C. F., E. J. Whitaker, B. F. Hammond, and R. W. Colman. 1993. Purification and characterization of a potent 70-kDa thiol lysyl-proteinase (Lys-gingivain) from Porphyromonas gingivalis that cleaves kininogens and fibrinogen. J. Biol. Chem. 268:7935-7942. 249. Seddon, S. V., H. N. Shah, J. M. Hardie, and J. P. Robinson. 1988. Chemically defined and minimal media for Bacteroides gingivalis. Curr. Microbiol. 17:147-149. 250. Shah, H. N., R. Bonnet, B. Matteen, and R. A. Williams. 1979. The porphyrin pigmentation of subspecies of Bacteroides melaninogenicus. Biochem. J. 180:45-50. 251. Shah, H. N., and D. M. Collins. 1990. Prevotella, a new genus to include Bacteroides melaninogenicus and related species formerly classified in the genus Bacteroides. Int. J. Syst. Bacteriol. 40:205-208. 252. Shah, H. N., and M. D. Collins. 1988. Proposal for reclassification of Bacteroides asaccharolyticus, Bacteroides gingivalis, and Bacteroides endodontalis in a new genus, Porphyromonas. Int. J. Syst. Bacteriol. 38:128-131. 253. Shah, H. N., and S. E. Gharbia. 1989. Ecological events in subgingival dental plaque with reference to Bacteroides and Fusobacterium species. Infection. 17:264-268. 254. Shah, H. N., and S. E. Gharbia. 1989. Lysis of erythrocytes by the secreted cysteine proteinase'of Porphyromonas gingivalis W83. FEMS Microbiol. Lett. 61:213-218. 255. Shah, H. N., S. E. Gharbia, D. Kowlessur, E. Wilkie, and K. Brocklehurst. 1990. Isolation and characterization of gingivain, a cysteine proteinase from Porphyromonas gingivalis strains W83. Biochem. Soc. Trans. 18:578-579. 139 256. Shah, H. N., S. E. Gharbia, D. Kowlessur, E. Wilkie, and K. Brocklehurst. 1991. Gingivain; A cysteine proteinase isolated from Porphyromonas gingivalis. Microbial Ecol. Health Dis. 4:319-328. 257. Shah, H. N., S. E. Gharbia, and C. M. OToole. 1992. Assessment of the relative cytotoxicity of Porphyromonas gingivalis cells, products, and components on human epithelial cell lines. J. Periodontal. 63:44-51. 258. Shah, H. N., S. E. Gharbia, A. Progulske-Fox, and K. Brocklehurst. 1992. Evidence for independent molecular identity and functional interaction of the haemagglutinin and cysteine proteinase (gingivain) of Porphyromonas gingivalis. J. Med. Microbiol. 36:239-244. 259. Shah, H. N., and R. A. D. Williams. 1987. Catabolism of aspartate and asparagine by Bacteroides intermedins and Bacteroides gingivalis. Curr. Microbiol. 15:313-318. 260. Shah, H. N., and R. A. D. Williams. 1987. Utilization of glucose and amino acids by Bacteroides intermedins and Bacteroides gingivalis. Curr. Microbiol. 15:241-246. 261. Shah, H. N., R. A. D. Williams, G. D. Bowden, and J. M. Hardie. 1976. Comparison of the biochemical properties of Bacteroides melaninogenicus from human dental plaque and other sites. J. Appl. Bacteriol. 41:473-492. 262. Shoemaker, N. B., R. D. Barber, and A. A. Salyers. 1989. Cloning and characterization of a Bacteroides conjugal tetracycline-erythromycin resistance element by using a shuttle cosmid vector. J. Bacteriol. 171:1294-1302. 263. Shoemaker, N. B., C. Getty, J. F. Gardner, and A. A. Salyers. 1986. Tn4351 transposes in Bacteroides spp. and mediates integration of plasmid R751 into the Bacteroides chromosome. J. Bacteriol. 165:929-936. 264. Shoemaker, N. B., C. Getty, E. P. Guthrie, and A. A. Salyers. 1986. Two Bacteroides plasmids, pBFTMlO and pB8-51, contain transfer regions that are recognized by broad-host-range IncP plasmids and by a conjugative Bacteroides tetracycline resistance element. J. Bacteriol. 166:959-965. 265. Shoemaker, N. B., E. P. Guthrie, A. A. Salyers, and J. F. Gardner. 1985. Evidence that the clindamycin-erythromycin resistance gene of Bacteroides plasmid pBF4 is on a transposable element. J. Bacteriol. 162:626-632. 266. Silhavy, T. J., M. J. Berman, and L. W. Enquist. 1984. Experiments with gene fusions. Cold Spring Harbor Laboratory, Cold Spring Harbor. 267. Singer, R. E., and B. A. Buckner. 1981. Butyrate and propinate: important component of toxic dental plaque extracts. Infect. Immun. 32:458-463. 268. Singh, U. 1989. The adherence properties of Bacteroides gingivalis, Ph.D. Theis, University of British Columbia, Vancouver, Canada, 269. Singh, U., D. Grenier, and B. C. McBride. 1989. Bacteroides gingivalis vesicles mediate attachment of streptococci to serum-coated hydroxyapatite. Oral Microbiol. Immunol. 4:199-203. 140 270. Slots, J. 1977. Microflora in the healthy gingival sulcus in man. Scand. J. Dent. Res. 85:247-254. 271. Slots, J. 1977. The predominant cultivable microflora of advanced periodontitis. Scand. J. Dent Res. 85:114-121. 272. Slots, J. 1979. Subgingival microflora and periodontal disease. J. Clin. Periodontal. 6:351-382. 273. Slots, J. 1981. Enzymatic characterization of some oral and nonoral gram-negative bacteria with the API ZYM system. J. Clin. Microbiol. 14:288-294. 274. Slots, J. 1982. Importance of black-pigmented Bacteroides in human periodontal disease, p. 27-45. In R. J. Genco, and S. E. Mergenhagen (ed.), Host-Parasite Interactions in Periodontal Disease. American Society for Microbiology, Washington, D.C. 275. Slots, J., L. Bragd, M. Wikstrom, and G. Dahlen. 1986. The occurrence of Actinobacillus actinomycetemcomitans, Bacteroides gingivalis and Bacteroides intermedins in destructive periodontal disease in adults. J. Clin. Periodontol. 13:570-577. 276. Slots, J., and G. Dahlen. 1985. Subgingival microorganisms and bacterial virulence factors in periodontitis. Scand. J. Dent. Res. 93:119-127. 277. Slots, J., and R. J. Genco. 1979. Direct hemagglutination technique for differentiating Bacteroides asaccharolyticus oral strains form non-oral strains. J. Clin. Microbiol. 10:371-373. 278. Slots, J., and R. J. Gibbons. 1978. Attachment of Bacteroides melaninogenicus subsp. asaccharolyticus to oral surfaces and its possible role in colonization of the mouth and of periodontal pockets. Infect. Immun. 19:254-264. 279. Slots, J., and M. A. Listgarten. 1988. Bacteroides gingivalis, Bacteroides intermedins and Actinobacillus actinomycetemcomitans in human periodontal diseases. J. Clin. Periodontol. 15:85-93. 280. Slots, J., and M. A. Taubman. 1992. Contemporary Oral Microbiology and Immunology. Mosby-Year Book, Inc., St. Louis. 281. Smalley, J. W., and A. J. Birss. 1987. Trypsin-like enzyme activity of the extracellular membrane vesicles of Bacteroides gingivalis W50. J. Gen. Microbiol. 133:2883-2894. 282. Smalley, J. W., and A. J. Birss. 1990. Stability of soluble and extracellular vesicle-associated trypsin-like protease (TLP) activity of Bacteroides gingivalis W50. FEMS Microbiol. Lett. 70:317-320. 283. Smalley, J. W., and A. J. Birss. 1991. Extracellular vesicle-associated and soluble trypsin-like enzyme fractions of Porphyromonas gingivalis W50. Oral Microbiol. Immunol. 6:202-208. 141 284. Smalley, J. W., A. J. Birss, H. M. Kay, A. S. McKee, and P. D. Marsh. 1989. The distribution of trypsin-like enzyme activity in cultures of a virulent and an avirulent strain of Bacteroides gingivalis W50. Oral Microbiol. Immunol. 4:178-181. 285. Smalley, J. W., A. J. Birss, and C. A. Shuttleworth. 1988. The degradation of type I collagen and human plasma fibronectin by the trypsin-like enzyme and extracellular membrane vesicles of Bacteroides gingivalis W50. Arch. Oral Biol. 33:323-329. 286. Smalley, J. W., D. Mayrand, and D. Grenier. 1993. Vesicles, p. 259-292. In H. N. Shah, D. Mayrand, and R. J. Genco (ed.), Biology of the species of Porphyromonas gingivalis. CRC Press, Inc., Boca Raton. 287. Smith, A. J., T. Minhas, J. Greenman, and G. Embery. 1993. The distribution and properties of some hydrolytic enzymes from Porphyromonas gingivalis W50. Microbios. 73:185-197. 288. Smith, D. E., and P. A. Fisher. 1984. Identification, developmental regulation, and response to heat shock of two antigenically related forms of a major nuclear envelope protein in Drosophilia embryos: application of an improved method for affinity purification of antibodies using polypeptides immobilized on nitrocellulose blots. J. Cell Biol. 99:20-28. 289. Sojar, H. T., J.-Y. Lee, G. S. Bedi, M.-I. Choi, and R. J. Genco. 1991. Purification, characterization, and immunization of fimbrial protein from Porphyromonas (Bacteroides) gingivalis. Biochem. Biophys. Res. Commun. 175:713-719. 290. Sojar, H. T., J.-Y. Lee, G. S. Bedi, and R. J. Genco. 1993. Purification and characterization of a protease from Porphyromonas gingivalis capable of degrading salt-solubilized collagen. Infect. Immun. 61:2369-2376. 291. Sorimachi, H., S. Imajoh-Ohmi, Y. Emori, H. Kawasaki, S. Ohno, Y. Minami, and K. Suzuki. 1989. Molecular cloning of a novel mammalian calcium-dependent protease distinct from both m- and u-types. J. Biol. Chem. 264:20106-20111. 292. Sorimachi, H., S. Ishiura, and K. Suzuki. 1993. A novel tissue-specific calpain species expressed predominantly in the stomach comprises two alternative splicing products with and without Ca2+-binding domain. J. Biol. Chem. 268:19476-19482. 293. Sorimachi, H., T. C. Saido, and K. Suzuki. 1994. New era of calpain research. FEBS Lett. 343:1-5. 294. Sorsa, T., V.-J. Uitto, K. Suomalainen, H. Turto, and S. Lindy. 1987. A trypsin-like protease from Bacteroides gingivalis: partial purification and characterization. J. Periodontal Res. 22:375-380. 295. Studier, F. W., and B. A. Moffatt. 1986. Use of bacteriophage T7 RNA polymerase to direct selective high-level expression of cloned genes. J. Mol. Biol. 189:113-130. 296. Suido, H., M. E. Neiders, P. K. Barua, M. Nakamura, P. A. Mashimo, and R. J. Genco. 1987. Characterization of N-CBz-glycyl-glycyl-arginyl peptidase and glycyl-prolyl peptidase of Bacteroides gingivalis. J. Periodontal Res. 22:412-418. 142 297. Suido, H., J. J. Zambon, P. A. Mashimo, R. Dunford, and R. J. Genco. 1988. Correlation between gingival crevicular fluid enzymes and the subgingival microflora. J. Dent. Res. 67:1070-1074. 298. Sundqvist, G., A. Bengtson, and J. Carlsson. 1988. Generation and degradation of the complement fragment C5a in human serum by Bacteroides gingivalis. Oral Microbiol. Immunol. 3:103-107. 299. Sundqvist, G., J. Carlsson, and L. Hanstrom. 1987. Collagenolytic activity of black -pigmented Bacteroides species. J. Periodontal Res. 22:300-306. 300. Sundqvist, G., J. Carlsson, B. Herrmann, and A. Tarnvik. 1985. Degradation of human immunoglobulins G and M and complement factors C3 and C5 by black -pigmented Bacteroides. J. Med. Microbiol. 19:85-94. 301. Sundqvist, G., D. Figdor, L. Hanstrom, S. Sorlin, and G. Sandstrom. 1991. Phagocytosis and virulence of different strains of Porphyromonas gingivalis. Scand. J. Dent. Res. 99:117-129. 302. Sundqvist, G. K., J. Carlsson, B. F. Herrmann, J. F. Hofling, and A. Vaatainen. 1984. Degradation in vivo of the C3 protein of guinea-pig complement by a pathogenic strain of Bacteroides gingivalis. Scand. J. Dent. Res. 92:14-24. 303. Sundqvist, G. K., M. I. Eckerbom, A. P. Larsson, and U. T. Sjogren. 1979. Capacity of anaerobic bacteria from necrotic dental pulps to induce purulent infections. Infect. Irnmun. 25:685-693. 304. Sveen, K. 1977. The capacity of lipopolysaccharides from Bacteroides, Fusobacterium and Veillonella to produce skin inflammation and the local and generalized Schwartzman reaction in rabbit. J. Periodontal Res. 12:340-350. 305. Sveen, K., T. Hofstad, and K. C. Milner. 1977. Lethality for mice and chick embryos, pyrogenicity in rabbits and ability to gelate lysate from ameobocytes of Limulus polyphemus by lipopolysaccharides from Bacteroides, Fusobacterium and Veillonella. ActaPatho. Microbiol. Scand. Sect. B. 85:388-396. 306. Sveen, K., and N. Skaug. 1980. Bone resorption stimulated by lipopolysaccharides from Bacteroides, Fusobacterium, and Veillonella, and by the lipid A and the polysaccharide part of Fusobacterium lipopolysaccharide. Scand. J. Dent. Res. 88:535-542. 307. Tabor, S., and C. C. Richardson. 1985. A bacteriophage T7 RNA polymerase/promoter system for controlled exclusive expression of specific genes. Proc. Natl. Acad. Sci. USA. 82:1074-1078. 308. Takada, H., J. Mihara, I. Morisaki, and S. Hamada. 1991. Induction of interleukin-1 and -6 in human gingival fibroblast cultures stimulated with Bacteroides lipopolysaccharides. Infect. Immun. 59:295-301. 309. Takahashi, N., T. Kato, and H. K. Kuramitsu. 1991. Isolation and preliminary characterization of the Porphyromonas gingivalis prtC gene expressing collagenase activity. FEMS Microbiol. Lett. 84:135-138. 143 310. Takazoe, L, T. Nakamura, and K. Okuda. 1984. Colonization of the subgingival area by Bacteroides gingivalis. J. Dent. Res. 63:422-426. 311. Takazoe, I., M. Tanaka, and T. Homma. 1971. A pathogenic strains of Bacteroides melaninogenicus. Arch. Oral Biol. 16:817-822. 312. Tally, F. P., D. R. Snydman, M. J. Shimell, and M. H. Malamy. 1982. Characterization of pBFTMlO, a clindamycin-erythromycin resistance transfer factor from Bacteroides fragilis. J. Bacterid. 151:686-691. 313. Tamura, M., M. Tokuda, S. Nagaoka, and H. Takada. 1992. Lipopolysaccharides of Bacteroides intermedins {Prevotella intermedia) and Bacteroides (Porphyromonas) gingivalis induce interleukin-8 gene expression in human gingival fibroblast cultures. Infect. Immun. 60:4932-4937. 314. Tang, C. M., J. Cohe, T. Krausz, S. V. Nooden, and D. W. Holden. 1993. The alkaline protease of Aspergillus fumigatus is not a virulence determinant in two murine models of invasive pulmonary aspergillosis. Infect. Immun. 61:1650-1656. 315. Tanner, A. C. R., C. Haffer, G. T. Bratthall, R. A. Visconti, and S. S. Socransky. 1979. A study of the bacteria associated with advancing periodontitis in man. J. Clin. Periodontol. 6:278-307. 316. Tanner, A. C. R., S. S. Socransky, and J. M. Goodson. 1984. Microbiota of periodontal pockets losing crestal alveolar bone. J. Periodontal Res. 19:279-291. 317. Terato, K., Y. Nagai, K. Kawanishi, and S. Yamamoto. 1976. A rapid assay method of collagenase activity using 14C-labeled soluble collagen as substrate. BBA. 445:753-762. 318. Tew, J. G., D. R. Marshall, J. A. Barmeister, and R. R. Rammey. 1985. Relationship between gingival crevicular fluid and serum antibody titres in young adults with generalized and localized periontitis. Infect. Immun. 49:487-493. 319. Tippet, B., K. W. Frisken, and J. R. Tagg. 1987. Quantification of the bacterium Bacteroides gingivalis in human dental plaque by detection of the trypsin-like protease activity of colonies imprinted on membrane filters. Arch. Oral Biol. 32:151-152. 320. Tisljar, U. 1993. Thimet oligopeptidase - a review of a thiol dependent metallo-endopeptidase also known as Pz-peptidase endopeptidase 24.15 and endo-oligopeptidase. Biol. Chem. Hoppe-Seyler. 374:91-100. 321. Toda, K., M. Otsuka, Y. Ishikawa, M. Sato, Y. Yamamoto, and R. Nakamura. 1984. Thiol-dependent collagenolytic activity in culture media of Bacteroides gingivalis. J. Periodontal Res. 19:372-381. 322. Tonzetich, J., and B. C. McBride. 1981. Characterization of volatile sulphur production by pathogenic and non-pathogenic strains of oral Bacteroides. Arch. Oral Biol. 26:963-969. 323. Touw, J. J. A., T. J. M. van Steenbergen, and J. de Graaff. 1982. Butyrate: a cytotoxin for vero cells produced by Bacteroides gingivalis and Bacteroides asaccharolyticus. Antonie van Leeuwenhoek. 48:315-325. 144 324. Travis, J., and G. S. Salvesen. 1983. Human plasma proteinase inhibitors. Annu. Rev. Biochem. 52:655-709. 325. Tsutsui, H., T. Kinouchi, Y. Wakano, and Y. Ohnishi. 1987. Purification and characterization of a protease from Bacteroides gingivalis 381. Infect. Immun. 55:420-427. 326. Uitto, V.-J. 1983. Degradation of basement membrane collagen by proteinases from human gingiva, leukocytes and bacterial plaque. J. Periodontol. 54:740-745. 327. Uitto, V.-J., M. Haapasalo, T. Laakso, and T. Salo. 1988. Degradation of basement membrane collagen by protease from some anaerobic microorganisms. Oral Microbiol. Immunol. 3:97-102. 328. Uitto, V.-J., H. Larjava, J. Heino, and T. Sorsa. 1989. A protease of Bacteroides gingivalis degrades cell surface and matrix glycoproteins of cultured gingival fibroblasts and induces secretion of collagenase and plasminogen activator. Infect. Immun. 57:213-218. 329. Uitto, V.-J., and A.-M. Raeste. 1978. Activation of latent collagenase of human leukocytes and gingival fluid by bacterial plaque. J. Dent. Res. 57:844-851. 330. Van Dyke, T. E., S. Offenbacher, D. Place, V. R. Dowell, and J. Jones. 1988. Refractory periodontitis: Mixed infection with Bacteroides gingivalis and other unusual Bacteroides species. J. Periodontol. 59:184-189. 331. van Steenbergen, T. J. M., and J. de Graaff. 1986. Proteolytic activity of black -pigmented Bacteroides strains. FEMS Microbiol. Lett. 33:219-222. 332. van Steenbergen, T. J. M., J. J. De Soet, and J. de Graaff. 1979. DNA base composition of various strains of Bacteroides melaninogenicus. FEMS Microbiol. Lett. 5:127-130. 333. van Steenbergen, T. J. M., F. G. A. Delemarre, F. Namavar, and J. de Graaff. 1987. Differences in virulence within the species Bacteroides gingivalis. Antonie van Leeuwenhoek. 53:233-244. 334. van Steenbergen, T. J. M., M. D. den Ouden, J. J. A. Touw, and J. de Graaff. 1982. Cytotoxic activity of Bacteroides gingivalis and Bacteroides asaccharolyticus. J. Med. Microbiol. 5:253-258. 335. van Steenbergen, T. J. M., P. Kastelein, J. J. A. Touw, and J. de Graaff. 1982. Virulence of black-pigmented Bacteroides strains from periodontal pockets and other sites in experimentally induced skin lesions in mice. J. Periodontal Res. 17:41-49. 336. van Steenbergen, T. J. M., F. Namavar, and J. de Graaff. 1985. Cheminoiluminescence of human leukocytes by black-pigmented Bacteroides strains from dental plaque and other sites. J. Periodontal Res. 20:58-71. 337. van Steenbergen, T. J. M., A. J. van Winkelhoff, D. Mayrand, D. Granier, and J. de Graaff. 1984. Bacteroides endodontalis sp. nov., an asaccharolytic black-pigmented Bacteroides species from infected dental root canals. Int. J. Syst. Bacteriol. 34:118-120. 145 338. van Winkelhoff, A. J., T. J. M. van Steenbergen, and J. de Graaff. 1988. The role of black-pigmented Bacteroides in human oral infections. J. Clin. Periodontol. 15:145-155. 339. Vincent, J. W., J. B. Suzuki, W. A. Falker Jr., and W. C. Cornett. 1985. Reaction of human sera from juvenile periodontitis, rapidly progression periodontitis, and adult periodontitis patients with selected periodontopathogens. J. Periodontol. 56:464-469. 340. Wahren, A., and R. J. Gibbons. 1970. Amino acid fermentation by Bacteroides melaniogenicus. Antonie van Leeuwenhoek. 36:149-159. 341. Washington, O. R., M. Deslauriers, D. P. Stevens, L. K. Lyford, S. Haque, Y. Yan, and P. M. Flood. 1993. Generation and purification of recombinant fimbrillin from Porphyromonas (Bacteroides) gingivalis 381. Infect. Immun. 61:1040-1047. 342. Watanabe, K.-L, T. Takasawa, F. Yoshimura, M. Ozeki, M. Kawanami, and H. Kato. 1992. Molecular cloning and expression of a major surface protein (the 75-kDa protein) of Porphyromonas (Bacteroides) gingivalis in Escherichia coli. FEMS Microbiol. Lett 92:47-56. 343. White, D., and D. Mayrand. 1981. Association of oral Bacteroides with gingivitis and adult periodontitis. J. Periodontal Res. 16:259-265. 344. Williams, G. D., and S. C. Holt. 1985. Characteristics of the outer membranes of selected oral Bacteroides species. Can. J. Microbiol. 31:238-250. 345. Wilson, M. 1993. Biological activities of lipopolysaccharide and endotoxin, p. 171-197. In H. N. Shah, D. Mayrand, and R. J. Genco (ed.), Biology of the species Porphyromonas gingivalis. CRC Press, Inc., Boca Raton. 346. Wilton, J. M. A., T. J. Hurst, and E. E. Scott. 1993. Inhibition of polymorphonuclear leucocyte phagocytosis by Porphyromonas gingivalis culture products in patients with adult periodontitis. Arch. Oral Biol. 38:285-289. 347. Wingrove, J. A., R. G. DiScipio, Z. Chen, J. Potempa, J. Travis, and T. E. Hugli. 1992. Activation of complement components C3 and C5 by a cysteine proteinase (gingipain-1) from Porphyromonas (Bacteroides) gingivalis. J. Biol. Chem. 267:18902-18907. 348. Winkler, J. R., V. Matarese, C. I. Hoover, R. H. Kramer, and P. A. Murray. 1988. An in vitro model to study bacterial invasion of periodontal tissue. J. Periodontol. 59:40-45. 349. Woese, C. R. 1987. Bacterial evolution. Microbiol. Rev. 51:221-271. 350. Wiinsch, E., and H.-G. Heidrich. 1963. Zur quantitativen Bestimmung der Kollagenase. Zeitschrift Fur Physiologische Chemie. 333:149-151. 351. Yamazaki, K., F. Ikarashi, T. Aoyagi, K. Takahashi, T. Nakajima, K. Hara, and G. J. Seymour. 1992. Direct and indirect effects of Porphyromonas gingivalis lipopolysaccharide on interleukin-6 production by human gingival fibroblasts. Oral Microbiol. Immunol. 7:218-224. 146 352. Yoneda, M., K. Maeda, and M. Aono. 1990. Suppression of bactericidal activity of human polymorphonuclear leukocytes by Bacteroides gingivalis. Infect. Immun. 58:406-411. 353. Yoshimoto, H., Y. Takahashi, N. Hamada, and T. Umemoto. 1993. Genetic transformation of Porphyromonas gingivalis by electroporation. Oral Microbiol. Immunol. 8:208-212. 354. Yoshimoto, H., and T. Umemoto. 1990. Characterization and physical mapping of plasmids of black-pigmented Sactero/d&s. Oral Microbiol. Immunol. 5:208-212. 355. Yoshimura, F., M. Nishikata, T. Suzuki, C. I. Hoover, and E. Newbrun. 1984. Characterization of a trypsin-like protease from the bacterium Bacteroides gingivalis isolated from human dental plaque. Arch. Oral Biol. 29:559-564. 356. Yoshimura, F., T. Sugano, M. Kawanami, H. Kato, and T. Suzuki. 1987. Detection of specific antibodies against fimbriae and membrane proteins from the oral anaerobe Bacteroides gingivalis. Microbiol. Immunol. 31:935-941. 357. Yoshimura, F., K. Takahashi, Y. Nodasaka, and T. Suzuki. 1984. Purification and characterization of a novel type of fimbriae from the oral anaerobic Bacteroides gingivalis. J. Bacteriol. 160:949-957. 358. Yoshimura, F., Y. Takahashi, E. Hibi, T. Takasawa, H. Kato, and D. P. Dickinson. 1993. Proteins with molecular masses of 50 and 80 kilodaltons encoded by genes downstream from the fimbrilin gene (fimA) are components associated with fimbriae in the oral anaerobe Porphyromonas gingivalis. Infect. Immun. 61:5181-5189. 359. Zambon, J. J., H. Reynolds, J. G. Fisher, M. Shlossman, R. Dunford, and R. J. Genco. 1988. Microbiological and immunological studies of adult periodontitis in patients with noninsulin-dependent diabetes mellitus. J. Periodontol. 59:23-31. 360. Zambon, J. J., H. S. Reynolds, and J. Slots. 1981. Black-pigmented Bacteroides spp. in the human oral cavity. Infect. Immun. 32:198-203. 147 

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