1. Molecular Analysis of a P-N-Acetyl-Hexosaminidase
Gene From Porphyromonas gingivalis W83
Archie Lovatt
Thesis presented for the degree of Doctor of Philosophy
University of Leicester
1994
4. Contents
Chapter 1 Introduction
1.1 Porphyromonas gingivaUs
1.1.1 General Biology
1.1.2 Role in Periodontal Disease
1.2 Bacterial Adhesion and Colonisation
1.2.1 Dental Plaque
1.2.2 Bacterial Colonisation Factors and their Interactions
9
9
10
1.3 The Degradation of Host Components and Extracellular
Matrix Molecules 14
1.3.1 Proteolytic Degradation 15
1.3.1.1 The Role of Proteolytic Enzymes from P. gingivalis 15
1.3.1.2 The Activation of Host Proteases by P. gingivalis 17
1.3.2 The Breakdown of Glycosaminoglycans by Exoglycosidases 18
1.4 P-N-Acetyl-Hexosaniinidases and
P-N-Acetyl-Glucosaminidases 22
1.4.1 Specificity 22
1.4.2 Possible Roles in Glycoconjugate Degradation 24
1.4.2.1 The Modification of Host Extracellular Matrix
Glycosaminoglycans 24
1.4.2.2 The Degradation of Glycoproteins and Glycolipids 26
1.4.2.3 The Hydrolysis of Asparagine-Linked Oligosaccharides on
Human IgG 29
1.4.2.4 Autolysins that Degrade Peptidoglycan 31
1.4.2.5 P-N-Acetyl-Hexosaminidases in Glycobiology 33
1.5 The Aims of This Th^is 36
5. Chapter 2 Materials and Methods 37
2.1 Bacterial Strains and Plasmids 37
2.1.1 Growth Condition and Media 37
2.2 Transformation of Bacterial Cells 39
2.2.1 Production of Competent Cells 39
2.2.1.1 Calcium Chloride Method 39
2.2.1.2 Electrotransformation Method 40
2.2.2 Transformation with Plasmid DNA 40
2.2.2.1 Calcium Chloride Method 40
2.2.2.2 Electrotransformation Method 41
2.2.3 Transformation with Bacteriophage DNA 41
2.3 Procedures for DNA Extraction 42
2.3.1 Extraction of Chromosomal DNA 42
2.3.2 Small Scale Extraction of Plasmid DNA 43
2.3.3 Large Scale Extraction of Plasmid DNA 43
2.3.4 Extraction of M13mpl8/19 Template DNA 44
2.3.5 Phenol Extraction and Ethanol Precipitation 45
2.4 Techniques used in Routine DNA Manipulation 45
2.5 DNA Hybridisation Proœdures 46
2.5.1 Transfer of DNA to Nylon Filters 46
2.5.2 Preparation of Filters for Colony Hybridisation 47
2.5.3 Production of RadiolabelW Probe 48
2.5.4 Hybridisation of DNA Immobilised on Filters with the Probe 48
2.6 DNA Sequencing 49
2.7 Polymerase Chain Reaction Proœdures 51
2.8 Radioactive Labelling of Proteins 52
2.8.1 Minicell Analysis 52
2.8.2 DNA Directed Transcription-Translation System 53
6. 2.9 Biochemical Assay of P-N-Acetyl-GIucosaminidase
and P-N-Acetyl-Galactmiaminidase Activity 54
2.10 Computer Analysis 55
2.10.1 Database Searching and Multiple Sequence Alignment 55
2.9.2 The Hydropathy Profile 56
Chapter 3 The Cloning and Expression of the nah Region
from P. gingivaUs W83 in E. coli 57
3.1 Introduction 57
3.2 Results 58
3.2.1 Detection of P-N-Acetyl-Hexosaminidase Activity in
f . gmghufw W83 58
3.2.2 Isolation and Restriction Endonuclease Analysis of the
nah Region from P. gingivalis W83 59
3.2.2.1 Construction of a P. gingivalis W83 Expression Library
in E. coli 59
3.2.2 2 Screening of the P. gingivalis W83 Gene Library
for Exo-P-N-Acetyl-Glucosaminidase Activity 61
3.2.2.3 Restriction Endonuclease Analysis of the nah Region 63
3.2.2.4 Southern Blot Analysis of Chromosomal DNA 65
3.2.3 Localisation and Expression of the Gene on pALl that
Encodes for P-N-Acetyl-Hexosaminidase Activity 67
3.2.3.1 Localisation on pALl 67
3.2.3 2 IPTG-Induced Expression of P-N-Acetyl-Hexosaminidase 69
3.2.3.3 Expression of Proteins in a Cell-Free and Minicell System 70
3.3 Discussion 72
IV
7. Chapter 4 Molecular Characterisation of the nah Region
by DNA Sequence Analysis 75
4.1 Introduction 75
4.2
4.2.1
4.2.1.1
4.2.1.2
4.2.1.3
4.2. 1.4
4.2. 1.5
4.3
Results 76
DNA Sequence and Computer Analysis of the nah Region
from P. gingivalis W83 76
The DNA Sequencing Strategy 76
The DNA Sequence of the nahA Gene and the Predicted
Amino-Acid Sequence of the P-N-Acetyl-Hexosaminidase
Protein (NahA) 76
Homology Between NahA and Other Enzymes 87
The NahA Protein of P. gingivalis is Homologous to the
Central Domain of CrylA(c) 6-endotoxin from Bacillus
thuringiensis 97
Analysis of orfl 98
Discussion 106
Chapter 5 Further Studies on the P-N-Acetyl-
Hexosaminidase From P. gingivalis W83 114
5.1 Introduction 114
5.2 Results and Discussion 115
5.2.1 Construction of Plasmid pMUT for the Generation of a
nahA Isogenic Mutant of P. gingivalis 115
5.2.2 Strategy for the Generation of a Glutathione
S-transferase-NahA Fusion Protein 120
9. List of Abbreviations
Amp Ampicillin
Cc Clindamycin
EDTA Ethylenediaminetetraacetic acid
Em Erythromycin
ECM Extracellular matrix
Fuc Fucose
GAG Glycosaminoglycan
Gal Galactose
GalNAc N-Acetyl-galactosamine
GlcA Glucuronic acid
GlcNAc N-Acetyl-glucosamine
IdoA Iduronic acid
IPTG Isopropyl-P-D-galactopyranoside
Kan Kanamycin
Man Mannose
MES 2-[N-Morpholino]ethanesulfonic acid
PEG Polyethylene glycol
SA Sialic acid
Sm Streptomycin
TEMED N, N, N', N'-Tetramethylethylenediamine
10. Acknowledgements
A special thanks to Dr I. S. Roberts for his continuous supervision,
guidance, help and encouragement throughout all of this work. I would also
like to thank past and present members of the Department of Microbiology and
Immunology, University of Leicester for their help and expertise. I thank Dr C.
Pazzani, A. Wallace and G. Rigg for their endless advice, assistance and
friendship. I thank Dr J. Milner, J. Maley, R. Pearce, Dr T. McGenity, R.
Willmouth and A. K. Barton and for their help, friendship and interest. Thanks
also to Dr J. Eastgate, S. Gordon, D. Simpson, Dr M. Camara, A. Smith, R.
Feely, A. Owtrowski, Dr C. R. Drake, N. Taylor, Dr C. Petite, Dr S. O’
Brien, Dr. R. Morse, Dr M. Coleman, B. J. Roberts, Dr J. Hill, A. Sheperd,
Dr T. Mitchell, P. Esumeh and J. R. Canvin for helping me in a number of
situations. For those who gave friendship, advice or inspiration, I extend my
thanks. I thank the Medical Research Council for my stipend and funding my
attendance at conferences. My extra special thanks goes to my parents for
everything they have done and continue to do.
vm
11. Abstract
Molecular Analysis of a P-N-Acetyl-Hexosaminidase Gene From
Porphyronwnas gingivaUs W83
Archie Lovatt
The black-pigmented Gram-negative anaerobe Porphyromonas gingivalis has
been implicated in human pericxlontal diseases and expresses a number of
exoglycosidase activities that may be capable of degrading the oligosaccharides
from host glycoconjugates. The first stage in characterising the role of the p-N-
Acetyl-hexosaminidase activity from this microorganism was the cloning of the
nahA gene from P. gingivalis W83 which encodes for a protein with this
enzyme activity. The nahA gene was cloned in E.coli by constructing a plasmid
expression library of 5fl«3A-generated P. gingivalis W83 chromosomal DNA
fragments. Expression of p-N-Acetyl-hexosaminidase activity was detectW by
cleavage of the fluorogenic substrates 4-melhylumbelliferyl-N-Acetyl-P-D-
glucosaminide and 4-methylumbelliferyl-N-Acetyl-P-D-galactosaminide.
Southern blot analysis suggested that the nahA gene was present as a single
copy in all P. gingivalis strains tested. In contrast, sequences homologous to
the nahA gene were not detectable in the closely related species P. endodontalis
and P. asaccharolytica. The nahA gene was 2331 base pairs long and encoded
a protein of 111 amino acids with a predicted molecular weight of 87 kDa. A
characteristic signal peptide for an acylated lipoprotein was present at the
amino terminus suggesting that the mature p-N-Acetyl-hexosaminidase may be
a lipoprotein attached to the outer membrane of P. gingivalis. Protein
homology studies suggested that NahA and p-N-Acetyl-hexosaminidases from
eukaryotes and prokaryotes contain homologous active site domains with
similar catalytic arginine residues. DNA sequence analysis 5’ to the nahA gene
identified another open reading frame, and a potential hairpin structure which
may be involved in regulating gene expression. Lastly, a suicide plasmid that
may allow the site specific inactivation of the nahA gene on the chromosome of
P. gingivalis was constructed. The results presented in this thesis may
contribute to future studies including the generation of an isogenic mutant of P.
gingivalis lacking p-N-Acetyl-hexosaminidase activity.
12. CHAPTER 1
Introduction
1.1 Porphyromonas gingivaUs
1.1.1 General Biology
Black-pigmented Gram-negative anaerobes were originally describe as
Bacterium melaninogenicum, producing dark brown-black pigmented colonies
when grown for 6-10 days on blood agar plates (Oliver and Wherry, 1921).
This species was shown to be very heterogenous, comprising of both
saccharolytic and asaccharolytic strains (Sawyer et al, 1962). These strains
were assigned to a single species, Bacteroides melanimgenicus, which
containW the three subspecies, melaninogenicus (saccharolytic), intermedius
(moderately saccharolytic) and asaccharolyticus (asaccharolytic). (Holdeman
and Moore, 1974). The saccharolytic (fermentative) species now belong to the
genus Prevotella and the asaccharolytic (non fermentative) species have been
placed into the genus Porphyromonas, which includes Porphyromonas
endodontalis, Porphyromonas asaccharolytica and Porphyromonas gingivalis
(Finegold et al., 1993; Kulekci et al., 1993; van Steenbergen et al., 1993).
When P. gingivalis is grown under chemostat conditions, the cells utilise
arginine, cysteine, histidine, serine and tryptophan as their carbon and energy
source, whereas sugars inhibit growth rate (Mayrand and Holt, 1988). P.
gingivalis utilises a polyaspartate/glutamate heteropolymer at a slower rate than
aspartate homopolymer (Shah and Gharbia, 1993). When grown on peptides die
yield of cells is greater than that grown on amino acids (Shah and Gharbia,
13. 1993). Moreover, when grown in an equimolar mixture of polyaspartate and
free aspartate, the uptake of free aspartate appears to be suppressed until there
is utilisation of the homopolymer. Such observations imply that peptides are the
favoured substrates for growth of P. gingivalis and amino acids are utilised to
lesser extent (Shah and Gharbia, 1993).
P. gingivalis requires vitamin K and the iron compound, haemin, for growth
(Gibbons and MacDonald, 1960; Mayrand and Holt, 1988). It has been
proposed that haemin and vitamin K function as electron carriers in the electron
transport system of P. gingivalis (Gibbons and MacDonald, 1960; Mayrand and
Holt, 1988). The black pigment produced by P. gingivalis has been described
as a ‘haemoglobin derivative’ and has been shown to be protohaemin with
traces of protoporphyrin (Duerden, 1975; Shah et al., 1979). The concentration
of haemin available for growth appears to have an effect on the physiology and
virulence of P. gingivalis (McKee et al., 1986). For example, P. gingivalis
cells grown with no haemin in their media are avirulent when injected into
mice, whereas those grown in haemin excess cause 100% mortality in mice
(MC Kee et al., 1986). Furthermore, those grown in haemin excess appear to
have more fimbriae than those grown under haemin limitation (M^ Kee et al.,
1986). A 26 IcDa protein is predominant on the cell surface of P. gingivaUs
when grown under haemin limitation, but is not expressed under haemin
excess. This protein, termed Omp26, appears to be exported to the outer
membrane for haemin binding and imported across the outer membrane for
haemin transport (Bramanti and Holt, 1992a & b).
Isolates of P. gingivalis from in vivo infections are often associated with a
mixed bacterial population (Mayrand and Holt, 1988). The presence of vitamin
K-related compounds, such as naphthoquinone, produced by associated bacteria
may enhance the growth of P. gingivalis (MacDonald et al., 1963). Succinate
14. has been shown to replace vitamin K or haemin as a growth factor for P.
gingivalis (Mayrand and Holt, 1988). When produced by Treponema denticola
via fermentation of amino acids, succinate may act as a growth factor for P.
gingivalis in vivo (Grenier, 1992a). Moreover, succinate may be incorporated
into the lipids and phospholipids of the cell envelope and is suggested to play an
important role in bacterial nutritional interaction during periodontal disease
(Lev et al., 1971; Grenier, 1992a).
1.1.2 Ro!e in Periodontal Disease
Periodontal disease is a collective term for a variety of conditions
characterised by the inflammation and degeneration of the gingivae, connective
tissue, periodontal ligament, cementum and the alveolar bone that supports
teeth. While gingivitis refers to inflammation that is confined to the gingival
tissues, periodontitis is considered to be an advancement of gingivitis into the
bone that surrounds the teeth. The initial symptoms of gingivitis are
enlargement and inflammation of the soft tissue with bleeding of the gums. The
inflammatory reaction results in swelling of the tissue and the formation of
periodontal pockets between the teeth and their supporting tissue (Hirsch and
Clarke, 1989; Tortora and Anagnostakos, 1990). At the advanced stage,
extensive degradation of the host extracellular matrix (ECM) takes place,
together with loss of alveolar bone and tooth support (Uitto, 1991). A
hypothetical representation of chronic adult periodontal disease is shown in
Figure 1.1.
15. Figure 1.1 A Hypothetical Representation of Chronic Adult Periodontal
Disease
Enamel
Ine pathological
periodontal pocket
GE
EPITHELIAL CELLS COLLAGEN ANCHOR
CELLULAR
MATRIX
GE=GmGIVAL EPITHELIUM
Cl^CONNEClTVE TISSUE
AB=ALVEOLAR BONE
PDL=PEmODONrAL LIGAMENT
CE=CEMENTUM
BASEMENT MEMBRANE
Figure 1.1 Hypothetical representation of chronic adult
periodontitis. The structure of the periodontium is shown. The small
boxed region of the periodontal pocket is shown in more detail
within the large box. Dental plaque bacteria colonise the periodontal
pocket and their toxins/enzymes/extracellular vesicles may gain
access from the pocket to the subepithelium. All host cells are
surrounded by a complex extracellular matrix. During disease,
bacteria may directly degrade host extracellular matrix and immune
components and indirectly trigger host-mediated tissue degradative
pathways (Constructed from information present in Saglie et al.,
1988; Holt and Bramanti, 1991; Uitto, 1991; Lamont et al., 1992;
Birkedal-Hansen, 1993; Couchman and Woods, 1993; Duncan et
al., 1993; Sandros et al., 1993).
16. The pathogenesis of periodontal diseases is thought to be a complex series of
events that involves both host factors and dental plaque. Although periodontal
diseases are multifactorial in nature, studies indicate that the inflammation and
destruction of tissues is initiated and maintained by the bacteria of dental plaque
(Lindhe et al., 1975; Christersson et al., 1991; Corbet and Davies, 1993;
Offenbacher et al., 1993; Ready and Jeffcoat, 1993). Dental plaque is a highly
organised mixture of bacterial species and the co-aggregation or cell-to-cell
recognition of genetically distinct partner-types is essential for its formation
(Kolenbrander, 1988; Kolenbrander and London, 1993). The estimated
composition of dental plaque is over 300 bacterial species, but only certain
types of Gram-negative anaerobes are associated with the various forms of
periodontal diseases (Christersson et al., 1991; Holt and Bramanti, 1991). For
example, Actinobacillus actinomycetemcomitans is associated with localised
juvenile periodontitis, Prevotella intermedius!Treponema denticola with acute
necrotizing ulcerative gingivitis and Capnocytophaga spp. with juvenile
diabetes advanced periodontitis (Holt and Bramanti, 1991). Porphyromonas
gingivalis has received considerable attention in past years and is thought to
play an important role in the formation of lesions during chronic adult
periodontal disease (Slots and Genco, 1984; Holt and Bramanti, 1991).
P. gingivalis is found in adult periodontitis lesions at high frequency and
increased numbers. However, in plaque from healthy patients P. gingivalis is
either not detected or shows significantly lower frequency and numbers
(Christersson et al., 1991; Dahlen, 1993). Therefore, it appears that an
ecological niche of P. gingivalis is the diseased periodontal pocket that occurs
between the tooth and its supporting tissue. Patients suffering from periodontitis
exhibit high levels of antibodies that are specific for P. gingivalis (Ebersole et
al., 1985; De Nardin et al., 1991; Lopatin and Blackburn, 1992; Kinane et al.,
1993), and eradication of this organism from the subgingival microflora
17. correlates with resolution of the disease (Loesche et al., 1981; van Dyke et al.,
1988). Several reports demonstrate that P. gingivalis can invade gingival and
oral epithelial cells in vitro (Lamont et al., 1992; Duncan et al., 1993; Sandros
et al., 1993). Such invasion and the detection of P. gingivalis within gingival
tissue (Saglie et al., 1988), may suggest that this organism penetrates the
mucosal epithelial barrier. Although the invasive mechanisms are not fully
understood, it is clear that P. gingivalis expresses an arsenal of potential
virulence factors (Holt and Bramanti, 1991).
Virulence factors that may aid colonisation, tissue destruction and
impairment of host defence mechanisms have been proposed for P.gingivalis.
These include extracellular hydrolytic enzymes such as trypsin-like protease,
collagenase, neuraminidase, p-N-Acetyl-hexosaminidase and glycosulphatase,
bacterial surface components such as lipopolysaccharide (LPS), polysaccharide
capsule, fimbriae and haemagglutinins (Slots and Genco, 1984; Mmhas and
Greenman, 1989; Holt and Bramanti, 1991; Socransky and Haffajee, 1991;
Meghji et al., 1993; Slomiany et al., 1993; Sundqvist, 1993; van WinkeUioff et
al., 1993). hi addition, P. gingivalis releases outer-membrane vesicles (OMV)
into the extracellular environment when grown in vitro and may also secrete
these factors in vivo (Grenier and Mayrand, 1987; Mayrand and Holt, 1988).
The OMV structures possess haemagglutinating, haemolytic, proteolytic and
exoglycosidase activity, mediate bacterial co-aggregation and may activate
alveolar bone resorption (Grenier and Mayrand, 1987; Smalley and Birss,
1987; Minhas and Greenman, 1989; Bourgeau and Mayrand, 1990; Kay et al.,
1990a; Mihara et al., 1993). The small size of OMV may allow them to cross
epithelial barriers that are impermeable to bacterial cells, hi this way, they
could serve as vehicles for toxins and enzymes that extend the ability of the
bacterial cell to obtain nutrients (Grenier and Mayrand, 1987; Mayrand and
Holt, 1988). Moreover, since OMV are highly proteolytic, they have been
18. 7
implicated in the degradation and penetration of epithelial tissue by P.
gingivalis (Grenier and Mayrand, 1987). These potential virulence factors may
also compete for antibodies and inhibit specific antibacterial immune defence
mechanisms (Mayrand and Holt, 1988).
Many studies have assessed the pathogenicity of P. gingivalis using
experimental animal model systems (Holt and Bramanti, 1991; Sundqvist,
1993). The cynomolgus monkey model, Macaca fascicularis has a similar
periodontal morphology to that of humans, with gingivitis developing
spontaneously in the presence of calculus and plaque (Komman et al., 1981;
Holt et al., 1988; Birek et al., 1989; Nemeth et al., 1993). The implantation of
P. gingivalis into the periodontal microbiota of this monlcey results in high
levels of antibodies to this microorganism plus rapid and significant alveolar
bone loss (Holt et al., 1988). Immunisation of the cynomolgus monlcey with
Idlled P. gingivalis protects against such bone loss (Dahlen, 1993).
Periodontal destruction can also be induced in gnotobiotic rats by mono
infection with P. gingivalis. Periodontal destruction is estimated by measuring
horizontal and vertical bone changes in the animal’s periodontium (Klausen et
al., 1991). Immunisation of the gnotobiotic rat with Idlled P. gingivalis cells or
highly purified fimbriae before gingival challenge with this microorganism
results in a reduction of periodontal bone loss (Klausen et al., 1991; Evans et
al., 1992). Recently, infection with a nonfimbriated mutant of P. gingivalis 381
showed that this strain was unable to induce the extent of periodontal bone loss
that was observed with the wild type strain (Malek et al., 1994)
19. Subcutaneous injection of mice with invasive strains of P. gingivalis
produces spreading lesions that frequently result in death (Mayrand and Holt,
1988). Invasive strains of P. gingivalis spread to distant sites and produce
abdominal abscesses, whereas non-invasive strains produce localised lesions at
the challenged site (Mayrand and Holt, 1988; Genco et al., 1991; Naito et al.,
1993). Although the mouse system does not mimic the human pathological
periodontal pocket, this model may be useful when studying the pathogenic
mechanisms of P. gingivalis. For example, Genco et al., 1991 have described
the development of the mouse subcutaneous chamber model. Bacteria within
the chamber can be studied throughout the course of infection and the chamber
contents can be used to examine specific host factors that are produced in
response to P. gingivalis.
hi summary, P. gingivalis is implicated in human adult periodontitis and
studies indicate that alveolar bone loss or experimental periodontitis in animal
models can be induced by P. gingivalis. P. gingivalis is suggested to be an
opportunistic periodontopathogen that survives and multiplies within the
periodontal pocket, resisting host defence mechanisms and damaging host tissue
(Slots and Genco, 1984; Holt and Bramanti, 1991; Loos et al., 1992). The
pathogenicity of P. gingivalis is more than likely multifactorial, requiring
several virulence factors that may play an important role in the pathogenesis of
adult periodontal disease. As a result, the determination and analyses of factors
that may influence bacterial colonisation, the impairment of host defences and
the destruction of host tissue is essential to understanding the pathogenesis of
human periodontal disease.
20. 1.2 Bacterial Adhesion and Colonisation
1.2.1 Dental Plaque
Dental plaque development on tooth surfaces begins with the precipitation of
a salivary pellicle (Christersson et al., 1991). The salivary pellicle is a thin coat
that covers the freshly cleaned tooth surface and consists of glycoproteins,
mucim and salivary enzymes (Mayhall, 1970; Kolebrander and London, 1993).
Bacterial colonisation of the salivary pellicle takes place rapidly and the first
microorganisms that attach to the tooth surface are mainly Streptococcus
species and Gram-positive rods. It is thought that facultative bacteria proliferate
first and create an environment suitable for the growth of anaerobes (Hirsch
and Clarke, 1989). As plaque matures, its composition becomes more complex
and the early colonising population diversifies to include Actinomyces,
Capnocytophaga, Haemophilus, Prevotella and Fusobacterium species
(Kolenbrander and London, 1993). Late colonisers which include A.
actinomycetemcomitans, P. gingivalis and T. denticola are linked to early
colonisers, such as A. israelii, C. gingivalis, H. parainfluenzae, Pr. loeochei
and S. oralis. The linkage that bridges the attachment of late colonisers to early
colonisers is thought to be Fusobacterium nucleatum (Kolenbrander and
London, 1993).
P. gingivalis attaches to a variety of bacteria from dental plaque, including
A. viscosus, A. naeslundii, A. israelii, S. sanguis, S. mitis, T. deiuicola and F.
nucleatum (Kolenbrander, 1990; Grenier, 1992b; Kolenbrander and London,
1993). The current idea is that P. gingivalis and T. denticola adhere to F.
nucleatum and to each other within bacterial plaque (Grenier, 1992b;
Kolenbrander and London, 1993). Moreover, the occurence of T. denticola in
diseased periodontal sites requires a detectable level of P. gingivalis (Simonson
21. 10
et al., 1992), and the co-aggregation and nutritional interaction between these
two microorganisms is thought to be important for the initiation and
progression of certain forms of periodontal disease (Grenier, 1992a & b; Nilius
et al, 1993). P. gingivalis may also provide growth factors for F. nucleatum,
which could explain why these two organisms frequently coexist in
periodontally diseased sites (Rogers et al., 1992).
1.2.2 Bacterial Colonisation Factors and their Interactions
P. gingivalis adheres to a variety of host components, including fibronectin-
collagen complexes (Naito and Gibbons, 1988), lactoferrin (Kalfas et al.,
1991), fibrinogen (Lantz et al., 1991), epithelial cells (Isogai et al., 1988) and
erythrocytes (Hoover et al., 1992b). Surface structures of P. gingivalis
proposed to be involved in adherence in vivo are fimbriae (Yosmimura et al.,
1984; Sharma et al., 1993), proteases (Grenier, 1992c; Hoover et al., 1992b;
Stinson et al., 1993) and haemagglutinins (Desluariers and Moutoun, 1992;
Dusek et al., 1993). Haemagglutinins of P. gingivalis have been reported, to
range in size (Chandad and Mouton, 1990; Dusek et al., 1993), may have
proteolytic activity (Grenier, 1992c; Hoover et al., 1992b), and form
complexes with proteases (Pilce et al., 1994). The protease-haemagglutinin
complexes of P. gingivalis may be involved in the adhesion and subsequent
hemolysis of host erythrocytes, thereby facilitating the acquisition of haemin in
vivo (Dusek et al., 1993; Pike et al., 1994). Besides agglutinating erythrocytes,
haemagglutinins may be involved in the aggregation of Actinomyces spp. that is
mediated by the OMV of P. gingivalis (Bourgeau and Mayrand, 1990).
Early studies indicated that fimbriae were associated with the
haemagglutination activity, but they are now believed not to be involved in this
process (Yoshimura et al., 1984 & 1985; Watanabe et al., 1992; Hamada et
22. 11
al., 1994; Malek et al., 1994). Fimbriae may play an important role as
adtiesins in vivo and bave been shown to be a major target for antibody
responses in patients with advanced periodontal disease (Yoshimura et al.,
1987). Fimbriae from P. gingivalis have been purified (Sojar et al., 1991), the
fimA gene has been characterised (Dickinson et al., 1988), and the structural
subunit fimbrülin has an apparent molecular weight of 43 IcDa (Washington et
al., 1993). The fimbriae of P. gingivalis are thought to be involved in the
adhesion to salivary pellicle, epithelial cells, collagen, periodontal ligament and
gingival fibroblasts (Watanabe et al., 1992; Naito et al., 1993). Naito et al.,
1993 suggest that the fimbriae of non-invasive strains (ATCC 33277, 381 and
Su63) have a higher relative hydrophobicity and stronger collagen binding
capacity than than the fimbriae firom invasive strains (ATCC 53977, ATCC
49417, 16-1 and W83). Further, a comparison of non-invasive and invasive
strains suggest that non-invasive strains have relatively more cell surface
hydrophobicity than invasive strains (Watanabe et al., 1992). These
observations have led to the hypothesis that non-invasive strains have fimbriae
which strongly attach to collagen in lesions, but because of their high cell
surface hydrophobicity can be readily phagocytosed. However, invasive strains
may bind weakly to collagen, and because of their high cell surface
hydrophilicity remain in lesions by avoiding phagocytosis (Naito et al., 1993).
Recently, Hamada et al., 1994 and Malek et al., 1994 have reported no change
in the relative cell surface hydrophobicity of fimA mutants of P. gingivalis
33277 and 381.
A number of investigators have described the construction and
characterisation offimA mutants of P. gingivalis (Hamada et al., 1994; Malek
et al., 1994). Hamada et al., 1994 showed that inactivation of the fimA gene m
P. gingivalis 33277 causes no alteration in haemagglutinating activity, however
decreases the adherence of this strain to human gingival fibroblasts. The
23. 12
adhesion of wM-type P. gingivalis to gingival fibroblasts causes changes in the
normal architechure of the fibroblast, with the appearance of long microvilli
surrounding large bacterial clumps. No such changes are observed with the
fimA mutant, which could imply that fimbriae trigger a sequence of events in
the fibroblast that facilitate bacterial contact (Hamada et al., 1994). hi a similar
report, Malek et al., 1994 have shown that Q.fimA mutant of P. gingivalis 381
has no change in haemagglutination, however is less able to bind saliva-coated
hydroxyapatite. Moreover, in the gnotobiotic rat model, this fimA mutant was
unable to induce the extent of periodontal bone loss that was observed with
wild-type strain (Malek et al., 1994). The failure of ihofimA mutant to cause
significant periodontal damage in gnotobiotic rats may be due to the inability of
the mutant to adhere to saliva-coated oral surfaces in the animal (Malek et al.,
1994). Alternatively, fimbriae may play a role in other reactions that are
important in periodontal disease. For example, they have been shown to
stimulate the release of interleuldn-1 (IL-1) from mouse monocytes (Hanazawa
et al., 1991). IL-1 stimulates osteoclastic bone resorption (Roodman, 1991),
and antibodies directed towards the fimbriae may inhibit their ability to
stimulate IL-1 production (Evans et al., 1992). This could explain why
immunisation of the gnotobiotic rat with highly purified fimbriae elicits an
immune response that interferes with bone loss induced by P. gingivalis (Evans
et al., 1992) (Section 1.1.2).
Many cell-to-cell adhesive interactions and bacterial co-aggregations can be
inhibited by the addition of simple sugars, suggesting that many adhesins are
lectin-like proteins. (Kolenbrander, 1988; Kolenbrander, 1989; Holt and
Bramanti, 1991; Kolebrander and London, 1993). The F. nucleatum-P.
gingivalis co-aggregation has been characterised and represents a typical
carbohydrate-lectin interaction. This interaction is inhibited by lactose and
appears to be mediated by a carbohydrate receptor on P. gingivalis that
24. 13
interacts with a 42 IcDa outer-membrane protein on F. nucleatum (Kinder and
Holt, 1993). The specific co-aggregation between F. gingivalis and T. denticola
is inhibited by D-galactosamine and arginine and is thought to be bimodal, that
is, both microorganisms contain specific adhesins that recognise complementary
receptors on the other partner cell (Grenier, 1992b; Kolenbrander and London,
1993). Further, the co-aggregation between A. israelii and C. gingivalis is
inhibited by SA, GalNAc and GlcNAc (Kagermeier et al., 1984). A GlcNAc
residue is thought to be a receptor for the attachment of T. denticola to
epithelial cells, fibroblasts and erythrocytes (Weinberg and Holt, 1988;
Grenier, 1991; Milcx and Keulers, 1992; Keulers et al., 1993). Interactions
between a GalNAc residue and a bacterial cell surface lectin appear to be
involved in the adhesion of Prevotella loeschei to both prokaryotic and
eukaryotic cells (London and Allen, 1989), and the co-aggregation between
Streptococcus sanguis 34 and Actinomyces viscosus T14V appears to be
dependent on a lectm-GalNAc association (Mclntre, 1985). The aggregation of
the plaque bacterium Eikenella corrodens with salivary glycoprotein is thought
to involve a bacterial cell surface adhesin that interacts with a complementary
GalNAc sugar receptor (Ebisu et al., 1992). Moreover, salivary glycoprotein is
thought to play an important role in the accumulation of dental plaque but its
aggregation with P. gingivalis does not involve GalNAc (Ebisu et al., 1992).
Several studies suggest that the binding of P. gingivalis to certain oral
bacteria and host components involves non-lectin type adhesin(s) (Olcuda et al.,
1986; Nagata et al., 1990; Bourgeau and Mayrand, 1990; Kalfas et al., 1991).
Unlike bacterial lectins, these adhesin(s) are not inhibited by sugars but are
inhibited by L-lysine or L-arginine (Olcuda et al., 1986; Bourgeau and
Mayrand, 1990; Nagata et al., 1990). Studies indicate that L-arginine can
inhibit the aggregation of Actinomyces spp. that is mediated by the OMV of P.
gingivalis, and the specific co-aggregation between S. mitis and P. gingivalis
25. 14
(Nagata et al., 1990; Bourgeau and Mayrand, 1990). L-arginine can also
inhibit the trypsin-like protease and haemagglutination activity from P.
gingivalis (NisMkata et al., 1989).
1.3 The Degradation of Host Immune Components and
Extracellular Matrix Molecules
The extracellular matrix (ECM) is made up of collagens and glycoconjugates
such as proteoglycans, glycosaminoglycans and glycoproteins. Collagens are
thought to be the main constituents of the connective tissue matrix, whüe
proteoglycans and the glycosaminoglycan hyaluronic acid are supposedly
present in the intercellular material of the epithelium. The subepithelial
basement membranes are specialised extracellular matrices and contain
collagen, chondroitin sulphate and glycoproteins such as lanainin and
fibronectin. (Uitto, 1991; Couchman and Woods, 1993). In periodontal
diseases, several factors may interfere with ECM interaction. Bacterial
enzymes and toxins may directly act on epithelial cells, resulting in degradation
of host cell surface and adhesion molecules. The host cells may react with
increased proliferation, production of inflammatory mediators and extracellular
hydrolytic enzymes, resulting in local degradation of the subepithelial basement
membrane (Uitto, 1991).
26. 15
1.3.1 Proteolytic Degradation
1.3.1.1 Ilu;31ole(%fIhroteolyde Ibazymes jBnanijP.apmypwYRKc
Proteolytic enzymes are produced by a number of microbial pathogens and
have been implicated as pathogenicity determinants (Stephen and Peitrowsld,
1986; Mirelman, 1988; Hase and Finlcelstein, 1993). It has been suggested that
the secretion of proteases by P. gingivalis may have important roles in the
degradation of host immune system and ECM components, and the generation
of oligopeptides or amino-acids for bacterial growth (Schenken, 1986; Shah and
Gharbia, 1989; Holt and Bramanti, 1991; Madden et al., 1992; Kato et al.,
1992). A secreted protease has been shown to lyse erythrocytes (Shah and
Gharbia, 1989), and P. gingivalis ean degrade IgAl, IgA2, IgG and the C3
component of complement (Kdian, 1981; Schenlcen, 1986). The inhibition of
the proteolytic degradation of IgG and C3 by P. gingivalis enhances the
phagocytosis of P. gingivalis suggesting that protease(s) contribute to
phagocytosis resistance (Cutler et al., 1993). Scott et al., 1993 have purified a
70 IcDa membrane bound thiol-protease from P. gingivalis that is able to render
fibrinogen non-clottable and suggest this enzyme is the one of the most potent
fibrases described to date. It is proposed that the fibrinolytic activity of P.
gingivalis may serve by degrading the fibrinous matrix within periodontal
lesions, allowing the baeteria to enter underlying connective tissue (Holt and
Bramanti, 1991; Lantz et al., 1991).
Trypsin-lilce activity is suggested to cause morphological changes in gingival
fibroblasts and polymorphonuclear leukocytes (PMN) (Morioka et al., 1993;
Sundqvist, 1993). It is also thought that trypsin-lilce activity of P. gingivalis
decreases the expression of complement (CRl) and IgG (FcyRII and FcyRIII)
reeeptors on PMNs. The decrease of CRl may impair the attachment of C3b-
27. 16
opsonised bacteria to PMNs. The decreased IgG receptor expression on PMNs
may result in reduced antibody-dependent cell cytotoxicity and phagocytosis of
IgG-coated bacteria (Tai et al., 1993). Sinee the PMN is thought to be
important in the maintenance of health in periodontal tissues, the impairment of
this immune eell by the trypsin-like protease activity from P. gingivalis may
allow the proliferation of this microorganism and other plaque bacteria within
periodontal tissue (Lambster and Novak, 1992).
The trypsin-lilce protease activity of P. gingivalis appears to be related to
virulence. Studies show that trypsin-like activity of the invasive strain W50 is
more than 3-fold higher than that of the avirulent mutant W50/BE1. Further,
the virulence of P. gingivalis W50 in the mouse lesion model is reduced under
haemin limitation, which is associated with a 3-fold reduction in trypsin-lilce
activity (Sundqvist, 1993). Recently, a gene from P. gingivalis that encodes
trypsin-lilce protease activity iprtT) has been isolated, characterised and the
deduced amino-aeid sequence eorrresponds to a 54 IcDa protein (Otogoto and
Kuramitsu, 1993). Two proteins (Mj. 120,000 and 150,000) that degrade
fibrinogen have been isolated from P. gingivalis and these enzymes appear to
have trypsin-lilce specificity (Lantz et al., 1991).
Collagenolytic activity is a characteristic feature of virulent P. gingivalis
strains and has been implicated in the degradation of collagen within underlying
connective tissue (Holt and Bramanti, 1991). A gene encoding collagenase
activity has been isolated from P. gingivalis and encodes a polypeptide with a
predicted molecular weight of 35 IcDa. The active enzyme, PrtC, appears to
behave as a dimer following gel filtration chromatography. The purified protein
degrades fibrillar type I collagen but not the synthetic collagenase substrate 4-
phenylazobenzyloxycarbonyl-Pro-Leu-D-Arg (Kato et al., 1992). Sojar et al.,
1993 have purified a collagenase that migrates as a 50 IcDa major band and a 60
28. 17
kDa minor band on SDS-PAGE. This enzyme hydrolysed type I collagen from
rat skin, rat plasma kininogen and transferrin. The authors also suggest that this
collagenase has a specificity for the Pro-X-Gly sequence found in several
proteins, including collagen (Sojar et al., 1993). Recently Bedi and Williams,
1994 demonstrated that a 55 kDa trypsin-like protease from P. gingivalis is
capable of degrading type I, III and IV collagen. This enzyme was also
reported to hydrolyse the C3 component of complement, fibrinogen,
fibronectin, al-antitrypsin, a2-macroglobulin, apotransferrin and human serum
albumin. These activities may suggest that this protein is involved in the
degradation of serum proteins, subepithelial basement membrane and
underlying connective tissue, which may be a potent virulence mechanism of P.
gingivalis (Bedi and Williams, 1994).
1.3.1.2 The Acüvaüomof Host Proteases by P. giMgiWis
Host matrix metalloprotemases (MMP) are a family of nine or more highly
homologous endopeptidases that cleave many components from ECM, and play
a role in the metabolic degradation of the ECM in health and disease (Birkedal-
Hansen et al., 1993). MMPs are thought to be stored in the ECM and may be
released during periodontal tissue damage (Uitto, 1991; Birkedal-Hansen et al.,
1993). The expression of MMPs can be stimulated in a variety of cultured cells
derived from human periodontal tissues, including PMNs, fibroblasts,
macrophages and kératinocytes (Birkedal-Hansen et al., 1993). MMP activity
may be stimulated either directly by microbial products or indirectly by
inflammatory mediators or cytokines generated in response to oral
microorganisms (Birkedal-Hansen, 1993).
29. 18
The fimbriae of P. gingivalis can induce the expression of IL-1 and TNFa in
mouse macrophages (Hanazawa et al., 1991; Murakami et al., 1993). Also, the
LFS of P. gingivalis can induce IL-1 production in human gingival fibroblasts
(Sisney-Durrant and Hopps, 1991). In response to IL-1 and TNFa, the host’s
fibroblasts, kératinocytes, macrophages, PMNs and endothelial cells may in
turn upregulate the synthesis and release of MMPs that degrade host tissue
(Birkedal-Hansen, 1993). Proteases of P. gingivalis may be capable of
converting mucosal kératinocytes and fibroblasts to destructive cellular
phenotypes by inducing expression of, or mediating activation of MMPs
(Birkedal-Hansen et al., 1984; Uitto et al., 1989; Birkedal-Hansen et al, 1993).
Therefore, the proteases, fimbriae and LPS of P. gingivalis may contribute to
periodontal tissue damage by activating host MMPs.
1.3.2 The Breakdown of Glycosaminoglycans by Exoglycosidases
Recent advances in glycoconjugate research have highlighted the significanee
of oligosaccharide structures in mammalian cellular interactions, protein
conformation, host-parasite recognition and antibody function (Rademacher et
al., 1988; Karlson, 1989; Feizi, 1991 & 1993; Hakomori, 1993; Oppdenaldcer
et al., 1993). The oligosaccharides linlced to glycolipids and glycoproteins are
generally branched struetures, where two to four monosaccharides can be
attached to a single monosaccharide within the chain (International Union of
Biochemistry (lUB) recommendations, 1976; Kornfeld and Komfeld, 1980).
This property distinguishes them from other glycoconjugates such as
peptidoglycans, proteoglycans and glycosaminoglycans (GAG), where
polypeptides are linked to linear carbohydrate chains composed of variable
numbers of specific repeating disaccharide units (lUB recommendations, 1980).
30. 19
Glycosaminoglycans (GAG) are present in mammalian connective tissue
(Roden, 1980; Rahemtulla, 1992), cell surfaces and the extracellular matrix
(Couchman and Woods, 1993; Yanagishita, 1993). GAGs are integral parts of
the proteoglycans (Alberts et a l, 1980; Roden, 1980; Couchman and Woods,
1993) and interact with a wide range of biolological molecules, including
fibronectins (Ruoslahti, 1988; Hynes, 1990), protease inhibitors (Bourin and
Lindahl, 1993; Yanagishita, 1993), blood coagulation factors (Bourin and
Lindahl, 1993), cytokines and growth factors (Wright et a l, 1992; Tanaka et
al, 1993). It is suggested that the GAGs are widely distribute in periodontal
tissue (Table 1.1) (Rahemtulla, 1992).
Table 1.1 Glycosaminoglycans within Periodontal Tissue
PERIODONTAL TISSUE
GLYCœiAMINOGLYCAN
(GAG)
Gingival
Fibroblasts
Gin^val
Epithelium
Periodontal
Ligament
Alveolar
Bone
Hyaluronic acid
4- 4- 4- 4-
Chondroitin sulphate
4- 4- 4- 4-
Dermatan sulphate
4- 4- ■+- -
Keratan sulphate
- - - 4-
Table 1.1 The distribution of glycosaminoglycans in periodontal tissue.
GAGs are thought to be involved in the overall integrity of gingival tissue,
where as protein-linked aggregates they interact with collagen and other cell
adhesive molecules to form a sieve-like, three-dimensional network (Bartold et
al. 1982; Uitto, 1991; Couchman and Woods, 1993). Hyaluronic acid may
represent between 30 and 40% of the total GAG in gingival tissue, with the
remainder being dermatan and chondroitin sulphate (Embery et a l, 1979). The
composition of GAGs in periodontal tissue is thought to alter during
31. 20
periodontitis and may fluctuate with disease severity (Kirldiam et al., 1992;
Rahemtulla, 1992; Shibutani et al., 1993). It has been reported that the ratio of
glucuronic acid to hexosamine is the same in normal and diseased human
gingivae, with the polysaccharide components of GAGs being significantly
reduced in the diseased tissue (Rahemtulla, 1992). It has been suggested that
direct enzyme attack on GAGs by bacterial plaque, or indirect attack by
lysosomal enzymes as part of an inflammatory response may lead to an
imbalance in the structure of periodontal tissue (Tipler and Embery, 1985). It is
possible that such disruption of GAGs is due to brealcdown of GAGs by
exoglycosidases, which in turn could have pronounced effects on the functional
integrity of periodontal tissue. Interestingly, the exoglycosidase P-N-Acetyl-
hexosaminidase participates in the step-wise degradation of the
glycosaminoglycans hyaluronic acid (Bach and Geiger, 1978; Roden, 1980;
Kresse and Glossl, 1987), keratan sulphate (Yutaka et al., 1982; Kresse and
Glossl, 1987), chondroitin sulphate and dermatan sulphate (Kresse and Glossl,
1987) (Figure 1.2).
32. 21
Figure 1.2 Structure and Sequential Hydrolysis of Glycosaminoglycan
Glycosidic Linkages by Exoglycosidases
coo
O alN AcYo, OalNAcOlcA
(%
B AA B
(i) Structure and sequential
degradation of hyaluronic acid.
A= P-glucuronidase
B=P-N-Acetyl-hexosaminidase
ooo
GalNAcGalNAc^GlcA
DB
O-SOj poo
Q y—
GalNAc
OhOH
at
D
O t-O -S O ; O t-O -aO ) ÇH20H qt-o-8 0 .
00
CHj
B D
(ii) Structure and sequential
degradation of dermatan sulphate.
A= p-glucuronidase
B and F=N-Acetyl-galactosamine-4-
sulphate sulphatase
C and H=P-N-Acetyl-hexosaminidase
D=iduronate-2-sulphate sulphatase
E=a-L-iduronidase
(iii) Structure and sequential
degradation of chondroitin-4/6-sulphate.
A and D=P-glucuronidase
B=N-Acetyl-galactosamine-6-sulphate
sulphatase
C and F= P-N-Acetyl-hexosaminidase
E=N-Acetyl-galactosamine-4-sulphate
sulphatase
(iv) Structure and sequential
degradation of keratan sulphate.
A= N-Acetyl-galactosamine-6-sulphate
sulphatase
B and E=P-galactosidase
C and F= N-Acetyl-glucosamine-6-
sulphate sulphatase
D and G= P-N-Acetyl-hexosaminidase
Figure 1.2 The sequential degradation of glycosaminoglycans by
lysosomal exoglycosidases. The exoglycosidases recognise
monosaccharides at the outer non-reducing end of the oligosaccharide and
release them in turn (Modified from Kresse and Glossl, 1987)
33. 22
1.4 p-N-Acetyl-Hexosaminidases and
P-N-Acetyl-Glncosaminidases
These enzyme activities have been reported in organisms from microbes to
higher animals (Gibson and Fullmer, 1969; Neufield, 1989; Esaiassen et al.,
1991; Conoi et at., 1992; Cesari gf oZ., 1992). P-N-Acetyl-hexosaminidases can
hydrolyse terminal non-reducing p-linked NAcGlc and NAcGal, while exo-P-
N-Acetyl-glucosaminidases are described as enzymes that hydrolyse terminal
non-reducing NAcGlc (Cabezas, 1989). Since P-N-Acetyl-hexosaminidases (EC
3.2.1.52) can also be termed as exo-P-N-Acetyl-glucosaminidases (EC
3.2.1.30) under enzyme nomenclature, both names have been used for the same
exoglycosidase activity (Cabezas, 1989). Exo-P-N-Acetyl-glucosaminidases are
also occasionally termed chitobiases (formerly EC 3.2.1.29 now EC 3.2.1.30)
due to their ability to release terminal non-reducing GlcNAc residues from the
homopolymer, chitin (lUB recomendations, 1978; Cabib, 1987; Flach et at.,
1992).
1.4.1 Specificity
Exoglycosidases usually show a very high degree of specificity for a
particular monosaccharide, which is controlled by at least two factors. Firstly,
the glycon specificity is directed towards the terminal sugar moiety, its
anomeric configuration (a/p) and its sterioisomericity (D/L) (Kobata, 1979;
Dwek et al, 1993). Secondly, the aglycon (the rest of the oligosaccharide)
influences the hydrolysis of a monosaccharide and in certain cases affects the
specificity of the enzyme (Kobata, 1979). For example, the P-N-Acetyl-
hexosaminidase isolated from Streptococcus pneumoniae culture filtrate
(Glasgow et al., 1977), can cleave terminal GlcNAc/GalNAcp 1-2Man linkages
34. 23
from N-linlced oligosaccharides only if the mannose is not substituted at C-6
(Furulcawa and Kobata, 1993) (Figure 1.3). Its reported activity against O-
linked oligosaccharides suggests that terminal GlcNAcpl-3Gal and GlcNAcpi-
6Gal can be hydrolysed (Yamashita et al., 1981). The P-N-Acetyl-
hexosaminidase from Jack Bean has a broad specificity (Li and Li, 1970), and
can cleave terminal GlcNAc/GalNAcP 1-2, 3, 4 or 6Man linkages (Welply,
1989; Dwek et a l, 1993). One study suggests that Jack Bean P-N-Acetyl-
hexosaminidase may not be able to hydrolyse the GlcNAcP l-4GlcNAc
oligosaccharides derived from chitin (Iwamoto et al., 1993).
The p-N-Acetyl-hexosaminidases from human lysosomes are extensively
characterised since defects result in lysosomal storage of their substrates and
neurodegenerative disease (Cantz and Kresse, 1974; von Figura and Hasilk,
1986; Neufield, 1989). The p-N-Acetyl-hexosaminidases from human
lysosomes are formed by dimérisation of two different polypeptide chains, a
and p. In normal human tissues, at least two different isoenzyme forms of P-N-
Acetyl-hexosaminidases occur, namely A (aP) and B (PP). The A form has a
broad substrate specificity and can remove terminal non-reducing p-linlced
NAcGal or NAcGlc from glycoconjugates that occur in human cells. The B
isoenzyme has a similar specificity with the key exception that it does not
hydrolyse the glycolipid GM% ganglioside or the synthetic substrate MU-
G1cNAc-6-S04 (Price and Dance, 1972; Kytzia and Sandhoff, 1985; Neufield,
1989).
P-N-Acetyl-hexosaminidases and exo-P-N-Acetyl-glucosannnidases can
hydrolyse terminal NAcGlc monosaccharides, while endo-P-N-Acetyl-
glucosaminidases usually cleave internal NAcGlc linkages within
oligosaccharides (Figure 1.3) (Kobata, 1979; Cabezas, 1989; Furulcawa and
35. Kobata, 1993). The exoglycosidases usually differ from the endoglycosidases in
terms of substrate specificity (Kobata, 1979; Valisena et al., 1982; Morinaga et
al., 1983; Greber et al, 1989; Sugai et al., 1989; Furulcawa and Kobata, 1993),
and amino-acid sequence similarity (Henrissat and Bairoch, 1993). Many endo-
P-N-Acetyl-glucosaminidases that have been described do not contain exo-P-N-
Acetyl-glucosaminidase activity (Valisena et al., 1982; Morinaga et al., 1983;
Greber et al, 1989; Sugai et al., 1989; Furulcawa and Kobata, 1993), however,
a Neisseria gonorrhoeae enzyme that contains both exo- and endo-P-N-Acetyl-
glucosaminidase activity has been reported (Gubish et al., 1982).
1.4.2 Possible Roles in Glycocopjngate Degradation
1.4.2 .1 The ModiHcation of Host Extracellular Matrix
Glycosaminoglycans
The invasive parasites Trichinella spiralis and Entamoeba histolytica contain
high levels of P-N-Acetyl-hexosaminidase (Lundblad et al., 1981; Rhoads,
1985). These microorganisms penetrate the intestinal mucosa, invade and
damage other host tissue (Ravidin, 1986; Prescot et al., 1990). Mice infected
with T. spiralis are reported to contain 40-50% less muscle-parasite when
immunised with the P-N-Acetyl-hexosaminidase from T. spiralis, suggesting
this enzyme is important for pathogenicity (Rhoads, 1988). The p-N-Acetyl-
hexosaminidase and several other hydrolases from E. histolytica are suggested
to be involved in the pathogenesis of invasive amoebiasis (Ravidin, 1986;
Mirelman, 1988). Lundblad et al., 1981 purified the secreted P-N-Acetyl-
hexosaminidase from E. histolytica strains and postulated that this enzyme has a
role in erythrocyte brealcdown or intestinal epithelial damage. In degradation
experiments. Worries et al., 1983 showed that the p-N-Acetyl-hexosaminidase
36. 25
from E. histolytica could degrade oligosaccharides from hyaluronic acid, which
may suggest that this enzyme participates in the degradation of host submucosa
ECM.
P. gingivalis is reported to degrade the GAG chondroitin sulphate (Holt and
Bramanti, 1991), and express an extracellular N-Acetyl-galactosamine-4-
sulphatase capable of releasing sulphate from gingival GAG (Slomiany et al.,
1993). P. gingivalis has several exoglycosidase activities that have the potential
to hydrolyse a number of host oligosaccharides. These enzyme activities
include P-galactosidase, neuraminidase and p-N-Acetyl-hexosaminidase
(Laughon et al., 1982; van WinlceUioff et al., 1985; Nash, 1987; Syed et al.,
1988; Minhas and Greenman, 1989; Moncla et al., 1990; A. Wallace, personal
communication, 1993).
The increased levels of P-N-Acetyl-hexosaminidase activity in gingival
crevicular fluid from periodontal disease patients and the serum of subjects with
cancer may implicate this enzyme activity in tissue damage (Dobrossy et al.,
1980; Tucker et al., 1980; Chatterjee et al., 1982; Beighton et a l, 1992). In
cancer, it is accepted that the joint action of host hydrolytic enzymes degrade
the ECM and contribute to tissue breakdown, whereas in periodontal disease,
tissue damage may be due to both bacterial and host enzymes (Nicolson, 1982;
Uitto, 1991; Lambster and Novak, 1992). Studies in cancer research have
demonstrated that highly metastatic murine leukemic and human ovarian
carcinoma cells release substantial amounts of glycosidases, with P-N-Acetyl-
hexosaminidase showing high secretory activity (Bosman and Bernacki, 1970;
Niedbata et al., 1987). Moreover, both tumour cell- and P-N-Acetyl-
hexosaminidase-mediated ECM degradation can be inhibited with sugar analogs
loiown to be competitive inhibitors of P-N-Acetyl-hexosaminidase (Niedbata et
al, 1987; Woynarowska et al., 1989 & 1992). Although the site of action for p
37. 26
-N-Acetyl-hexosaminidase in ECM is unclear, it can be postulated that this
enzyme may act on macromolecules such as the glycosaminoglycans. Also, the
treatment of basement membrane material with P-N-Acetyl-hexosaminidase
significantly increases its sensitivity to trypsin degradation (Jensen and Lendet,
1986). Therefore, it could be implied that p-N-Acetyl-hexosaminidase may
unmask new proteolytic sites of action allowing more efficient brealcdown of
the ECM (Woynarowska et al., 1989). If, in a similar manner, the P-N-Acetyl-
hexosaminidase and trypsin-like protease of P. gingivalis can degrade oral
basement membrane, this could farther implicate these enzyme activities as
virulence factors of P. gingivalis. With this in mind, it is tempting to speculate
about the increased level of p-N-Acetyl-hexosaniinidase activity in gingival
crevicular fluid during periodontal disease. For example, this activity could be
due to the action of bacterial exoglycosidases and/or host lysosomal enzymes as
part of an inflammatory response. Moreover, this enzyme activity from P.
gingivalis may play a role in ECM degradation during chronic adult periodontal
disease.
1.4.2.2 The Degradaüom of (glycoproteins and Glycolipids
The oligosaccharides of host cell surface glycosphingolipids and
glycoproteins have roles as molecular attachment sites for a number of
pathogenic bacteria (Karlson, 1989). Moreover, the action of glycosidases on
cell surface structures may influence bacterial colonisation or attachment
(Hoskins, 1991). For example, the attachment of T. denticola to laminin and
fibrinogen is significantly decreased when the bacteria are treated with a
mixture of glycosidases (Haapasalo et al., 1991). Treatment of epithelial cells
with neuraminidase increases the binding of P. gingivalis but decreases the
attachment of Streptococci to these host cells (Socransky and Haffajee, 1991).
Therefore, it could be suggested that the extracellular neuraminidase activity of
38. 27
P. gingivalis may eliminate the binding of Streptococci to SA residues on
epithelial cells, thereby facilitating the attachment of P. gingivalis to these
cells. Since a number of bacterial adhesins recognise complementary
GlcNAc/GalNAc residues (Section 1.2.2), a decrease of GalNAc/GlcNAc
lectin-receptors on the surface of host cells, bacteria and salivary glycoprotein
would directly hinder the adhesive ability of certain oral bacteria in the
periodontal microbiota. If the extracellular P-N-Acetyl-hexosaniinidase from P.
gingivalis can deplete GalNAc/GlcNAc lectin-receptors this enzyme may
modulate bacterial adhesion.
p-N-Acetyl-hexosaminidase activity is also assumed to play a role in the
sequential degradation of digestive tract mucin oligosaccharides (Prizont, 1982;
Hoskins, 1991; Boureau et al., 1993). Mucins are glycoproteins that form a gel
covering the mucosal surface, which protects the delicate epithelial cells against
the extracellular environment and selects substances for binding and uptake by
these epithelia (Süberberg, 1989; Strous and Dekker, 1992). Mucins contain
oligosaccharide structures with large quantities of GalNAc, GlcNAc, Gal, Fuc,
and SA, and carbohydrate may be roughly 50 % of the dry weight (Strous and
Dekker, 1992). Since non-covalent interactions between carbohydrate moitiés
on mucins are thought to be important in the formation mucin-clusters and gel
like properties (Strous and Deldcer, 1992), the degradation of oligosaccharide
structures on mucins may modify the effectiveness of epithelial hmction and
protection (Hoskins, 1991). Moreover, Eposito et al., 1983 has shown that in
vivo treatment of rat mucosa with a mixture of the exoglycolytic enzymes P-N-
Acetyl-hexosaminidase, neuraminidase, P-galactosidase, sulphatase, a-
mannosidase, P-glucosidase and P-glucuronidase causes the mucosa-to-serosa
permeability to increase dramatically.
39. 28
It is known that certain faecal bacteria can degrade mucin oligosaccharides
(Salyers et al., 1977; Robertson and Stanley, 1982; Hoskins et al., 1985;
Ruseler van Embden et al., 1989). The bacterial degradation of
oligosaccharides on mucin glycoproteins and intestinal glycosphingolipids
appears to require extracellular neuraminidase, a-glycosidases, p-galactosidase
and p-N-Acetyl-hexosaminidase activities (Hoskins and Boulding, 1981;
Hoskins et al., 1985; Larson et al., 1988; Falk et al., 1990; Hoskins, 1991). hi
Bacteriodes fragilis, the production of exoglycosidases with the potential to
degrade mucin is inversely related to growth rate when mucin is provided as
the sole carbon and nitrogen source (MacFarlane and Gibson, 1991). Mucin has
been shown to induce the production of the both extracellular and cell-bound
neuraminidase, a-glucosidase, p-galactosidase and p-N-Acetyl-hexosaminidase
activities of feacal bacteria (MacFarlane et al., 1989). Moreover, when
subgingival plaque bacteria are grown in BM medium (medium that supports
the growth of black-pigmented anaerobes see section 2.1.1), with added mucin,
these glycosidase activities and P-glucuronidase increase significantly (Beighton
et a/., 1988). Since mucin increases the production of exoglycosidases in the
bacteria of subgingival plaque, it is possible that these enzymes may degrade
the oligosaccharides of mucin structures within the gingival epithelium and
salivary pellicle (Dabelsteen et al., 1991). Additionally, in host cell plasma
membranes, the oligosaccharides that are linlced to glycoproteins and
glycosphingolipids are thought to be important for host cell-to-cell adhesion and
interaction (Hakomori, 1993). Consequently, the combined action of
glycosidases degrading these oligosaccharides could have important effects on
host cell-to-cell interactions.
The release of carbohydrate molecules might also enhance the growth of
certain bacterial species. This hypothesis is based on the observation that a lack
of glycosidic enzymes in Clostridium difficile may be correlated with its
40. 29
nutritional supression and inability to compete for sugars in the normal colonic
microflora. Moreover, free sugars added to a normal colonic microflora in
continuous culture enhance the growth of C. difficile (Wilson and Perini,
1988). Further, a faecal strain of Ruminococcus gnavus which has a-
galactosidase activity but lacks P-N-Acetyl-hexosaminidase activity cannot
degrade the backbone of blood group B oligosaccharides on salivary
glycoprotein. When this strain is grown in symbiotic association with a strain
that produces P-N-Acetyl-hexosamdnidase, extensive degradation occurs and
bacterial growth is enhanced (Hosldns et al., 1985; Hoskins, 1991). If
comparable circumstances exist in the periodontal microflora, then it can be
postulated that the action of p-N-Acetyl-hexosamdnidase on salivary
glycoprotein and cell surface glycoconjugates may provide growth factors that
nutritionally enhance certain saccharolytic members of bacterial plaque.
1.4.2.3 Hydrolysis of Asparagine-Linked Oligosaccharides on
Hnman IgG
The two CH2 domains in the Fc region of human IgG are thought to be held
apart by two asparagine-linlced oligosaccharides of the general structure in
Figure 1.3. Some IgG molecules contain al-3 arms and al-6 arms that lack the
terminal SA-Gal and terminate in GlcNAc. The al-3 arms from each sugar are
thought to provide a bridge between the two CH2 domains, whereas the al-6
arms are thought to be directed towards the surface of the CH2 domains
(Rademacher, 1988; Roitt, 1991). The al-3 and al-6 arms can be completely
removed by the sequential enzymatic action of neuraminidase, P-galactosidase
and P-N-Acetyl-hexosaminidase (Figure 1.3) (Koide et al., 1977).
41. 30
Figure 1.3 The Structure and Enzymatic Degradation of the Desialylated
Oligosaccharides of Human IgG
S. pneumoniae Jack Bean or S. pneumoniae
fi-galactosidase /J-N-Acetyl-hexosaminidase
X X
±Gaip 1—4GlcNAcP 1—2M anal +Fucal
Jack Bean
P-N-Acetyl-hexosaminidase^ 5 5
±GlcNAcP 1—4ManP 1—4GlcNAcP 1—4G 1cN A c-asp
3 X
/ S. pneumoniae
±Gaip 1—4GIcNAcP 1—2M anal endo-p-N-Acetyl-glucosandnidase
% %
S. pneumoniae Jack Bean or S. pneumoniae
p-galactosidase P-N-Acetyl-hexosandnidase
monosaccharide sequence
►
Figure 1.3 The action of P-N-Acetyl-hexosaminidase, P-
galactosidase and endo-P-N-Acetyl-glucosaminidase on desialylated
(neuraminidase-treated) asparagine-linked sugar chains of human
IgG. The exoglycosidases cleave each glycosidic linkage down the
monosaccharide sequence begining at the outer non-reducing end.
The basic structure required for endoglycosidic hydrolysis by the
endo-P-N-Acetyl-glucosaminidase is shown in bold. The endo-P-N-
Acetyl-glucosaminidase hydrolyses the internal glycosidic linkage
most efficiently when the al-3-linked mannose residue is not
substituted at C-2 (Modified from Kiode et at., 1977; Mizuochi et
at., 1982; Kobata era/., 1989; Furukawa and Kobata, 1993).
If the neuraminidase, p-galactosidase and P-N-Acetyl-hexosaminidase
activities from P. gingivalis can sequentially remove the al-3 and al-6 arms of
each oligosaccharide in vivo, this may aid modification of IgG interactions.
This idea is based on the importance of these oligosaccharides in antibody-
monocyte interaction and the molecular stability of IgG (Leatherbarrow et al.,
42. 31
1985; Rademacher, 1988; Oppdenaldcer et al., 1993). Moreover, the treatment
of intact IgG with neuraminidase and P-galactosidase has no effect on rosette
formation or antibody-dependent cell-mediated cytotoxicity. However,
additional manipulation with p-N-Acetyl-hexosaminidase and endo-P-N-Acetyl-
glucosaminidase from S. pneumoniae decreased these immunological reactions
(Koide et al., 1977).
1.4.2.4 Autolysins that Degrade Pepddoglycan
Bacterial peptidoglycan hydrolases that can degrade their own cell walls are
refered to as autolysins (Ghuysen et al., 1966). Autolysins can be classified as
N-Acetyl-muramidases, endopeptidases, transglycosidases, N-Acetyl-muramyl-
L-alanine-amidases and P-N-Acetyl-glucosaminidases (Ghuysen et al., 1966).
Autolysins may be required for several important cellular functions, including
cell wall growth, turnover and splitting of the septum for cell separation (Holtje
and Tuomanen, 1991). Some autolysins may be important for bacterial
pathogenicity and may cause release of highly inflammatory cell wall
components or may lyse a portion of the cell that results in the liberation of
toxins (Berry et al., 1989; Holtje and Tuomanen, 1991; Wuenscher et al.,
1993)
Endo-P-N-Acetyl-glucosaminidase mutants of Bacillus subtilis 168 appear to
form long chains of unseparated cells. Therefore, this endoglycosidase activity
has been suggested to participate in cell growth, division and shape formation
(Fein and Rogers, 1976). However, an endo-P-N-Acetyl-glucosaminidase gene
from B. subtilis AC327 was disrupted using site specific mutagenesis and the
mutant strain appeared normal in growth and morphology (Rashid et al., 1993).
It was proposed that the endo-p-N-Acetyl-glucosaminidase together with
43. 32
amidase may be important for B. subtilis motility rather than growth and
morphology (Rashid et al., 1993). Endo-P-N-Acetyl-glucosaminidases may
play a significant role in bacterial pathogenicity (Berry et al., 1989; Valisena et
al., 1991). For example, an endo-P-N-Acetyl-glucosaminidase from
Staphylococcus aureus inhibits the response of human lymphocytes to mitogens
and interferes with the production of antibodies in mice (Valisena et al., 1991).
This enzyme is also suggested to share common epitopes with microbial and
mammalian exoglycosidic P-N-Acetyl-hexosaminidases (Guardati et al., 1993).
Bacterial exo- and endo-P-N-Acetyl-glucosaminidases are reported to
hydrolyse peptidoglycan and they may play a role as endogenous prokaryotic
hydrolases that modify peptidoglycan during cell growth (Kawagishi et al.,
1980; Gubish e/fl/., 1982; Chapman and Perldns, 1983). However, in bacterial
peptidoglycan modification, exo-P-N-Acetyl-glucosaminidase activity has an
uncertain physiological role and some authors are sceptical of its dhect
involvement in peptidoglycan turnover or metabolism (Doyle and Koch, 1987).
This may be due to the observation that exo-P-N-Acetyl-glucosaminidase
mutants of Bacillus subtilis B and Escherichia coli K12 appear normal in
growth, division and morphology (Ortiz, 1974; Yem and Wu, 1976). Also, it
has been reported that B. subtilis cells do not reincorporate pre-radiolabelled
cell wall GlcNAc into their 'new' peptidoglycan during growth, which could
suggest a more indirect role for exo-P-N-Acetyl-glucosanimidase enzymes in
wall metabolism or turnover (Doyle and Koch, 1987). Further studies are
needed to examine the role of these exoenzymes in bacterial cell wall
metabolism.
44. 33
1.4.3 p-N-Acetyl-Hexosaminidases in Glycobiology
The deglycosylation enzymes are powerful tools in the analyses of
oligosaccharide structure and function (Maley et al., 1989; Welply, 1989;
Dwek et a l, 1993; Furukawa and Kobata, 1993). A variety of specific
glycosidases are generally used and enzymatic deglycosylation is carried out by
sequential exoglycosidase digestion or by endoglycosidases (Nageswara and
Bahl, 1987; Welply, 1989; Dwek et a l, 1993; Furukawa and Kobata, 1993).
The detennination of oligosaccharide structures linked to the glycoconjugates
shown in Table 1.2 involved the use of P-N-Acetyl-hexosaminidase cleavage. P
-N-Acetyl-hexosaminidase cleavage has also contributed to the analysis of the
oligosaccharides linked to fibrinogen (Townsend et al., 1982), intracellular
glycoproteins (Hanover et al., 1987; Holt et al., 1987), human chorionic
gonadotropin (Goverman et al., 1983), plus the epidermal growth factor
(Cummings et al., 1985) and acetyl-choline receptors (Herron and Schimerlick,
1983). The wide use of P-N-Acetyl-hexosaminidases in mixed glycosidase
digestion procedures that analyse oligosaccharide structure and function
demonstrates that these enzymes are valuable tools for glycobiologists.
Therefore, it is lUcely that commercially available and novel P-N-Acetyl-
hexosaminidases will play an integral role in the future of carbohydrate
research. Together, P-N-Acetyl-hexosaminidases and other glycosidases may
be pivotal in the identification and commercialisation of new developments in
glycobiology.
45. 34
Table 1.2 Some Selected Glycoproteins and Glycolipids
Glycoconjugate Function Role of CHO References
Immunoglobulin G Host defence Fc interaction
Stability
Protease resistance
Mizuochi et al., 1982
Taniguichi et al., 1985
Rademacher et al., 1986
Rademacher et al., 1988
aj-acid
glycoprotein
Immunomodulator ? Jeanloz, 1972
Yoshima etal., 1981
Bouten etal., 1992
Interleukin-6 T/B cell activation ? van Snick, 19W
Parekh et al., 1992
p67 Eukaryote
translation
Regulation? Datta etal., 1989
ABH cell surface
antigens
Tissue markers
and
Cellular adhesion
Cell-cell interactions Ito et at., 1989a&b
Dabelsteen et al., 1991
Hakomori, 1993
a2 -HS-glycoprotein Bone metabolism? ? Colclasure etal., 1988
Watzlawick etal., 1992
Fetuin Development
and
Lipogenesis
? Takasaki & Kobata, 1986
Cayatte et al., 1990
Table 1.2 Selected examples of glycoconjugates where P-N-Acetyl-
hexosaminidase cleavage has been applied to determine
oligosaccharide structure or function. The oligosaccharide functions
are shown.
In summary, new advances in glycobiology have demonstrated that
oligosaccharides play a key role in interactions between pathogens and host
cells. Since oligosaccharides are also involved in many host cell fimctions and
molecular interactions, then extracellular glycosidic enzymes liberated by
pathogenic bacteria may be important in the pathogenesis of disease. Although
microbial p-N-Acetyl-hexosaminidases have been shown to hydrolyse a wide
range of carbohydrate structures, there is little information about the exact
role(s) of these enzymes from pathogenic microorganisms. However, the
46. 35
growing interest in microbial P-N-Acetyl-glucosamdnidases and P-N-Acetyl-
hexosaminidases is reflected by the cloning of the genes from Vibrio harveyi
(Jannatipour et al., 1987; Soto-Gil and Zyskind, 1989), V. vulnificus (Wortman
et al., 1986; Somerville and Colwell, 1993), V. fumissii (Bassler et a l, 1991),
V. parahaemolyticus (Zhu et a l, 1992), Serratia liquefaciens (Joshi et a l,
1988), S. marcescens (Kless et a l, 1989; Tews et a l, 1992), P. gingivalis
(Lovatt and Roberts, 1991), Dictyostelium discoideum (Graham et a l, 1988)
and Candida albicans (Cannon et a l, 1994). P-N-Acetyl-glucosaniinidases and
P-N-Acetyl-hexosanhnidases might function as virulence factors that degrade
host oligosaccharides, and play a role in the hydrolysis of peptidoglycan, acting
as autolysins. Although numerous studies have characterised and exploited
enzymes within this group of exoglycosidases, there is sthl much to loiow about
their regulation and physiological significance in the degradation of
glycoconjugates.
Lastly, the occurence of densely packed mixtures of diverse bacterial species
in dental plaque suggests that bacterial interactions play an important role in
species survival. The in vivo formation of bacterial aggregates may provide a
network for the growth and retention of selected oral bacterial species. Some
interspecies relationships may be favourable, in that one species produces
growth factors for, or facilitates the attachment of, another. Other relationships
may be antagonistic due to the competition for nutrients, or the production of
substances that inhibit the growth or attachment of a second species. For
successful colonisation of the periodontal pocket, the adhesion to host
components may be required, and the fimbriae, trypsin-lilce proteases and
haemagglutinins of P. gingivalis may serve in the attachment process. The
production of proteases and glycosidases by P. gingivalis may modulate
bacterial adhesion and damage host immune components. In addition, proteases
and glycosidases may degrade bacterial/host cell surfaces and ECM molecules
47. 36
in an effort to gain ecological advantage, perhaps by providing nutrients and/or
assisting in microbial attachment and spreading. The glycosidases produced by
P. gingivalis and the bacteria of subgingival plaque may release the
carbohydrate from human IgG, protective mucin glycoconjugates and basement
membrane glycosaminoglycans. In response to ECM damage by bacteria,
hydrolytic enzymes and LPS, the host may amplify tissue destruction by
releasing or expressing matrix-metalloproteinases that degrade the subepithelial
basement membrane. Although there is no evidence that P-N-Acetyl-
hexosaminidase activity contributes to the degradation of host ECM molecules
during periodontal disease, considerations may offer future research directions
that wül contribute to the understanding of molecular mechanisms in tissue
damage and periodontopathogenesis.
1.5 The Aims of This Thesis
Firstly, the aim of this thesis is to speculate on the function of the P-N-
Acetyl-hexosaminidase from P. gingivalis and to adopt a genetic manipulation
approach for defining the role of this enzyme. Secondly, to use this strategy to
isolate and produce structural information on the gene that encodes for P-N-
Acetyl-hexosaminidase activity. To use this information to construct a site-
directed gene replacement strategy for the generation of a isogenic mutant of P.
gingivalis lacldng P-N-Acetyl-hexosaminidase activity, which can be compared
with the wild type P.gingivalis in appropriate model systems. The last aim is
for this thesis is to provide guidance and inspiration for future work and new
investigators.
48. 37
CHAPTER 2
Materials and Methods
2.1 Bacterial Strains and Plasm ids
The bacterial strains and plasmids that were used in this study are listed in
Table 2.1 and Table 2.2 respectively.
2.1.1 Growth Conditions and Media
Porphyromonas gingivalis, Porphyromonas asaccharolytica and
Porphyromonas endondontalis strains were grown anaerobically at 37°C in
Bacteroides Medium (BM) (10 g/litre ' trypticase peptone; lOg/litre ' proteose
peptone; 5g/litre'^ yeast extract; 5g/litre ' glucose; 5g/litre ‘ NaCl; 0.7g/litre ‘
cysteine HCl; Ig/litre^ NaHCOg; 5pg/ml ' hemin; lOpg/ml ' menadione) with
the addition of 1.5% w/v agar (BBL) as required or on 7% v/v horse-blood
agar plates (Oxoid). E. coli srains were grown in Luria broth (L-broth) (25
g/litre ' peptone; 12.5g/litre ‘ NaCl; 25g/litre ‘ yeast extract) at 37°C with the
addition of 1.5% w/v agar as required. B-agar (10 g/litre ' peptone; 8g/litre '
NaCl, 1.5% w/v agar) was used where stated. For detecting P-N-Acetyl-
hexosaminidase activity, the flourogenic substrates 4-methylumbelliferyl-N-
Acetyl-p-D-glucosanunide (MUAG) and 4-methylumbelliferyl-N-Acetyl-P-D-
galactosaminide (MUAGal) were added to media at a concentration of lOOp
g/ml. Antibiotics at concentrations of lOOpg/ml ampicillin, 25pg/ml
tetracycline and 25pg/ml kanamycin were used when required. Antibiotics and
substrates were obtained from Sigma Chemical Comp. Ltd.. Bacterial cells
49. 38
were routiiüey harvested by centrifiigation in a Sorval centrifiige (3300g at 4°C
for 10 minutes (mins)) or in a bench top minifuge (13400g at room temperature
for 5 mins).
Table 2.1 Bacterial Strains
Bacterial strain Relevant characteristic Source *
E.COÜ
SURE™ recB r e d sbcCZOl uvrC
umuC::Tn5 (konO lacA
(hsdRMS) endAl gyrA96 thi
relAl supE44 F'lproAB'^
lacB lacZAMlS TnlO (tef)]
Stratagene® Cloning
Systems
JMlOl supE thil A(lac-proAB)
F'[froD36 proAB'^ lacP
Yanisch-Perron et al., 1985
DS410 minA minB ora xyl m l azi
thi
Dougan and Sherratt, 1977
P. gingivaüs
W83 Clinical specimen H. Shah"
WpH35 Clinical specimen MRC Dental Unit, London
23A3 Clinical spœimen MRC Dental Unit, London
ATCC 33277 Type strain ATCC^
P. endodontalis
ATCC 35406 Type strain ATCC
P. asacharolytica
ATCC 8503 Type strain ATCC
Table 2.1 Bacterial strains. * Addresses; ^, Eastman Dental
Hospital, London. , American Type Culture Collection, Rockville,
Maryland, USA
50. 39
Table 2.2 Bacterial Plasmids / Vectors
Plasmid/vector Relayent characteristic Construction or source
pTTQlS pUC based expression
vector (Amp' lacN)
Stark et at., 1987
M13mpl8/19 M13 cloning/sequencing
vector
Yanisch -Perron
gfo/., 1985
pNJR6 Colonic Bacteroides suicide
vector derived from the
shuttle cosmid pNJRl and
does not contain the pB8-51
region that enables
replication in Bacteroides
hosts (Kan' Cc' Em' Sm')
Shoemaker et al., 1989
PGEX-2T Vector that directs the
synthesis of foreign
polypeptides as fusions with
glutathione S-transferase
(Amp')
Smith and
Johnson, 1988
Table 2.2 Bacterial plasmids / vectors.
2.2 Transformation of Bacterial Cells
2.2.1 Production of Competent CeUs
2.2.1.1 Calcium Chloride Method
lOOpl of an E. coli overnight culture grown at 37°C in 10ml L-broth were
diluted 1:100 with 10ml of L-broth and grown to mid exponential phase
(ODgoty» 0.4). The cells were harvested (3300g at 4°C for 10 mins), washed in
10ml of lOmM NaCl, pelleted (3300g at 4°C for 5 mins) and resuspendW in
4ml ice-cold CaCfy (lOOmM). The cells were placed on ice for 30 mins and
collected by gentle centrifugation (1800g) at 4°C for 5 mins. The cell pellet was
51. resuspended in 1ml ice-cold CaCl] (lOOmM) and used immediately in
transformation,
2.2.1.2 Electrotransfbrmadon Method
lOOpl of an overnight culture grown at 37°C in 10ml L-broth were diluted
1:100 with 10ml of L-broth and grown to mid exponential phase (ODggo^O.S).
The cells were chilled on ice for 15 mins and harvested (3300g at 4°C for 10
mins). The cell pellet was washed 4 times in 10ml of nanopure water and once
in 10% v/v glycerol with centrifugation as before between the washes. The cell
pellet was then resuspended in SOpl of 10 % v/v glycerol and used immediately
in transformation.
2.2.2 Transformation with Plasmid DNA
2.2.2.1 Calcium Chloride Method
Competent cells (lOOpl) were mixed with 5-20pl of DNA (in water) and
placed on ice for 1 hour (hr). The cells were heat shocked at 42°C for 3 mins.
Immediately after heat-shocldng, SOOpl of L-broth were added and the cells
incubated for 1 hr at 37°C. The transformed cells were plated onto L-agar
plates (lOOpl per plate), which contained the appropriate antibiotic(s).
52. 41
2.2.2.2 Electrotransfbrmation Method
Competent cells (40pl) were mixed with l-2pl of DNA (in water) and
transferred to an ice-cold BIO-RAD 0.2cm gene puiser cuvette. The cell
suspension was pulsed (2.4kVcm-i, 25pF, 2000) on a BIO-RAD Gene-Pulser
apparatus. Immediately after pulsing, 1ml of ice cold SOC recovery medium
(20 g/litre'^ tryptone; 5g/litre'^ yeast extract; lOmM NaCl; 2.5mM KCl; lOmM
MgSO^; 20mM glucose) was added and the cells were incubated for Ihr at 37°
C with shaldng. The transformed cells were plated onto agar plates (lOOpl per
plate), which contained the appropriate antibiotic(s).
3 TTraauüRariaadioïkivith IkacterioidbaypeiDOSYl
Competent cells of E. coli JMlOl (lOOpl; see Section 2.2.1.1) were mixed
with DNA, incubated on ice for 1 hr and heat-shocked at 42°C for 3 mins. The
transformed cells were mixed with lOOpl of JMlOl (ODggg^O.S) and then 3ml
of molten B-agar (held at 45°C and containing 20pl lOOmM IPTG, SOpl 2%
w/v X-gal in dimethylformamide) were added. The suspension was immediately
mixed and poured onto a B-agar plate, rocked to disperse and once set
incubated at 37°C overnight.
53. 42
2.3 Procedures for DNA Extraction
DNA extraction protocols used the following solutions:
solution I : 50mM glucose
25mM Tris-HCl pH 8.0
lOmM EDTA
5mg/ml lysozyme
solution II 0.2M NaOH
35mM sodium dodecyl sulfate (SDS)
solution III : 5M acetate (11.5ml glacial acetic acid)
3M potassium ions (60ml 5M potassium acetate)
nanopure water (28.5ml)
2.3.1 Extraction of Chromosomal DNA
The method used to extract chromosomal DNA was based on that described
by Saito and Muira, 1963. Bacterial cells from 10ml stationary phase cultures
were washed in lOmM NaCl and resuspended in 5ml solution I for 30 mins on
ice. SDS was added to a final concentration of 35mM and EDTA to 50mM.
The preparation was left at room temperature until the solution was clear
(typically 20 mins). The protein was removed by repeated phenol:chloroform
(1:1 w/v) extraction followed by one chloroform:isoamyl alcohol (24:1 v/v)
extraction (see Section 2.3.5). Chromosomal DNA was finally retrieved by
gently pouring 2 volumes of ethanol (chilled at -20°C) down the side of a tube
containing the clear aqueous phase. DNA precipitated at the interface and was
54. 43
spooled out using the rounded end of a pasteur, resuspended in sterile nanopure
water and stored at -20°C.
2.3.2 Small Scale Extraction of Plasmid DNA
Small scale preparation of plasmid DNA used 1.5ml of an overnight culture.
Cells were suspended in lOOpl of a freshly made solution I for 30 mins on ice.
Solution II (200|Lil) was added and the tube was gently mixed and placed on ice
for 5 mins. HOpl of solution III were added to the clear mixture and the tube
was gently mixed and left on ice for 5 mins. The supernatant was recovered
after centrifugation (13400g for 10 mins) avoiding the white pellet. Protein was
removed by one phenol:chloroform (1:1 w/v) extraction followed by one
chloroform:isoamyl alcohol (24:1 v/v) extraction. The DNA was precipitated
by adding two volumes of ethanol (see Section 2.3.5)
2.3.3 Large Scale Extraction of Plasmid DNA
Overnight cultures (400ml) were used for large scale preparation of plasmid
DNA (Birboim and Doly, 1979). The cells were collected in large pots by
centrifugation (3300g at 4°C for 10 mins), resuspended in 10ml of a freshly
made solution I and left on ice for 30 mins. Fresh solution II (20ml) was added,
gently mixed and the whole left on ice for another 10 mins. Solution III (7.5ml)
was added, gently mixed and the whole left on ice for 10 mins. Cell debris was
removed from the plasmid preparation by centrifugation at 4°C for 20 mins at
35000g. Isopropyl alcohol (0.6 volumes) was added to the supernatant, mixed
and left to stand at room temperature for a minimum of 15 mins. DNA was
collected by centrifugation at 4000g for 30 mins at 20°C. The DNA pellet was
air dried for 15 mins and resuspended in sterile nanopure water to a final
55. 44
volume of 17ml. Caesium chloride was added to a final concentration of
Img/ml and ethidium bromide to SOpg/ml. Chromosomal and plasmid DNA
were separated by centrifugation at 40000 rpm using a Sorval TV850 rotor in a
Sorval OTD 60 centrifuge for 20 hrs at 20°C. DNA was visualised under UV
light and the lower band of plasmid DNA extracted. Ethidium bromide was
removed by equilibration with caesium chloride-saturated isopropanol. Caesium
chloride was removed by exhaustive dialysis against distilled water at room
temperature. Plasmid DNA was stored dissolved in sterile distilled water at -20
°C.
2.3.4 Extraction of M13mpl8/19 Template DNA
The recombinant bacteriophage M13mpl8/19 were transformed into JMlOl
(Section 2.2.3) and white plaques were picked into 5ml L-broth containing 100
pi of an overnight culture and incubated at 37°C for 5 hrs with vigorours
aeration. Replicative form DNA and template DNA were obtained from two
1.5ml aliquots of a bacterial culture. The replicative form DNA was extracted
as described for small scale extraction of plasmid DNA (see Section 2.3.2),
whereas the template DNA was isolated from the supernatant. The supernatant
(1.2ml) was mixed with 300pl of a solution containing 2.5M NaCl and 20%
w/v PEG 6000. The mixed solution was left at room temperature for 30 mins.
The phage pellet was recovered by two sequential centriftigations (the second to
remove traces of PEG 6000), then resuspended in 120pl I.IM sodium acetate
pH 7.0, and extracted with an equal volume of phenol:chloroform (1:1 w/v)
followed by one chloroform:isoamyl alcohol (24:1 v/v) extraction (see Section
2.3.5). The template DNA was precipitated with ethanol (see Section 2.3.5).
The template DNA was collected by centrifugation at 13400g for lOmins, dried
in vacuo and resupended in 20pl nanopure water. 2pl of template DNA were
56. 45
visualised by agarose gel electrophoreseis (see Section 2.4) and 4-7pl were
typically used in a sequencing reaction (see Section 2.6)
2.3.5 Phenol Extraction and Ethanol Precipitation
Phenol extraction was performed using one volume of phenol:chloroform
(1:1 w/v) containing 0.1% w/v hyroxyquinoline and equilibrated with lOOmM
Tris-HCl pH 8.0. Chloroform extraction was performed using one volume of a
mixture of chloroform:isoamyl alcohol (24:1 v/v). The aqueous phase was
separated in a Sorval centrifuge (3300g at 20°C for 20 mins) or in a bench top
minifuge for 5 mins at 13400g, then collected avoiding the inter-phase. Ethanol
precipitation was performed with sodium acetate to a final concentration of
300mM and 2 volumes of ethanol at -20°C for minimum of 30 mins. The DNA
was collected by centrifugation either in a bench top niinifuge for 5 mins at
13400g or in a Sorval centrifuge for 30 mins at 3500g.
2.4 Techniques Used in Routine DNA Manipulation
Restriction endonucleases and DNA modifying enzymes were purchased
from Pharmacia Biochemicals Inc. or Life Technologies Ltd (GIBCO/BRL) and
used according to the manufacturers recomendations. Restriction endonuclease
cleavage of DNA was performed typically in 20pl reactions with one unit of
enzyme per pg of DNA at 37°C. T4 DNA ligase was used at 14°C overnight in
T4 ligase buffer (50mM Tris-HCl pH 7.5; lOmM MgClz; ImM ATP). DNA
fragments were separated by agarose gel electrophoresis using 0.7-1.2% w/v
Seakem agarose in TAE buffer (40mM Tris-acetate; ImM EDTA) with 0.5p
g/ml ethidium bromide and visualised using a longwave UV transilluminator.
DNA samples were mixed with the appropriate volume of 6x gel-loading buffer
57. 46
(0.25% w/v bromophenol blue; 0,25% w/v xylene cyanol; 15% w/v Ficoll)
prior to loading. The DNA size markers used was 1 kb ladder (BRL/GIBCO).
For subcloning, the DNA fragment was excised from the gel and the DNA
recovered by Sephaglass-Band-Prep-Kit (Pharmacia) according to the
manufacturers recommendations.
For routine dephosphorylation of plasmid vector, lOpg DNA was incubated
in 100pi calf-intestinal-phosphatase (CIP) buffer (50mM Tris-HCl pH 9.0;
lOmM MgCl2 ; ImM ZnCl2 ; lOmM spermidine) containing ten units of calf-
intestinal alkaline phosphatase and incubated at 37°C for 30 mins. Another ten
units of calf-intestinal phosphatase was added to the reaction and the whole
incubated once more at 37°C for 30 mins. Nanopure water was added to a final
volume of 300pl. The DNA was extracted twice with an equal volume of
phenol:chloroform (1:1 w/v), then once with an equal volume of
chloroform:isoamyl alcohol (24:1 v/v) (section 2.3.5). The phosphatased DNA
was precipitated with ethanol and sodium acetate and collected by
centrifugation at 13400g for 5 mins. The DNA was washed in fresh 70% v/v
ethanol, centrifuged as before, and resuspended in a final volume of 50pl
2.5 DNA Hybridisation Procedures
2.5.1 Transfer of DNA to Nylon Filters
DNA was transferred to filters as described by Southern, 1975. DNA
samples were separated by agarose gel electrophoresis as described above and
the gel photographed along side a linear rule. The DNA was de-purinated by
soaking the gel in 0.25M HCl for 7 mins. The gel was rinsed briefly in distilled
water and placed in denaturing solution (0.5M NaOH; 1.5M NaCl) for 30 mins
58. 47
with occasional shaking. The gel was again rinsed in distilled water and placed
in neutralising solution (0.5M Tris-HCl pH 7.5; 3M NaCl) for another 30 mins
with occasional shaking as before. The gel was rinsed again and placed on six-
sheets of pre-wet (20x SSC) Whatman paper (3mm) without trapping any air
bubbles (20x SSC is 3M NaCl; 0.3M trisodium citrate). A pre-wet (3x SSC)
sheet of nylon membrane (Hybond-N, Amersham International pic) was placed
on the gel with a pre-wet sheet (3x SSC) Whatman paper on top, again taking
care to avoid bubbles. Four sheets of dry Whatman paper were placed above
this with a stack of paper towels. Finally, a glass plate and a 500g weight were
placed on top. The lower sheets of Whatman paper were regularly soaked with
20x SSC and the paper towels changed. The apparatus was left overnight for
the DNA to transfer and then dismantled. The nylon filter was air dried,
wrapped in Saran wrap and exposed to UV light from a long wave
transilluminator for 5 mins to fix the DNA to the filter. Filters were stored at
room temperature in the dark until required for DNA hybridisation (see Section
2.5.4).
2.5.2 Preparation of Filters for Colony Hybridisation
Bacteria were grown overnight at 37°C on an L-agar plate containing the
appropriate antibiotics. A nylon filter (Hybond-N) was placed on top of the
bacterial colonies and left for 10 mins. Whatman paper (3mm) was placed m a
shallow tray and soaked in denaturing solution. The nylon filter was removed
from the L-agar plate and placed on the soaked Whatman paper (colony side
up) for 5 mins. The filter was transfer to Whatman paper, this time soaked in
neutralising solution for a further 5 mins and air dried. For details on the
composition of the solutions see Section 2.5.1. DNA was fixed to the filter by
exposing to longwave UV light from a transilluminator for 5 mins. Cell debris
59. 48
was removed from the filters by gentle scrubbing in 5x SSC using polymer
wool and filters left to air dry in preparation for DNA hybridisation.
2.5.3 Production of a Radiolabelled Probe
Plasmid DNA was cleaved with the appropriate restriction endonucleases and
the fragments separated by agarose gel electrophoresis on a 1% w/v low
melting point agarose gel (BRL). The required DNA fragments were excised
from the gel and added to sterile nanopure water (1.5ml water per gram of
agarose). The sample was placed in a boiling water bath for 7 mins then stored
at -20°C. Prior to use the sample was boiled for an additional 3 mins.
Approximately lOng of DNA was radiolabelled using random hexanucleotide
primers exactly as described by Feinberg and Vogelstein, 1983. Nucleotides
and hexanucleotides were obtained from Pharmacia and [a-32p]dCTP from
Amersham International pic.
2.5.4 Hybridisation of DNA Dnmobilised on Filters with the Probe
Southern blot or colony hybridisation filters were shaken at 65°C m 100ml
of pre-hybridisation solution (see below) for 2 hr. This solution was discarded
and replaced by 20ml hybridisation solution (see below) containing the
radiolabelled probe DNA which had been boiled for 5 mins before adding. The
filter was shaken overnight at 65°C. Hybridisation solution (3x SSC; 2x
Denhardts; 200|Lig/ml salmon sperm DNA; 0.1% w/v SDS; 6% w/v PEG
6000). Prehybridisation solution is the same except with 5x Denhardts (lOOx
Denhardts is 2% w/v Bovine serum albumin V; 20% w/v Ficoll 400; 2% w/v
polyvinylpyrollidone). Solutions were stored at -20°C without salmon sperm
60. 49
DNA. Salmon sperm DNA was sheared by forcing it through a narrow gauge
syringe needle and denatured by boiling prior to use.
After the hybridisation period the filters were washed twice by shaldng in
250ml 2x SSC 0.1 % w/v SDS at 65°C for 15 mins and twice in 0.5x SSC 0.1 %
w/v SDS (0.5x SSC 0.1% w/v SDS is a high stringency wash condition that
allows for approximately 75% DNA homology) (Drake, 1991). SDS
concentration (0.1% w/v) and temperature (65°C) were not varied. The filters
were then air dried completely. The filters were wrapped in Saran wrap for
autoradiography and placed in a cassette carrying intensifying screens. Kodak
X-Omat AR film was exposed to the filters at -70°C. Films were developed in
an Agfa-Geveart automatic processing machine.
2 .6 D N A Sequem clng
Nucleotide sequence was determined by the chain termination method
described by Sanger et al., 1977, in which DNA synthesis from
deoxynucleotide triphosphates is terminated by the addition of
dideoxynucleotide triphosphates. The M13 cloning vectors, M13mpl8 and
M13mpl9 were used to generate single stranded DNA templates (Section
2.3.4). Sequence reactions were performed using the Sequenase Version 2.0 Idt
produced by United States Biochemical Corporation, U.S.A.. The protocol
recommended by the manufacturers was followed using the universal (-40)
primer or oligonucleotide primers synthesised for this purpose. DNA fragments
were radiolabelled by incorporating [a-35S]dATP in the extension reactions.
The radiolabelled fragments were separated by gradient gel electrophoresis
(Biggin et al., 1983). Preparation of the gels used the following solutions.
61. 50
Gel Solution 1 Gel Solution 2
7ml 5x TBE acrylamide/urea mix 40ml O.Sx TBE acrylamide/urea mix
45pi 10% w/v ammonium persulphate 180pl 10% w/v ammonium persulphate
2.5pl TEMED 7.5pl TEMED
0.5x TBE acrylamide/urea mix 5x TBE acrylamide/urea mix
430g urea 430g urea
50ml lOx TBE 150ml lOx TBE
150ml 40 % acrylamide 150ml 40 % acrylamide
per litre 50g sucrose
50mg bromophenol blue
per litre
Electrophoresis grade ammonium persulphate was purchased from BIO
RAD, TEMED from Sigma Chemical Company Ltd. and SEQUEGEL 40%
acrylamide from BDH. lOx TBE is 0.089M Tris-borate, 0.089M boric acid,
0.002M EDTA. To prepare the gel, gel plates (20cm x 50cm) were taped
together separated by 0.4mm spacers. 10ml gel solution 2 followed by 14ml gel
solution 1 were drawn up into a 25ml pipette. Air bubbles were introduced to
form a rough gradient. The liquid was run between gel plates and the cavity
filled with the remaining of gel solution 2. The comb was positioned and the
plates clamped along each side. Gels were routinely freshly made. A vertical
electrophoresis system was used. Running buffer in the top tank was 0.5x TBE
and the lower Ix TBE in accordance with the gradient itself. The gel was
clamped in position with aluminium sheets of a similar dimension as the gel
plates on either side for even heat distribution. The gel was pre-run for 30 mins
at a constant power of 40W and the wells rinsed with running buffer prior to
loading. Electrophoresis was performed at constant power of 40W for 3, 7 and
62. 51
9 hrs. After electrophoresis the gel plates were prised apart and the gel was
soaked in fixing solution (10% v/v methanol and 10% v/v acetic acid) for 15
mius and then rinsed with distilled water. The gel was transferred to a pre-wet
filter paper, covered with Saran wrap and dried under vacuum at 80°C.
Autoradiography used Dupont Cronex film and took place at room temperature.
2.7 Polymerase Chaim Reaction Procedures
PCR amplification reactions contained Ix Taq buffer (lOmM Tris-HCl pH
8.8; 1.5mM MgCfy; 50mM KCl; 0.001% w/v gelatin), lOng template DNA,
primers at 2.5pM, dNTPs at 40|LiM and 2.5 units of Taq polymerase (Sigma
Chemical Company Ltd.) in a total volume of lOOpl. Ultrapure dNTPs were
purchases from Pharmacia Biochemicals Inc.. Reaction mixtures were UV
irradiated on a transilluminator for 15 mins prior to the addition of template
and Taq polymerase. Reaction mixtures were vortexed collected by
centrifugation and overlain with lOOpl sterile mineral oü prior to amplification.
DNA amplification was performed in a Perkin-Elmer Cetus thermal cycler. 30
cycles of the following conditions were performed:
Denaturing step 95°C 1 min
Annealing step 55°C 1 min
Extension step 72°C 3 mins
Following amplifications the PCR product was analysed by agarose gel
electophoresis and the DNA band excised and purified. Direct cloning of the
PCR product was by a modified method of Holton and Graham, 1991. Plasmid
vector DNA (0.5pg aliquots) (for direct cloning of the PCR product) was
cleaved with Sma restriction endonuclease to generate linear blunt end vector.
63. 52
incubated at 70°C for 2 hrs in Ix Taq buffer with 10 pM ddTTP and 5 units of
Taq polymerase. The T-tailed vector was extracted once with
phenol:chloroform (1:1 w/v) and once with chloroform:isoamyl alcohol (24:1
v/v), resupended in nanopure water, pooled and stored at -20°C.
2.8 Radioactive Labelling of Proteins
2.8.1 Minicell Analysis
Minicells were isolated using the procedure described by Hallewell and
Sherratt, 1976. Minicell strains {E. coli DS410) were grown to stationary phase
in 400ml Brain Heart Infusion (if necessary the appropriate antibiotic was
added to the medium). The cells were separated from culture by centrifugation
at 600g for 5 mins. The supernatants were centrifuged at 8500g for 15 mins
and the pellets retained. The pellets were resuspended in 3ml of Ix M9 salts
and niinicells were further purified by two successive sedimentations through
20ml linear gradients of 5-20% sucrose (w/v) in Ix M9 salts at 4650g for 20
mins (4°C). Purified minicells were collected by centrifugation at 9500g for 10
mins and resuspended in Ix M9 salts to a final OD6qo=2.0. Minicells were
either immediately used for protein labelling or aliquoted (lOOpl) in 30% v/v
sterile glycerol and stored at -20°C (for a period of time not exceeding 3
months).
lOx M9 salts per litre: 60g Na2 HP0 4 (337mM)
30g KH2PO4 (220mM)
5g NaCl ( 85mM)
lOg NH4CI (187mM)
64. 53
Proteins were labelled for 45 mins at 37°C in Ix M9 nainimal medium
containing 35§_niethionine (lOOpCi/ml). After a 15 mins chase with cold-
methionine-supplemented broth minicells were lysed by boiling m loading
buffer (0.08M Tris-HCl pH 6.8; O.IM dithiothreitol; 2% w/v SDS; 10% v/v
glycerol; O.lmg/ml bromophenol blue). 20pl (from a final volume of 25pi)
were analysed on a SDS polyacrylamide gel. The upper stacldng gel contained
4.5% acrylamide in 0.125M Tris-HCl pH 6.8 and 0.1% SDS. The lower
running gel contained 15% acrylamide in 0.037M Tris-HCl pH 8.8 and 0.1%
SDS. The running buffer contained 0.02M Tris-HCl, 0.2M glycine, 0.1% SDS
and 2.4mg/l sodium thioglycollate. The gel was run at a constant current of
25mA. After running the gel was soaked in fixing solution (10% v/v acetic
acid; 25 % v/v isopropanol) for 30 mins and treated with Amplify (Amersham)
according to the manufacturers recommendations. The gel was dried under
vacuum at 80°C and autoradiographed at room temperature.
2.8.2 DNA Directed Transcription-Translation System
DNA directed transcription-translation was carried out and used according to
the manufacturers recommendations. The DNA directed transcription-
translation kit was purchased from Amersham International pic. DNA was
prepared as described in Section 2.3.3. DNA (2-5pg) was transcribed and
translated in vitro with an E. coli S30 extract at 37°C for 60 mins. Proteins
were labelled in vitro with L-[35S]methionine (lOOpCi/ml) and the reactions
terminated by placing on ice. The samples were diluted 1:1 with loading buffer
(0.08M Tris-HCl pH 6.8; O.IM dithiothreitol; 2% w/v SDS; 10% v/v glycerol;
O.lmg/ml bromophenol blue) and heated to 100°C for 5 mins prior to loading.
lOpl of sample was loaded onto a SDS polyacrylamide gel and analysed as
described in Section 2.8.1.
65. 54
2.9 Biochemical Assay of p-N-Acetyl-Glacosaminidase
and p-N-Acetyl-Galactosaminidase Activity
Bacterial cultures (100ml) supplemented with the appropriate antibiotics
were incubated at 37°C until mid exponential phase (OD6Qo=0.5), at which
point isopropyl-p-D-thiogalactopyranoside (IPTG) was added to a final
concentration of lOmM when necessary. The cultures were then grown to
00^00=1.0, harvested by centrifugation (3300g at 4°C for 10 mins) and
resuspended in 3ml O.IM MES buffer pH 6.5. The samples were kept on ice
and sonicated with a Braun Labsonic 200 sonicator using a medium probe. The
samples were sonicated for 15 seconds (secs) with 30 secs cooling intervals,
repeatedly, until clearing was visible. Quantitative P-N-Acetyl-glucosaminidase
(EC 3.2.1.30) and N-Acetyl-galactosaminidase (EC 3.2.1.53) biochemical
assays performed in a final volume of 1.0ml. 500pl of O.OIM p-nitrophenyl-N-
Acetyl-P-D-glucosaminide or p-nitrophenyl-N-Acetyl-P-D-galactosaminide,
300-480pl of O.IM MES buffer pH 6.5 and 20-200pl of cellular sonicate were
mixed on ice then incubated at 37°C for 1 hr. The 1ml reaction was terminated
by adding 3mls of 0.2M Borate pH 9.8 and the absorbance measured at 420nm
(measures the amount of p-nitrophenol liberated from the substrate). One
enzyme unit was defined as the amount of enzyme which produced Immol of p-
nitrophenol in 1 min. The amount of protein in the assay reaction was estimated
using a BIO-RAD protein assay Idt with lysozyme as a standard. Enzyme
activity was expressed as units per milligram of protein.
