Multilevel control of extracellular sucrose metabolism in Streptococcus salivarius

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Multilevel control of extracellular sucrose metabolism in Streptococcus salivarius
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Thesis (Ph. D.)--University of Florida, 1991.
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Includes bibliographical references (leaves 96-100).
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by Patricia Dianne Thibos Lawman.
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Vita.

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MULTILEVEL CONTROL OF EXTRACELLULAR SUCROSE
METABOLISM IN STREPTOCOCCUS SALIVARIUS









BY


PATRICIA DIANNE THIBOS LAWMAN


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY



UNIVERSITY OF FLORIDA


1991























Copyright 1991

by

Patricia Dianne Thibos Lawman

























To Michael













ACKNOWLEDGEMENTS


The author wishes to express her sincere gratitude to Dr. Arnold S.
Bleiweis, chairman of her supervisory committee, for his support,
guidance, and friendship. Most of all she would like to thank him for his
faith in her judgment and the freedom given her to create, experiment, fail,
and grow. She would also like to thank the other members of her
committee, Dr. H. Baker, Dr. M. Brown, Dr. D. Duckworth, Dr. A.
Lewin, Dr. A. Progulske-Fox and Dr. B. Chassy, for their helpful
suggestions. Dr. R. Curtiss III is acknowledged for his most graciously
given insights and encouragement. A great deal of appreciation goes to the
members of the Bleiweis laboratory, past and present, for their assistance
and for making this laboratory a great place to work: Dr. G. Ayakawa, L.
Boushell, Dr. J. Brady, B. Buehler, P. Crowley, R. Cullinan, Dr. S. Lee, J.
Hernandez, and D. Piacentini.
This study was supported in part by a Ford Foundation Predoctoral
Fellowship to the author and a Public Health Service grant No. DE-08007
to Dr. Bleiweis.
Special thanks go to the author's parents, Ira and Minnie Lee Thibos,
whose support and faith in her were always present. The support, patience,
and acceptance of the author's children, Josh, Jed, and Sommer are
gratefully acknowledged.









The author would particularly like to thank her husband, Michael, to
whom this dissertation is dedicated, for his unstinting moral support, his
invaluable input, his firm belief in her, and his brilliant sense of humor.














TABLE OF CONTENTS

page

ACKNOWLEDGMENTS.................................................... iv

LIST OF TABLES.............................................................. viii

LIST OF FIGURES............................................................ ix

ABSTRACT................................................. ................... xi

CHAPTERS

1 INTRODUCTION................................................. 1

2 MULTILEVEL CONTROL OF EXTRACELLULAR
SUCROSE METABOLISM IN
STREPTOCOCCUS SALIVARIUS BY
SUCROSE....................................................... 12

Introduction................................................... 12
M ethods........................................................ 14
Results........................................................... 18
Discussion...................................................... 23

3 THE EXTRACELLULAR ENDODEXTRANASE OF
STREPTOCOCCUS SALIVARIUS:
MOLECULAR CLONING AND STUDIES OF
ENZYME REGULATION............................ 39

Introduction.............................................. 39
M ethods...................................................... 40
Results............................................................ 49
Discussion................................................. 54








4 ANALYSIS OF THE EXTRACELLULAR
ENDODEXTRANASE GENE OF
STREPTOCOCCUS SALIVARIUS................... 68

Introduction....................................... ............. 68
M ethods........................................... .............. 70
Results............................................................ 74
Discussion................................................ 77

5 SUMMARY AND CONCLUSIONS......................... 85

LIST OF REFERENCES............................................. 96

BIOGRAPHICAL SKETCH................................................. 109













LIST OF TABLES

2-1. Effect of Antibiotics on Dextranase and
Fructanase Production............................... 36

2-2. Effect of Antibiotics on Glucosyltransferase
Production........................................................ 37

2-3. Effect of Antibiotics on Fructosyltransferase
Production................................................... 38

3-1. Carbon Source Utilization by
S. salivarius PC-1...................................... 67

5-1. Sequence analysis at the nucleotide level of
streptococcal genes involved in sucrose
metabolism .................................... ............. 94

5-2. Sequence analysis at the protein level of
streptococcal genes involved in sucrose
metabolism............................... .......... 95


viii













LIST OF FIGURES


page
1-1. Diagramatic representation of sucrose metabolism
in S. salivarius..... .................................................... 11

2-1. Carbon source utilization by S. salivarius PC-1............. 30

2-2. Effect of carbon source on dextranase and fructanase
production by S. salivarius PC-1............................. 31

2-3. Effect of carbon shift on production of dextranase
and fructanase in S. salivarius PC-1.......................... 32

2-4. Effect of carbon source on cell-associated and
extracellular GTF production in S. salivarius PC-1.... 33

2-5. Effect of carbon source on cell-associated and
extracellular FTF production in S. salivarius PC-1..... 34

2-6. Effect of sucrose shift on production and distribution of
GTF and FTF in S. salivarius PC-1........................ 35

3-1. Screening the S. salivarius genomic library
for dex and gtf recombinants.............................. 61

3-2. Detection of electroblotted dextranase activity on
blue dextran-agarose ............................................... 62

3-3. Substrate specificity of native and recombinant
dextranases........................................................... 63

3-4. Product analysis of native and recombinant
dextranases............................................................. 64

3-5. Sucrose-mediated release of dextranase
inhibition in PC-1 cell-free supernatant................. 65











3-6. Effects of CDM/galactose and CDM/galactose
plus sucrose cell-free S. salivarius PC-1
supernatants on recombinant dextranase
activity.............................................................. 66

4-1. Autoradiographs of L-[S35] methionine-labeled proteins
from in vitro translations of Lambda ZAP II
and PD1.......................................... ......... ... 80

4-2. Excision of pBluescript and dex insert...................... 81

4-3. Partial restriction map of pPD13 containing the 2.6 kb
Xho I/Not I fragment of PD1.................................. 82

4-4. Construction of the original PD1 clone and deduced
restriction map of the 6.3 kb fragment containing
the dex gene...................................... .......... 83

4-5. Southern blot analysis of complete restriction digests of
the S. salivarius PC-1 chromosome............................. 84









Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy


MULTILEVEL CONTROL OF EXTRACELLULAR SUCROSE
METABOLISM IN STREPTOCOCCUS SALIVARIUS

By

Patricia Dianne Thibos Lawman

May, 1991




Chairman: A. S. Bleiweis, Ph.D.
Major Department: Immunology and Medical Microbiology

In order to study the role of sucrose in the regulation and control of
its metabolism by Streptococcus salivarius, standardized experimental
conditions were established. It was found that de novo synthesis was
required for the production of extracellular glucosyltransferase activity
which, upon the addition of sucrose, became associated with the cell
surface. Conversely, cell-associated fructosyltransferase activity required
genetic induction for production and cell-surface association, but required
sucrose for release from the surface framework. Extracellular fructanase
activity was twofold higher when cells were grown in sucrose than when
cells were grown in other sugars. This increase occurred within 5 minutes
but was diminished by rifampicin and chloramphenicol. The extracellular
dextranase activity of cells grown in sucrose was tenfold higher than that of
cells grown in other sugars. This activity increased within 5 minutes
following the addition of sucrose to galactose-grown cells and was affected









by neither rifampicin nor chloramphenicol. Dextranase appeared to be
tightly controlled by a dextranase inhibitor that was displaced by sucrose,
and by at least one other factor, that regulated or directed hydrolysis of
dextran substrates.
To analyze the function and regulation of the extracellular
endodextranase (xa-l,6-glucan hydrolase, EC 3.2.1.11) from S. salivarius, a
molecular approach was taken. Dextranase-positive recombinants from a
genomic library were identified by their ability to clear zones in blue
dextran-containing agar. One clone, PD1, contained a promoter element
and structural gene which encoded an active 190 kilodalton protein. The
native dextranase had an apparent molecular mass of 110 kilodaltons.
Streptococcus salivarius contained a single copy of the endodextranase
gene. Product analysis of dextranase activity on various substrates by thin
layer chromatography revealed the expected isomaltosaccharides produced
by the recombinant enzyme, but was unable to resolve the larger
polysaccharide products of the native supernatant. Streptococcus salivarius
utilized neither substrates nor products of dextran hydrolysis for growth,
suggesting that dextranase has a synthetic role in this organism. The
versatility in both genetic and biochemical control mechanisms, of this
complex set of enzymes allows tight regulation and quick response to
environmental stimuli, a potentially invaluable characteristic for an
inhabitant of the oral cavity.













CHAPTER 1
INTRODUCTION

Streptococcus salivarius is a Gram-positive coccoid bacterium which
is capable of growing in short or long chains (Hardie, 1986). This
organism grows in the presence of oxygen and can be found in the oral
cavity of humans and animals where it is primarily associated with the
tongue and saliva, but also can be isolated from fecal material. Some
strains have been linked to infectious endocarditis and dental caries in
gnotobiotic rats (Hardie, 1986).
It also has been shown that a significant number of S. salivarius
isolates have the ability to adhere to other structures in the oral cavity
besides epithelial surfaces, namely, the pellicle on the teeth (Weerkamp &
McBride, 1980). Streptococcus salivarius may be a primary colonizer,
providing an initial attachment site for more fastidious, less oxygen-
tolerant organisms. As S. salivarius is able to attach to tooth surfaces, but
is found in low numbers in dental plaque (Weerkamp & McBride, 1980), it
has been suggested that some adherence capabilities are rarely or never
used. Furthermore, the ability of S. salivarius to adhere may require
protease-sensitive components (Weerkamp & McBride, 1980) and may also
involve covalent linkage to the peptidoglycan structure (Weerkamp &
Jacobs, 1982).
Streptococcus salivarius is a successful inhabitant of the human oral
cavity. In addition, it is capable of forming aggregates with several oral
anaerobic, Gram-negative bacteria (Gibbons & Nygaard, 1970; Weerkamp









& McBride, 1981) and may form the basis for their initial colonization.
The ability to attach to host surfaces and to adhere to other bacteria may be
mediated by surface appendages associated with a fibrillar layer, termed
the "fuzzy coat." This extracellular layer is present on S. salivarius
(Gibbons et al., 1972) and other streptococci, such as Streptococcus
pyogenes (Ellen & Gibbons, 1972). Two morphologically distinct types of
fibrils and several factors mediating attachment to host surfaces and other
oral bacteria (Weerkamp & McBride, 1981) are found on the surface of S.
salivarius (Handley et al., 1984).
The role of sucrose-derived extracellular polysaccharides synthesized
by oral streptococci in cariogenicity, adherence, and colonization has been
well established (Hamada & Slade, 1980). Extracellular enzymes of S.
salivarius involved in sucrose metabolism include: glucosyltransferases
(GTFs; EC 2.4.1.10), which catalyze the synthesis of glucans from sucrose;
fructosyltransferases (FTFs; EC 2.3.1.10), which catalyze the synthesis of
fructans, either levan or inulin, from sucrose; dextranase (a-l,6-glucan
hydrolase; EC 3.2.1.11), which partially degrades glucan; and fructanase
(P-D-fructan fructohydrolase; EC 3.2.1.80), which releases free fructose
by hydrolyzing fructan polymers.
These enzymes are part of a very complex system including catabolic
enzymes which break down not only sucrose, but products derived from
sucrose, such as dextran, levan, and inulin (Chassy et al., 1974; Hamada et
al., 1975; Schachtele et al., 1975b; Russell, 1979; Takahashi et al., 1985;
Bume et al., 1987) (Figure 1-1). There are also anabolic enzymes which
catalyze the synthesis of products using sucrose or sucrose-derived
substrates (Slee & Tanzer, 1980; Hamada & Slade, 1980; Furata et al.,
1985). These processes occur both intracellularly and extracellularly and,









in the former case, require specialized transport systems for sucrose and
sucrose-derived products (Ellwood & Hamilton, 1982; Postma & Lengeler,
1985; Lodge & Jacobson, 1988; Reizer et al., 1988; Russell & Ferretti,
1990).
Despite the large amount of information available pertaining to the
a-D-glucosyltransferases, there are still many fundamental questions which
remain unanswered. It has been difficult to make valid comparisons
between the different GTFs because their properties depend on the strain
producing them, the isolation procedures used, growth conditions and the
number of GTFs produced (Montville et al., 1977). In fact, the exact
number and nature of GTFs required for glucan synthesis is still
controversial. A number of enzymes tend to form aggregates which
include invertase, FTF, dextranase, uncharacterized proteins, as well as
lipids, peptidoglycan, teichoic acids, and carbohydrates (Montville et al.,
1977), making isolation extremely difficult. It is reasonable to assume that
many studies have been performed with non-homogeneous fractions and
that inaccurate conclusions have been drawn. The analysis of the glucan
products of these enzymes, i.e., the determination of the glycosidic
linkages, the distribution of branches, and molecular weights has been
equally difficult (Walker et al., 1988). Since growth conditions and rates
determine the characteristics of the glucan produced, the ratio of a-(1,3)
to a-(1,6) glycosidic bonds, and consequently the solubility properties of
the dextran product, vary accordingly (Walker & Jacques, 1987). It has
not been possible, even with recombinant DNA techniques, to clarify the
relationship between individual enzymes and the type of linkage synthesis
they catalyze or the cooperation between multiple forms of
glucosyltransferases.









Even less progress has been made in the characterization of the P-D-
fructosyltransferases and the fructans they produce. The levan produced
by S. salivarius has been characterized as a levan possessing P-(2,6)
linkages with 3-(2,1) branchpoints (Hancock et al., 1976). It is thought
that a single FTF is capable of synthesizing both P-(2,6) and 3-(2,1)-
fructans (Jacques et al., 1985b). The relative amount of fructan to dextran
in plaque and saliva is small, probably due to rapid hydrolysis of the
former offsetting its greater rate of synthesis (Wood, 1969). Although S.
salivarius makes large amounts of fructan, it also produces a P-D-fructan
fructohydrolase (Burne et al., 1987) which hydrolyzes the levan polymer
to free fructose, which in turn is catabolized mainly to lactic acid.
Bacteria have evolved mechanisms by which full advantage is made
of exogenously supplied nutrients, even though entry of these nutrients into
the cell is not without cost. Virtually all biosynthetic pathways,
polymerization reactions, and assembly processes require coordination if
orderly growth is to occur. Control of such metabolic pathways can be
established by regulating the kinds and amounts of macromolecules made,
by control of gene expression and subsequent control of enzyme activity.
Modulation of enzymatic activity by inhibition or activation of allosteric
proteins by their specific effectors and by substrate/product concentrations
is best understood (Ingraham et al., 1983).
Until recently, the enzymes involved in sucrose metabolism in oral
streptococci had been assumed to be constitutive as they are produced in the
absence of substrate (Janda & Kuramitsu, 1978; Wenham et al., 1979).
Although there have been suggestions in the literature that sucrose
metabolizing enzymes may be under some form of regulation (Keevil et
al., 1983; Walker et al., 1983; Walker & Jacques, 1987), it has been









difficult to show this conclusively due to the difficulties in isolating
individual enzymes, variation between strains, differences in enzyme
production due to growth conditions, and the general complexity in the
network of these and other enzymes which metabolize sucrose.
Understanding the regulation and control of enzymes involved in sucrose
metabolism will be necessary for an in-depth analysis of the role of this
organism in oral pathology.
Work presented in this dissertation shows that this organism uses a
variety of mechanisms to control this set of enzymes, allowing the
regulation of their expression and function at several widely separated
stages in their production. A first step in establishing whether or not these
enzymes are under any form of regulation was to determine the role of
sucrose in the control of its metabolism. Once this was accomplished, the
scope of the study was narrowed to one enzyme, dextranase.
A major goal of this study was to determine the contribution of
dextranase to the physiology of S. salivarius. It was important to
distinguish between two proposals: 1) that the primary function of
dextranase, although a hydrolytic enzyme by definition, is, in fact to act as
a synthetic catalyst; or 2) that its major contribution is the degradation of
large glucans to smaller polymers serving as substrates for catabolism.
In a putative biosynthetic capacity, it is possible that dextranase could
serve to provide primer or branchpoints for the glucosyltransferases
(Walker, 1972; Germaine et al., 1977; Felgenhauer & Trautner, 1983).
The ratio of a-(1,3)- to a-(1,6)-glycosidic linkages and therefore the
solubility properties of dextran made by S. salivarius and other organisms
may be altered, if involvement in glucan formation is a major function of
dextranase. Since the degree of solubility of glucans has been shown to be









an important factor in colonization, plaque formation, and caries
development, dextranase then would have a great potential for impacting
on these processes (Walker & Jacques, 1987).
Alternatively, if dextranase were acting in a catabolic pathway, to
break down dextran eventually to yield glucose, it would be acting as an
antagonist to the extracellular GTFs, thus explaining the low apparent
levels of GTF activity (determined by the amount of dextran produced) in
this organism (Hamada et al., 1975; Schachtele et al., 1975). If this were
the case, the presence of dextranase would obscure any conclusions made in
studies of GTF in any organism producing both these enzymatic activities
and consequently, any reaction influenced by dextran, i.e., adherence,
aggregation, and the distribution of GTFs between cell-surfaces and culture
supernatants (Walker et al., 1981).
Extracellular endodextranase hydrolyzes large dextran molecules to
smaller polymers, predominantly oligoisomaltosaccharides (Dewar &
Walker, 1975). These short isomaltosaccharides should be readily
transported across the cell membrane, where a second enzyme, such as the
intracellular exodextranase described for Streptococcus mutans (Burne et
al., 1986; Russell & Ferretti, 1990), would hydrolyze them completely to
glucose. The tandem action of the endo- and exo-dextranases would be an
energy efficient system for the rapid hydrolysis and utilization of dextran.
If this were true, the extracellular endodextranase of S. salivarius could
play a key catabolic role in the physiology of this species and enable it to
use the dextran product of its own GTFs and/or those produced by other
oral organisms as short term energy reservoirs. Parker and Creamer
(1971) demonstrated that polysaccharides could serve as significant









reserves and that some glucans could support growth of some oral
streptococci.
Considering the fact that dextranase is produced by many strains of
oral streptococci (Dewar & Walker, 1975; Schachtele et al., 1975a; Ellis &
Miller, 1977; Felgenhauer & Trautner, 1983), it is surprising how little is
known about this enzyme. Most information to date on dextranase in oral
streptococci has been obtained from Streptococcus sobrinus (Barrett et al.,
1986; Walker et al., 1988). The preliminary studies presented in this
dissertation are the first dealing with dextranase from S. salivarius.
Barrett et al. (1987) developed a purification scheme for dextranase
present in S. sobrinus culture supernatant fluids. These investigators
recovered the majority of the dextranase in two forms, having molecular
weights of 175 KD and 160 KD. The lower molecular weight form was
thought to be a proteolytic breakdown product of the 175 KD dextranase.
This same group (Wanda, 1990) reported the molecular mass of the native
enzyme to be 280 KD 200 KD, by gel filtration. When S. sobrinus gene
libraries were screened for dextranase activity, three phenotypes of
recombinant clones were identified. A Pst I contiguous library produced a
175 KD dextranase, a Sau 3A noncontiguous library produced a 150 KD
dextranase, and an Eco RI noncontiguous library produced both
dextranases (Barrett et al., 1987). The variation in molecular weight was
ascribed to either deletions of C-terminal ends in the noncontiguous
libraries, "scrambling" of the dextranase structural gene, or the cloning of
two separate genes. The authors believe the latter is unlikely due to genetic
characterizations of S. sobrinus dextranase mutants which suggested a
single dextranase gene. The clone from the Sau 3A library was chosen for
further study. The dextranase from this construct was determined to be a









periplasmic enzyme in E. coli with a molecular weight ranging from 140
KD to 80 KD (by SDS-PAGE with blue dextran) (Wanda, 1990). Several
subclones were obtained by a partial digestion with Pvu II and religation.
The majority of the dex gene was found to reside on a 2.6 kb Pvu II
fragment; however, this fragment did not contain the dex promoter.
Attempts to obtain sequence of the dex gene of S. sobrinus have not been
successful to this point (R. Curtiss HI, personal communication).
The following chapters show that extracellular dextranase activity
was higher in sucrose-grown S. salivarius PC-1 cells compared to cells
grown in glucose, fructose, or galactose (Figure 2-2); that dextranase
activity increased 100-fold when sucrose was added to cells growing in
galactose (Figure 2-3) and that this increase was not affected by
transcriptional or translational inhibitors (Table 2-1); that dextranase was
active when cells were grown in sucrose and inactive in galactose-grown
cells (Figure 3-5a); and that sucrose added to cell-free S. salivarius
CDM/galactose culture supernatant activated the native dextranase (Figure
3-5b). These results suggest that activation of native dextranase by sucrose
may involve the displacement of a dextranase inhibitor.
Hamelik and McCabe (1982) concluded that the presence of an
inhibitor in batch-grown culture fluids of S. mutans accounted for the
absence of endodextranase activity in strains known to produce this
enzyme. This finding was in direct conflict with earlier conclusions that
endodextranase production was growth-dependent and that the enzyme was
labile at pH 5.0 (Walker et al., 1981). Sun et al. (1990) have cloned and
sequenced the dextranase inhibitor gene (dei) from S. sobrinus. The dei
gene specifies a 330 amino acid protein with a molecular weight of
approximately 36 KD and carries its own promoter. DNA from serotypes









a,d, and g of mutans streptococci was recognized by Southern hybridization
using the dei probe. The widespread distribution of a dextranase inhibitor
(Sun et al., 1990) may represent a common theme of postgenetic regulation
of this enzyme, making this a feasible model for studying enzyme
regulation in oral streptococci. The apparent presence of such an inhibitor
in S. salivarius is explored further in Chapter 3.
Since S. salivarius is a common inhabitant of the oral cavity, is
among the earliest colonizers after birth, and constitutes a major
proportion of the streptococci found on soft tissue in the mouth, it is likely
that any extracellular enzyme produced by this organism would have a
profound effect on the oral ecology. All indications are that dextranase has
the ability to participate in plaque formation and modification through its
effects on the polysaccharides thought to be important in bacterial
aggregation and adherence (Gibbons & Van Houte, 1975). Dextranase may
act directly in conjunction with GTFs in the modification of dextran
polymers, by providing primer or branchpoints and thus may be involved
in determining the degree of dextran solubility. This enzyme may also
break down the accumulated dextran in plaque, destabilizing the plaque
matrix and at the same time providing glucose for consumption by plaque
organisms. It also is conceivable, as previously suggested, that dextranase
may be important in the physiology and growth of S. salivarius, giving this
species a competitive edge in the oral environment.
The fact that dietary sucrose would serve to activate dextranase is of
considerable interest since it is under these nutritional conditions that dental
caries has been shown to occur. This is especially true since the ability to
make dextran has been held responsible for allowing colonization by
cariogenic organisms. An alternative explanation for the correlation









between sucrose and caries may be that dextranase provides additional
substrate for acid production as it breaks down the stored dextran. To test
this hypothesis further, the polysaccharides which serve as substrates for
dextranase were determined. The ability of S. salivarius to grow in
dextran or possible products of glucan hydrolysis also was established. The
results of these studies are presented in Chapter 3.
Proteins which bind glucans and enzymes which are involved in the
metabolism of glucans contain glucan binding regions (Ferretti et al.,
1988; Sato & Kuramitsu, 1988; Uedo et al., 1988; Sato et al., 1989; Banas
et al., 1990). It would be of interest to compare the nucleotide and amino
acid sequences of the dex gene to sequences of other such proteins. This
should allow the division of this protein into functional domains, i.e.,
glucan binding and catalytic regions. The DNA sequence should also give
specific information about initiation and termination signals, promoter and
coding regions, and a more accurate estimate of the molecular weight of
dextranase. To accomplish this, it was first necessary to clone and
characterize the S. salivarius dex gene and its product. This information is
found in Chapters 3 and 4.









Extracellular


Exodextranase


IMs r

II Porin


Glucose


GTF Dextran
so_ +
Fructose
SUCROSE


Glucose PEP PTS
Permease (ATP)


FTF Glucose II
+ II
Fructan





Fctos II
Fructose


Sucrose PEP PTS
Permease (ATP)


Invertase
Glu + Fru

SS-6-P


S-6-P-hydrolase


Fructose PEP PTS
Permease (ATP)


ATP-dep Proton Motive Force


Figure 1-1.
salivarius.


Diagramatic representation of sucrose metabolism in S.


Glucose

Fructose
Fructose


Intracellular


rT-














CHAPTER 2
MULTILEVEL CONTROL OF EXTRACELLULAR SUCROSE
METABOLISM IN STREPTOCOCCUS SALIVARIUS BY SUCROSE


Introduction


Sucrose has a marked impact on the microbial ecology of the oral
cavity. This common disaccharide enhances plaque development and is
associated with dental caries (de Stoppelaar et al., 1970; Carlsson &
Johansson, 1973; Gibbons & van Houte, 1973; Hamada & Slade, 1980).
Streptococcal enzymes which metabolize sucrose, some to synthesize
polysaccharides important in microbial adhesion or cohesion (Gibbons &
van Houte, 1975; Wenham et al., 1981; Koga et al., 1986), and others to
hydrolyze these polysaccharide products, have been shown to be virulence
factors in the mutans group of oral streptococci (Curtiss et al., 1987; Sato
et al., 1987; Schroeder et al., 1989). However, little has been done to study
analogous enzymes in Streptococcus salivarius, a major inhabitant of the
soft tissues of the oral cavity.
The complex system of extracellular enzymes involved in sucrose
metabolism include glucosyltransferases (GTFs; EC 2.4.1.5), which
catalyze the synthesis of glucans from sucrose; fructosyltransferases
(FTFs; EC 2.4.1.10), which catalyze the synthesis of fructans, either levan
or inulin, from sucrose; dextranase (a-1,6-glucan hydrolase, EC
3.2.1.11), which partially degrades glucan; and fructanase (p-D-fructan









fructohydrolase, EC 3.2.1.80), which releases free fructose by
hydrolyzing fructan polymers. Since these enzymes are produced in the
absence of substrate, they have been assumed to be constitutive in oral
streptococci (Janda & Kuramitsu, 1978; Wenham et al., 1979). Although
there have been suggestions in the literature that this might not be the case
(Keevil et al., 1983; Walker et al., 1983; Walker & Jacques, 1987), it has
been difficult to show conclusively that these enzymes are under any kind
of regulatory control. This has been due to difficulties in isolating
individual enzymes, variation between strains, differences in enzyme
production due to growth conditions, and the general complexity in the
network of these and other enzymes which metabolize sucrose.
Streptococcus salivarius is a common inhabitant of the oral cavity,
colonizing preferably the tongue dorsum and buccal epithelium (Weerkamp
& McBride, 1980) as well as being found in the saliva. Although it is not
known to be a major oral pathogen, this species possesses a variety of
anabolic (i.e. GTF, FTF) and catabolic (i.e. dextranase, fructanase)
extracellular enzymes associated with sucrose metabolism (Chassy et al.,
1976; Takahashi et al., 1983; Houck et al., 1987). Studies described below
indicate that sucrose plays a role in the regulation of its metabolism in this
microbial species and that significant versatility is maintained in the
mechanisms by which this is accomplished (Townsend & Bleiweis, 1989).









Methods

Bacterial strain and growth conditions,

A fresh isolate of S. salivarius was used in these studies. This isolate,
designated PC-1, was isolated in this laboratory. Subculturing was kept to
a minimum by using stock cultures maintained at -70oC in 25% (v/v)
glycerol.
A chemically defined medium (CDM), prepared according to
Terleckyj et al. (1975) was used for bacterial growth in all experiments
described in this paper. The energy source (sugar) varied depending on
the experimental conditions as described in Results. Reagent grade sugars
were purchased from Sigma Chemical Co. All cultures were grown at
370C.

Cell growth, harvest and preparation.

Growth curves were generated by inoculating CDM or CDM/sugar
(glucose, fructose, galactose, glucosamine, or sucrose) (10 mM) with a
CDM/glucose (10 mM) starter culture at a 1:10 ratio. A Klett-Summerson
photoelectric colorimeter, with a No. 54 filter, was used to take optical
density measurements every 30 minutes until stationary phase was reached.
Each data point in Figure 2-1 represents the mean of triplicate samples
from a representative experiment. The pH of each culture was determined
at the same time intervals using colorpHasTM indicator sticks (MC/B
Manufacturing Chemists, Inc.).
To establish the basal level of the enzymatic activities in S. salivarius
cells, a CDM/glucose (10 mM) starter culture was added at a ratio of 1:10









to CDM/glucose, fructose, galactose, or sucrose. When cultures reached
the desired density (Klett 75, mid-exponential phase) the cells were
harvested by filtration using a 0.2 mm membrane filter (Gelman Sciences,
Inc.), washed once with 50 mM potassium phosphate buffer, pH 6.35 (the
buffer used throughout this study), collected by centrifugation (4000g,
200C, 5 minutes), and then resuspended to a tenfold concentration
equivalent in the same buffer containing 10 mM NaF (McCabe & Smith,
1975). The cells were held at 40C until enzymatic assays were performed
(within 24 hours). The supematants from these cultures were concentrated
immediately using CentriprepT concentrators (Amicon) at 3000g, dialyzed
overnight and reconstituted to a tenfold concentration equivalent in the
potassium phosphate buffer. The supernatants were assayed for enzymatic
activity after dialysis.
For sucrose shift assays, a PC-1 starter culture, grown as above, was
added to CDM/galactose. This culture was grown to a Klett reading of 75.
At this time sucrose was added to a final concentration of 10 mM and
aliquots were taken over time as indicated in Figures 2-3 and 2-6. The
cells and culture fluids then were harvested and concentrated as above.
Aliquots of cells collected from CDM/galactose cultures also were washed
vigorously two times with 0.04% SDS and several concentrations of Tween
80 (0.02, 0.04, 0.06%), and twice with 50 mM potassium phosphate buffer,
pH 6.35. The cells were harvested by centrifugation (4000g, 200C, 5
minutes) after each wash, finally resuspended to a tenfold concentration
equivalent in the same buffer containing 10 mM NaF, and assayed for
retention of GTF and FTF activities.
Antibiotic inhibition studies were conducted in a manner similar to
the sucrose shift assays. A starter culture was added to CDM/galactose in









the proportions given above and grown to the same optical density. At this
time, sucrose (10 mM), chloramphenicol (100 mg ml-1), rifampicin (100
mg ml-1), or combinations of these were added to aliquots of the culture
and then incubated at 370C for 1 hour. The cells and supernatants were
treated as above prior to enzyme assays. Control cultures contained none
of these reagents.

Assay of enzyme activities.

Dextranase and fructanase activities were determined using a
standard assay for reducing sugars designed by Somogyi (1951) and Nelson
(1944). Substrate, either dextran Type 100C (Sigma Chemical Co.) or
levan from Aerobacter levanicum (Sigma Chemical Co.) at a final
concentration of 0.25 mg ml-1 was incubated with cells or supernatant as
prepared above. Reactions were stopped after 180 minutes. The amount
of reducing sugar released (glucose or fructose depending on the substrate)
was determined by comparison with glucose or fructose standards. One
unit of enzyme activity (U) was defined as the amount of dextranase or
fructanase catalyzing the release of 1 mmol reducing sugar (glucose or
fructose) min-1 ml-1. Each data point shows the mean (+/- SD) of
triplicate samples from a representative experiment.
GTF and FTF activities were quantitated by a modification of the
standard procedure developed by Robrish et al., (1972) where sucrose,
labeled either in the glucose or fructose moiety, was incorporated into
ethanol-precipitable polymer. Reaction mixtures consisted of tenfold
concentrated sample (cells or supernatant), 1 mM sucrose carrier, 0.25 mg
ml-1 commercially obtained dextran or levan primer, and labeled sucrose,









[glucose-14C(U), 7.7 x 10-5 mmol ml-1, at 261.0 mCi mmol-1] or
[fructose-14C(U), 7.5 x 10-5 mmol ml-1, at 267.0 mCi mmol-1] (NEN
Research Products). Reactions were carried out in a total volume of 100 pl
in 12 x 75 mm borosilicate disposable culture tubes at 370C for 90
minutes. The polymer was precipitated with 300 ml ice-cold 95% (v/v)
ethanol for at least 30 minutes. The ethanol insoluble product was
collected on 25 mm extra thick glass fiber filters (Gelman Sciences, Inc.)
cut to fit disposable MicrofoldsT (V & P Scientific, Inc.). The microfold
system is a 96 chamber micro-plate with a small opening in the bottom of
each well. The filter rests on top of the opening and immobilizes the
precipitate when vacuum is applied to the bottom of the manifold. Each
filter was washed once with ice-cold 95% (v/v) ethanol and air-dried
before being placed in vials with 3 ml aqueous counting scintillant (ACS)
(Amersham). Each sample was counted for 5 minutes in a Beckman LS
3801 Liquid Scintillation System. The adaptation of this procedure to a
microassay system made it possible to handle multiple samples with greater
accuracy and reproducibility. One unit of enzyme activity (U) was defined
as the amount of GTF or FTF that catalyzed the incorporation of 1 nmol of
the glucose or fructose moeity of labeled sucrose min-1 ml-1. Each data
point represents the mean (+/- SD) of triplicates from a representative
experiment. Depending on experimental conditions employed, 65-95% of
the polymer collected from GTF activities and 90% of the fructan
synthesized in the FTF assays was water-soluble. No attempt was made to
correlate these properties with regulatory effects as that distinction was
beyond the limits of these assays. Therefore, only the total amount of
polymer produced was reported.









Results

Carbon source utilization of S. salivarius PC-1.

In order to develop appropriate culture conditions, S. salivarius PC-
1, was grown in a number of carbohydrate carbon sources (Figure 2-1).
When cells were grown in glucose the doubling time at mid-exponential
phase was 49 minutes. It was determined that the growth rates at mid-
exponential phase were similar for cells grown in either fructose (60
minutes), sucrose (60 minutes), or galactose (62 minutes); however, the
lag time for fructose-grown cells was longer than for the other cultures.
Cells did not grow in glucosamine. Cells taken at a Klett reading of 75
were at mid-exponential phase regardless of the carbon source. This
allowed the normalization of all subsequent experiments to this value, and
the use of galactose as a "non-sucrose related" carbon source in sucrose
shift assays. Cultures grown in the various sugars and harvested at an
optical density of Klett 75 had a range of pH values between 6.1 and 6.4.
Over the duration of the growth experiments, all cultures shifted from an
initial pH of 7.0 to a final pH of 4.0 when glucose, galactose, fructose, or
sucrose was used as the growth substrate.

Dextranase and fructanase activities of S. salivarius PC-1.

Cultures of PC-1 were grown in 10mM glucose, fructose, galactose,
or sucrose, and the dextranase and fructanase activities of cells and culture
fluids were measured. Under all conditions tested, cell-associated activity
for these enzymes was not detected indicating that both dextranase and
fructanase are extracellular enzymes. If an intracellular dextranase exists









in this organism, as in related streptococci (Dewar & Walker, 1975;
Walker et al., 1980; Walker et al., 1981; Russell & Ferretti, 1990), its
activity was not measurable under these conditions. Dextranase levels were
similar whether the cells were grown in glucose, fructose, or galactose; the
same held true for fructanase (levanase) levels (Figure 2-2). Growth in
sucrose; however, resulted in a twofold higher fructanase activity and a
tenfold higher dextranase activity. From these preliminary experiments it
appeared that growth in sucrose resulted in significantly increased levels of
both dextranase and fructanase activities in culture supematants.

Effects of sucrose shift on dextranase and fructanase activities.

Having noted the effect that growth in sucrose had on the production
of both dextranase and fructanase, it was of interest to measure the
response of these enzymatic activities to the addition of sucrose to cells
growing in galactose (Figure 2-3). The addition of sucrose resulted in an
immediate (< 5 minutes) increase in both dextranase and fructanase
activities followed by a period of stabilization. While fructanase levels
increased by 64% upon the addition of sucrose, dextranase levels increased
more than 100-fold.

Effect of antibiotics on dextranase and fructanase production.

To determine whether the sucrose effect described above was being
implemented at the genetic level, previously determined inhibitory
concentrations of antibiotics specifically targeted to transcription
(rifampicin) or translation (chloramphenicol) were tested. In the absence
of sucrose, dextranase levels in culture fluids remained very low whether









the cells were untreated, or treated with chloramphenicol or rifampicin
(Table 2-1). However, there was a significant increase (100-fold) in
dextranase activity upon the addition of sucrose, even in the presence of
chloramphenicol or rifampicin. It appears, therefore, that the increases in
dextranase activity observed in these experiments do not reflect genetic
induction, and that regulation at the post-translational level is likely.
Fructanase activity measured under the conditions described was relatively
high whether the cells were grown in glucose, fructose, or galactose
(Figure 2-2); therefore, the net increase upon the addition of sucrose was
not as dramatic as that seen with dextranase activity (Figure 2-3).
Furthermore, unlike dextranase, this increase was reduced significantly,
from 78% with sucrose alone to 47% and 33%, respectively, by
transcriptional and translational inhibitors (Table 2-1), indicating a possible
genetic induction.

GTF and FTF activities of S. salivarius PC-1.

These experiments were performed with and without a dextran or
levan primer for GTF or FTF respectively. The addition of primer
increased the incorporation of radiolabeled glucose in the GTF assays by
approximately 15-25% and the incorporation of radiolabeled fructose in
the FTF assay by 10-15%. Data presented here are from experiments that
include primer.
The measurable activities of both cell-associated or released GTF
were quite low. This probably was due to the presence of dextranase
which was able to solubilize the radiolabeled products. Supporting this
theory was the apparent inverse relationship between the amount of GTF









activity (Figure 2-4) and dextranase activity (Figure 2-2) measured under
these conditions. Cell-associated GTF activity was similar in cells grown on
sucrose, galactose, or glucose, and higher in cells grown on fructose
(Figure 2-4). Glucose-grown cells produced the highest apparent activity of
the secreted form of GTF as compared to fructose-, sucrose-, or galactose-
grown cells.
Cell-associated FTF activity was greater than extracellular activity
when cells were grown in glucose, fructose, or galactose (Figure 2-5).
However, soluble FTF activity increased when cells were grown on
sucrose. The total FTF activity for sucrose-grown cultures was about
twofold greater than for galactose-grown cultures (Figure 2-5).

Effect of sucrose shift on GTF and FTF activities.

When sucrose was added to cells growing in galactose, cell-associated
GTF activity appeared to increase initially, but remained low (Figure 2-
6a). In contrast, released activity showed an initial decline before
returning to original values, suggesting a sucrose-mediated, cell-association
of secreted GTF. As seen in Figure 2-5, the majority of FTF activity was
cell-associated when the cells were grown in galactose. However, within 5
minutes after the addition of sucrose, there was a complete reversal of the
location of FTF activity (Figure 2-6b). This translocation effect was
immediate and profound.
To investigate the nature of the cell-surface interactions of these
enzymes, galactose-grown cells were subjected to a number of different
washing procedures and then assayed for GTF and FTF activities. The
0.04% SDS wash removed 95% of cell-associated GTF activity, but









reduced cell-associated FTF by only 34%. No FTF activity was dissociated
from the cell surface by Tween 80 at any concentration, whereas cell-
associated GTF activity was reduced 57% by 0.06% Tween 80, 71% by
0.04% Tween, and 81% by 0.02% Tween.
It became important to determine if there was genetic induction
following the addition of sucrose that might account for the ultimate
increase in total GTF and FTF activities over time.

Effect of antibiotics on GTF and FTF production.

An inhibition experiment similar to that described for dextranase and
fructanase (Table 2-1) was performed for the glycosyltransferases. The
untreated control and rifampicin-, or chloramphenicol-treated cells
exhibited levels of cell-associated GTF which were less than those of the
extracellular form of this enzyme (Table 2-2). The total GTF activity in
response to each of these treatments was very similar, although rifampicin
treatment caused an unexpected release of GTF activity into the fluid phase.
Upon the addition of sucrose, extracellular activity remained constant while
cell-associated activity increased almost fourfold. The total GTF activity in
the presence of sucrose was twofold higher than in the absence of sucrose.
Transcriptional and translational inhibitors, in conjunction with sucrose;
however, decreased the amount of extracellular GTF activity by 69-71%
when compared with untreated control and sucrose-exposed cultures. The
effects of rifampicin and chloramphenicol may be due to the inhibition of
all cellular protein synthesis.
The cell-associated form of FTF predominated in galactose-grown,
and in rifampicin- and chloramphenicol-treated cells; however, upon the









addition of sucrose, extracellular FTF activity increased almost sevenfold
while cell-associated activity remained unchanged (Table 2-3). When
chloramphenicol or rifampicin was added together with sucrose,
extracellular FTF levels remained constant, but cell-associated FTF activity
decreased sharply (71-73%). Total FTF activity was highest when sucrose
was added without chloramphenicol or rifampicin. Taken in aggregate,
these data indicate both genetic and posttranslational regulation of FTF by
its substrate sucrose.

Discussion

In studies such as those presented above, differential enzyme
production due to growth phase variations or enzyme degradation in aging
cultures must be avoided. Growth experiments; therefore, were run
extensively to determine the optimum stage at which cells grew at a
consistent rate without exhausting nutrients. This made it possible to
normalize all experiments by growing the cells to a predetermined point in
mid-exponential phase regardless of further treatment. The use of single
sugars as sole carbon sources in a chemically-defined medium minimized
the effects of nutrition which could complicate interpretation of results.
Fortunately, S. salivarius PC-1 utilized galactose at a rate similar to that of
sucrose and other sugars. This allowed the use of galactose as a "non-
sucrose related" carbon-source for the sucrose shift experiments. The pH
levels (6.1-6.4) of S. salivarius PC-1 cultures grown to an optical density
of Klett 75, regardless of the sugar used, were well within the ranges
determined by others for optimal GTF (McCabe & Smith, 1973),









dextranase (Walker et al., 1980), FTF (Wenham et al., 1979), and
fructanase (Jacques et al., 1985b) production.
Cell preparation and treatment also were designed to be consistent
within all experiments. This uniform approach provided cells at equivalent
mass, growth phase, and concentration and allowed for the comparison of
cell-associated and extracellular enzymatic activities under different
experimental conditions. The enzymatic activities measured were taken to
be an indication of the relative amounts of enzyme produced. The products
assayed in the GTF and FTF experiments were the net result of synthesis
(by GTF and FTF) and also degradation (by dextranase and fructanase).
On the other hand, the substrate for GTF and FTF (sucrose) was not
present in the dextranase and fructanase assays and therefore could not
have contributed to net amount of product.
Chassy et al. (1976) found that the presence of sucrose reduced GTF
activity in culture fluids of S. salivarius, but did not see a shift to the cell-
associated form. These investigators were unable to distinguish between
the possibilities of repression of GTF by sucrose or some modification in
activity of constitutive GTF levels. In the present study, when sucrose was
added to growing cells, GTF activity became associated with the cell
surface (Figure 2-6a). The nature of this surface interaction is different
from that observed with FTF, since GTF activity, but not FTF activity, can
be washed from the surface with SDS and low concentrations of Tween 80.
Recently, Aduse-Opoku et al. (1989), have shown that the glucan-
binding protein of Streptococcus mutans becomes bound to the cell surface
on exposure to sucrose. If dextran were to form a bridge between the
glucan binding domain of GTFs (Mooser & Wong, 1988) and these
surface-bound glucan-binding lectins (Drake et al., 1988; Banas et al.,









1990), washing with detergents may solubilize the dextran, releasing the
GTF activity. McCabe and Smith (1973) reached similar conclusions
concerning the binding of the GTFs of S. mutans. Another theory,
proposed by others (Umesaki et al., 1977; Wittenberger et al., 1978;
Jacques et al., 1985a) is that GTF synthesis and release are regulated by
changes in the membrane fluidity determined by its fatty acid composition.
The gtfB and gtfC genes from S. mutans have been cloned (Aoki et
al., 1986; Pucci et al., 1987; Hanada & Kuramitsu, 1988) and sequenced
(Shiroza et al., 1987; Ueda et al., 1988). The nucleotide sequences
designated the gene products as extracellular enzymes in that putative signal
sequences were present and membrane anchor sequences were absent. This
further supports the conclusion that cell-association of GTF activity is
mediated by factors other than wall- or membrane-anchoring domains in
the GTF molecule and that cell-associated and extracellular activities may
be mediated by the same enzyme species.
Inhibitor studies (Table 2-2) and studies by other investigators (Janda
& Kuramitsu, 1976; Montville et al., 1977; Janda & Kuramitsu, 1978)
indicate that de novo protein synthesis may be required for the synthesis of
at least one extracellular GTF. Recently, Hudson and Curtiss (1990)
showed an increase in the expression of gtfB/C from S. mutans in the
presence of sucrose, thereby providing additional evidence of genetic
regulation of this enzyme by its substrate in the oral streptococci. This
production, followed by association of GTF activity to whole cells, may be
part of a dynamic process involving glucan-binding lectins or still-
unknown surface proteins. The changes in GTF activity seen upon the
addition of sucrose are difficult to define biochemically due to multiple
effects this molecule may have on the regulation (genetic induction),









location (cell-association), and measureable activity (antagonistic effects of
dextranase) of this enzyme. Furthermore, each of these phenomena may
differ for the individual GTFs in S. salivarius as recently described by
Pitty et al. (1989). However, data presented here and by others (Janda &
Kuramitsu, 1976; Montville et al., 1977; Janda & Kuramitsu, 1978; Hudson
& Curtiss, 1990) suggest that sucrose regulates the synthesis and
distribution of GTF and that these two phenomena are independent of each
other.
The gene encoding the S. mutans GS-5 FTF has been cloned (Sato &
Kuramitsu, 1986) and sequenced (Shiroza & Kuramitsu, 1988). All
indications have been that this enzyme, like GTF, is a secreted protein.
Yet, Figure 2-6b and Table 2-3 clearly show that in the absence of sucrose
the majority of FTF activity is found on the cell surface of S. salivarius, as
previously demonstrated by Chassy et al. (1976). Several hypotheses
regarding the location and activity of FTF have been formulated. It has
been postulated that the cell-associated form of FTF is inactivated by
proteolytic degradation (Jacques & Wittenberger, 1981), that FTF may be
associated with the cytoplasmic membrane (Jacques, 1985), and that the
lipid content of the membrane is responsible for the regulation and release
of this enzyme (Pitty & Jacques, 1987). However, the suggestion can be
made that the rapid release of this activity from the cell surface upon the
addition of sucrose as seen in Figure 2-6b and Table 2-3, and recently
confirmed by Milward and Jacques (1990), is due to the FTF's greater
affinity for sucrose relative to its affinity for the surface framework to
which it may be bound. This establishes a biological basis for the
translocation phenomenon and also suggests that the distribution of FTF is
regulated independently of its synthesis. Since the cell-associated FTF









activity cannot be washed from the cell surface with either ionic or non-
ionic detergents it would argue against the enzyme being held through
nonspecific hydrophilic or hydrophobic interactions, but would strongly
suggest a more specific form of binding.
While a significant amount of the FTF activity can be recovered in
the supernatant after the addition of sucrose, this does not fully account for
the twofold increase in total activity (Table 2-3). Hudson and Curtiss
(1990) have shown that in S. mutans such an effect may be due to increased
transcriptional levels. Nucleotide sequence analysis has revealed regions of
the S. mutans FTF gene which may be involved in its genetic regulation
(Shiroza & Kuramitsu, 1988). Streptococcus salivarius also may regulate
FTF production by substrate induction, but the sucrose-mediated release of
FTF shown above clearly requires a surface-localized binding site for FTF.
It is presently unclear whether this specialized binding protein must be
synthesized and transported to the cell surface along with the FTF.
However, data presented in Table 2-3 indicate de novo protein synthesis is
associated, in some way, with the induction of cell-bound FTF by sucrose.
Streptococcus salivarius dextranase activity resides in the
supernatant, away from the cell, and this activity increases at least 100-fold
in the presence of sucrose. This is most likely not a genetic induction since
neither chloramphenicol nor rifampicin affects dextranase production in
the presence of sucrose. This should be compared with the effects of these
inhibitors on fructanase (Table 2-1) where there is significant inhibition,
indicating possible genetic induction. Sucrose may activate an otherwise
inactive form of dextranase through some type of conformational change.
However, since recombinant dextranase is active in the absence of sucrose
(see Chapter 3), sucrose may act to release dextranase from a putative









dextranase inhibitor. Hamelik and McCabe (1982) have shown that in
Streptococcus sobrinus the majority of dextranase is in a tightly bound,
inactive, enzyme-inhibitor complex. An affector (presumably sucrose)
with a high affinity for the inhibitor would be required for activation of
dextranase.
Dextranase could act on the dextran polymer to provide primer or
branch points for GTF (Walker, 1972; Germaine et al., 1977). If
involvement in glucan formation is a major function of dextranase, the
ratio of a-(l,3) to a-(1,6) glycosidic linkages and therefore the solubility
properties of dextran made by S. salivarius and other organisms may be
altered. This would have a potential impact on colonization, plaque
formation and caries development. Activation of dextranase by sucrose in
order to modify dextran synthesis, therefore, is readily understandable.
Alternatively, dextranase could act at the beginning of a catabolic
pathway resulting in the breakdown of dextran to eventually yield glucose
(Hamada et al., 1975b; Schachtele et al., 1975). It is known that
extracellular dextranase acts as an endoenzyme, cleaving the dextran into
smaller polymers, predominantly isomaltosaccharides (Dewar & Walker,
1975). A regulatory role for sucrose in the catabolic function of
dextranase is less obvious. However, from an energetic standpoint, the
synthesis and degradation of glucans relies solely on bond energy. The
short isomaltosaccharides released by the extracellular endodextranase
should be readily transported across the cell membrane where an
intracellular exodextranase, such as those recently cloned from S. mutans
by Burne et al. (1986) and Russell and Ferretti (1990) would further
degrade these short polymers to glucose. If this energy efficient system for
obtaining glucose exists in S. salivarius, the extracellular endodextranase









could play a key catabolic role in the physiology of this species and
regulation by sucrose would be understandable.
Furthermore, in this catabolic capacity, dextranase would be acting
as an antagonist to the extracellular GTFs, thus explaining the apparent low
levels of GTF activity in this organism. In fact, the presence of dextranase
may obscure any conclusions made in studies of GTF in any organism
producing both these enzymatic activities and consequently, any reaction
influenced by dextran (aggregation, adherence, the distribution of GTFs
between cell-surfaces and culture supematants) (Walker et al., 1981).





















180
160
140
m 120
100
80o
y 60
40
20

0 100 200 300 400 500
Time (minutes)




Figure 2-1. Carbon source utilization by S. salivarius PC-1
Cells were grown in CDM/glucose (10 mM) and aliquots transferred
to either: CDM (open squares), CDM/glucosamine (closed squares),
CDM/galactose (open diamonds), CDM/sucrose (dotted squares),
CDM/fructose (closed diamonds), and CDM/glucose (solid dotted squares).
Each sugar concentration was 10 mM, cells were incubated at 370C, and
Klett readings were taken as shown.






















2, 8

6*



U._
o 2-

0
SGlucose Fructose Galactose Sucrose
e
a Carbon source






Figure 2-2. Effect of carbon source on dextranase and fructanase
production by S. salivarius PC-1
Cells were grown in CDM/glucose (10 mM) and transferred to CDM
containing either; glucose, fructose, galactose, or sucrose (10 mM each).
When cultures reached Klett 75, supernatants were assayed for dextranase
(solid bars) and fructanase (hatched bars). One unit of enzyme activity (U)
was defined as the amount of dextranase or fructanase catalyzing the
release of 1 tmol reducing sugar (glucose or fructose) min-1 ml-1. Each
bar represents the mean + SD of triplicates from a representative
experiment.





















>, 15.0

o 12.5

S 10.0

o 7.5

5.0
0
2.5

U 0.0 -
0 10 20 30 40 50 60 70
a Time (minutes)






Figure 2-3. Effect of sucrose shift on production of dextranase and
fructanase in S. salivarius PC-1
Sucrose (10 mM) was added to cells grown in CDM/galactose (10
mM) to Klett 75. Aliquots were removed at time intervals from
suspensions held at 37oC. Supematants were assayed for dextranase (open
squares) and fructanase (closed squares) activities. One unit of enzyme
activity (U) was defined as the amount of dextranase catalyzing the release
of 1 gmol reducing sugar (glucose or fructose) min-1 ml-1. Each data
point shows the mean + SD of triplicates from a representative experiment.
















20


15
E

> 10


5-



Glucose Fructose Galactose Sucrose
Carbon source



Figure 2-4. Effect of carbon source on cell-associated and extracellular
GTF production in S. salivarius PC-1
Cells were grown in CDM/glucose (10 mM) and transferred to CDM
containing either; glucose, fructose, galactose, or sucrose (10 mM each).
Cultures were harvested at Klett 75 after growth at 370C and cell-
associated (solid bars) and extracellular (hatched bars) GTF activities were
assayed. One unit of enzyme activity (U) was defined as the amount of
GTF that catalyzed the incorporation of 1 nmol of the glucose moiety of
labeled sucrose min-1 ml-1. Each bar represents the mean + SD of
triplicates from a representative experiment.



















60

50
E
g 40

S 30

20
U-
I-
I. 10
10 -

Glucose Fructose Galactose Sucrose
Carbon source

Figure 2-5. Effect of carbon source on cell-associated and extracellular
FTF production in S. salivarius PC-1
Cells were grown in CDM/glucose (10 mM) and transferred to CDM
containing either; glucose, fructose, galactose, or sucrose (10 mM each).
Cultures were harvested at Klett 75 after growth at 370C and cell-
associated (solid bars) and extracellular (hatched bars) FTF activities were
assayed. One unit of enzyme activity (U) was defined as the amount of
FTF that catalyzed the incorporation of 1 nmol of the fructose moiety of
labeled sucrose min-1 ml-1. Each bar represents the mean + SD of
triplicates from a representative experiment.










a


4


3


> 2


0 1 1

0
0 10 20 30 40 50 60 70
Time (minutes)
b


100

_- 80
E
60

40-
o 40

I--
U. 20

0
0 10 20 30 40 50 60 70
Time (minutes)
Figure 2-6. Effect of sucrose shift on production and distribution of GTF
and FTF in S. salivarius PC-1.
Sucrose (10 mM) was added to S. salivarius PC-1 grown in
CDM/galactose (10 mM) at 370C to Klett 75. Aliquots were removed at
timed intervals as shown and cell-associated (open squares) and
extracellular (closed squares) glycosyltransferase activities were
determined. (A) represents GTF activity, (B) represents FTF activity.
One unit of enzyme activity (U) was defined as the amount of GTF or FTF
that catalyzed the incorporation of 1 nmol of the glucose or fructose
moiety, respectively, of labeled sucrose min-1 ml-i. Each data point shows
the mean + SD of triplicates from a representative experiment.










Table 2-1. Effect of Antibiotics on Dextranase and Fructanase Production
Enzyme Activity (U)
Treatment Dextranase Fructanase


None 0.16 + 0.03 6.58 + 0
St 10.67 + 0 11.72 + 0.47
C* 0.10 + 0.01 6.27 + 0.12
S&C 9.89 + 0.64 9.68 + 0.60
Rt 0.10 + 0.03 6.98 + 0.38
S&R 10.43 + 0.25 8.76 0.23


ts = Sucrose
*C = Chloramphenicol
SR = Rifampicin

A S. salivarius PC-1 CDM/galactose culture was grown to mid-exponential
phase (Klett 75). Sucrose (10 mM), chloramphenicol (100 mg ml-1),
rifampicin (100 mg ml-1), or combinations of these were added to aliquots
of the culture and then incubated at 370C for 1 hour. Control cultures
contained none of these reagents. The supematants were assayed for
extracellular dextranase and fructanase activities. One unit of enzyme
activity (U) was defined as the amount of dextranase or fructanase
catalyzing the release of 1 .Cmol reducing sugar min-1 ml-1.












Effect of Antibiotics on n


GTF Activity (mU)


Treatment


Cell-Associated


Extracellular


0.74 + 0.13
3.52 + 0.10
0.34 + 0.09
3.21 + 0.04
0.43 0.03
3.31 + 0.07


2.95 + 0.84
3.43 + 0.12
2.86 + 0.06
1.08 + 0.24
5.44 + 0.23
1.00 + 0.02


tS = Sucrose
*C = Chloramphenicol
:R = Rifampicin

A S. salivarius PC-1 CDM/galactose culture was grown to mid-exponential
phase (Klett 75). Sucrose (10 mM), chloramphenicol (100 mg ml-1),
rifampicin (100 mg ml-1), or combinations of these were added to aliquots
of the culture and then incubated at 370C for 1 hour. Control cultures
contained none of these reagents. The cells and supematants were assayed
for GTF activity. One unit of enzyme activity (U) was defined as the
amount of GTF that catalyzed the incorporation of 1 nmol of the glucose
moiety of labeled sucrose min-1 ml-1.


None


Total


S&C
Rt
S&R


3.69
6.95
3.20
4.29
5.87
4.31


T!;hli-~~~~~~~~, 99 EfcofA tboiso lcwtaseaeP dcin


-UI A I


Table 2-2












Table 2-3. Effect of Antibiotics on Fructosyltransferase Production
FTF Activity (mU)
Treatment Cell-Associated Extracellular Total


None 31.44 + 0.46 5.06 + 0.01 36.50
st 40.31 + 1.76 34.38 + 0.25 74.69
C* 35.76 + 1.94 5.01 + 0.01 40.77
S&C 11.52 + 0.03 30.77 + 0.15 42.29
Rt 47.28 + 0.26 4.03 + 0.09 51.31
S&R 10.89 + 0.01 26.67 + 1.91 37.56


tS = Sucrose
*C = Chloramphenicol
tR = Rifampicin

A S. salivarius PC-1 CDM/galactose culture was grown to mid-exponential
(Klett 75). Sucrose (10 mM), chloramphenicol (100 mg ml-1), rifampicin
(100 mg ml-1), or combinations of these were added to aliquots of the
culture and then incubated at 370C for 1 hour. Control cultures contained
none of these reagents. The supernatants were assayed for FTF activity.
One unit of enzyme activity (U) was defined as the amount of FTF that
catalyzed the incorporation of 1 nmol of the fructose moiety of labeled
sucrose min-1 ml-1.














CHAPTER 3
THE EXTRACELLULAR ENDODEXTRANASE OF STREPTOCOCCUS
SALIVARIUS: MOLECULAR CLONING AND STUDIES OF ENZYME
REGULATION


Introduction

Streptococcus salivarius, although not a major oral pathogen, is a
common inhabitant of the oral cavity (Weerkamp & McBride, 1980). This
species is among the earliest colonizers of the human mouth after birth and
constitutes about 40-60% of the streptococci in saliva and on the tongue
dorsum (Hamada & Slade, 1980). Its presence in large numbers, and
therefore its potential impact on the oral ecology (Staat et al., 1982), make
the study of this organism and the extracellular enzymes it produces, e.g.
glycosyltransferases, fructanase, and dextranase (Chassy et al., 1976;
Takahashi, et al., 1983; Houck et al., 1987) of considerable importance.
Understanding the regulation and control of these enzymes which are
involved in sucrose metabolism will be necessary for a better
understanding of the role of this organism in the oral cavity.
Until recently, these enzymes had been assumed to be constitutive, as
they are produced in the absence of substrate (Janda & Kuramitsu, 1978;
Wenham et al., 1979). However, it now has been shown that sucrose plays
a role in the regulation of its metabolism in S. salivarius and that
significant versatility is maintained in the mechanisms by which this is
accomplished (Townsend-Lawman & Bleiweis, 1991). Preliminary work









has shown that this organism uses a variety of mechanisms to control this
set of enzymes, allowing the regulation of their expression and function at
several widely separated stages in their production. Among these,
dextranase (ao-1,6-glucan hydrolase, EC 3.2.1.11) appears to be the only
enzyme controlled primarily at the posttranslational level. Dextranase
activity increased immediately upon the addition of sucrose to galactose-
grown cells, a phenomenon which was not affected by inhibitors of
transcription (rifampicin) or translation (chloramphenicol). These results
suggest that the increase in dextranase activity, in response to sucrose, may
involve the displacement of a dextranase inhibitor (Townsend-Lawman &
Bleiweis, 1990). This chapter describes experiments which further suggest
the presence of such an inhibitor, its displacement by sucrose, and also
indicate other levels of posttranslational control of this enzyme. Genetic
and biochemical approaches have been taken in order to study the
coordination of these regulatory events and the biological significance of
dextranase to this organism.
Methods

Bacterial strains and bacteriophages.

Escherichia coli XL1-Blue: endAI, hsdR17 (rk-, mk+), supE44, thi-
1, lambda-, recAl, gyrA96, relAl, (lac-) [F', proAB, laclIZA M15, TnlO
(tetR )] was purchased from Stratagene. XL1-Blue was maintained on
Luria-Bertani (LB)/tetracycline (12.5 gtg ml-1) (Sigma Chemical Co.) agar
plates and routinely grown in LB medium (Maniatis et al., 1982). To
prepare the XL1-Blue host cells for all manipulations described in this
manuscript a single colony was inoculated into LB broth supplemented with









0.2% maltose and 10 mM MgSO4. The cultures were grown overnight at
370C with vigorous shaking, centrifuged at 1,000g for 10 minutes, and
resuspended in 0.5 volumes of 10 mM MgSO4. Streptococcus salivarius
PC-1, a fresh isolate (Townsend-Lawman & Bleiweis,1991), was grown in
chemically defined medium (CDM) (Terleckyj et al., 1975) and stored at -
700C in 25% (v/v) glycerol. Streptococcus salivarius PC-1 CDM culture
supernatants were prepared by removing the cells by centrifugation
(4,000g, 200C, 5 minutes) followed by filtration using a 0.2 gpm membrane
filter (Gelman Sciences, Inc.). The culture supernatants were concentrated
using CentriprepTM concentrators (Amicon) and dialyzed overnight at 40C
in potassium phosphate buffer, pH 6.35.

Isolation of S. salivarius genomic DNA.

Chromosomal DNA was isolated from S. salivarius PC-1 as follows:
PC-1 cells were grown in 200 ml CDM from an overnight inoculum in the
same medium to an O.D.600 of 0.6. The culture was centrifuged for 10
minutes at 10,000g. Cells were washed once with ice-cold 2 M NaCl (100
ml) and once with ice-cold dH20 (100 ml) and centrifuged for 10 minutes
at 10,000g. The cell pellet was resuspended in 4.5 ml glucose-Tris (GT)
buffer (20 mM Tris, pH 7.0, and 20% glucose) containing 0.5 mg
mutanolysin (Sigma Chemical Co.) and incubated at 370C for 1 hour. One
ml sucrose-Tris-EDTA (STE) buffer (1% sucrose, 100 mM Tris, pH 8.0,
and 200 mM EDTA, pH 8.0) was added and incubated at 370C for 15
minutes. The reaction mixture was chilled briefly on ice and 1.5 ml 5 M
sodium percholate was added. The lysate was extracted with 7.5 ml
phenol/chloroform (1:1), shaken gently for 10 minutes, and centrifuged at









5,000g for 10 minutes. This extraction step was repeated 4 times. Genomic
DNA was precipitated by mixing 2 volumes of ice-cold 95% ethanol with
the aqueous solution, freezing at -700C for 20 minutes and centrifuging at
10,000g for 10 minutes at 4oC. The DNA pellet was washed once with
70% ethanol, air dried, and dissolved in 1 ml Tris-EDTA (TE) buffer (10
mM Tris, pH 7.4 and 1 mM EDTA, pH 8.0). RNase (Sigma Chemical Co.)
was added to a final concentration of 100 gg ml-1 and incubated at 370C
for 1 hour. Proteinase K (Sigma Chemical Co.) was added to a final
concentration of 200 gg ml-1 and incubated at 370C for 2 hours. This
solution was extracted once with an equal volume of chloroform. The DNA
was precipitated with ethanol and resuspended in 1 ml TE.

Construction of the S. salivarius genomic library.

The S. salivarius chromosomal DNA was partially digested with Eco
R1 (Promega), and ligated to Eco Rl-cut and dephosphorylated Lambda
ZAP II vector arms (Stratagene) using T4 DNA ligase (IBI). After in vitro
packaging using a packaging extract from Stratagene, the genomic library
was generated by infecting XL1-Blue indicator cells. To test the quality of
the packaged ligation product, samples (1-500 pl) of the packaged reaction
were plated with 200 pl XL1-Blue cells. Phage and bacteria were
preincubated at 370C for 15 minutes. Three milliliters of NZ Amine
(NZY) top agar (0.5% NaC1, 0.2% MgSO4.7 H20, 0.5 % yeast extract,
and 1% casein hydrolysate) containing 50 pl of 0.5 M isopropyl-P-thio-
galactopyranoside (IPTG, in H20) and 50 pl of 125 mg ml-1 5-bromo-4-
chloro-3-indoyl-P-d-galactopyranoside (X-gal) in N,N-dimethylformamide
(DMF) were added to phage and bacteria and plated onto NZY agar plates.








Screening the S. salivarius genomic library for dextranase clones.

In order to isolate dextranase clones, recombinant phage from the
genomic library were infected into XL1-Blue indicator cells, mixed with 8
ml soft agar containing NZY medium and 1% blue dextran (Sigma
Chemical Co.) to give approximately 1400 plaques per 150 mm petri dish
and overlaid onto M9 agar (Maniatis et al., 1982). Dextranase clones were
identified as plaques surrounded by zones of clearing on the blue dextran
agar (Barrett et al., 1987) and were plaque purified for future use (Figure
3-la). Recombinant phage expressing dextranase activity appeared at a
frequency of 1.2 x 10-3.

Preparation of E. coli lysates and S. salivarius supernatants expressing
dextranase activity.

Dex phage lysates were prepared as follows: Dextranase-positive
plaques were cored from the blue dextran agar plate and transferred to a
sterile microfuge tube containing 500 gl SM (0.58% NaCI, 0.2% MgSO4,
50 mM Tris, pH 7.5, and 0.01% gelatin) and 20 g1 chloroform. One
milliliter of this phage stock was infected into 5 ml XL1-Blue cells and
incubated at 370C for 15 minutes. LB medium (1 L) containing 0.2%
maltose and 10 mM MgSO4 was added and the culture incubated at 370C
for 12 hours with vigorous shaking. Aliquots of these lysates and in
separate experiments, PC-1 CDM culture supematants, were precipitated
with ammonium sulfate. The bacterial debris was pelleted (8,000g, 45
minutes, 40C) and the supernatant filtered with 47 mm metrigard superfine
prefilters (Gelman Instrument Co.) and sterilized with 1000 ml NalgeneM
sterilization filter units, Type S, CN (Nalge Co.). Sodium azide, to 0.02%,









was added along with 430 g L-1 solid ammonium sulfate (Fisher Scientific
Co.). These solutions were stirred at 4oC for 72 hours. The resulting
precipitate was collected by centrifugation (9,000g, 4oC, 4 hours),
resuspended in a small volume of potassium phosphate buffer, pH 6.35,
dialyzed overnight with frequent changes of the same buffer, and stored at
4oC until use. The phage lysates and supernatants were monitored
periodically for dextranase activity by radial diffusion from wells cut in a
1% agarose gel matrix containing 0.2% blue dextran and 50 mM potassium
phosphate buffer, pH 6.35. Activity was visualized as zones of clearing
surrounding the wells.

Protein analysis of native and recombinant dextranases.

Proteins were separated by sodium dodecyl sulfate (SDS)-
polyacrylamide gel electrophoresis on 7.5% gels by using the discontinuous
buffer system of Laemmli, (1970). High range prestained SDS-PAGE
standards (Bio-Rad Laboratories) were run simultaneously in order to
determine approximate molecular weights) of the PC-1 and recombinant
proteins. SDS has been shown to inhibit dextranase activity (Barrett &
Curtiss, 1986). To circumvent this problem, the proteins from PC-1 CDM
culture supernatant (Centriprep-concentrated and ammonium sulfate-
precipitated) and dex phage lysates (unconcentrated and ammonium sulfate-
precipitated) were transferred onto nitrocellulose membranes at 200 mA
for 1 hour in the buffer of Towbin et al., (1979). The blots were laid over
a 1% agarose gel matrix containing 0.4% blue dextran and 50 mM
potassium phosphate buffer, pH 6.35, and incubated at 370C overnight.
The proteins were able to renature sufficiently such that dextranase activity









was visualized as clear bands surrounded by blue color caused by the
infiltration of blue dextran from the gel to the nitrocellulose filter.

Collection of polysaccharides from S. salivarius recombinants.

Recombinant clones encoding sucrolytic enzymes were selected from
the S. salivarius genomic library by their ability to produce polysaccharide
on M9 agar containing 1% sucrose (Gilpin et al., 1985). Since E. coli
XL1-Blue cannot utilize sucrose as a carbon source, NZY soft agar was
used to give a weak bacterial lawn. Plaques surrounded by substantial
growth suggested the presence of cloned S. salivarius sucrolytic enzymes.
These plaques appeared at a frequency of 3.3 x 10-3. One third of these
plaques produced globules of polysaccharide when incubated for longer
periods of time (Figure 3-1b). Ten such plaques (recombinants PG1 to
PG10) were plaque-purified and related on XL1-Blue. The plates were
incubated at room temperature for 2 weeks. The polysaccharide was
scraped from the surface of the agar and dissolved in water (20 ml). After
pelleting the agar and cell debris by centrifugation (4,000g, 200C, 5
minutes), two volumes of ice-cold 95% ethanol were added to precipitate
the polysaccharide, which was spooled out with a capillary tube. The
polysaccharide was resuspended in water (20 ml) and extracted with
ethanol at least twice before being air-dried and weighed.

Determination of the sugar component in purified polvsaccharides.

In order to determine whether the gene encoding glucosyltransferase
(gtf) or fructosyltransferase (ftf) had been cloned and which type of
polymer (glucan or fructan) was being produced, the sugar components)









of the polysaccharides was determined by thin layer chromatography
(TLC) following acid hydrolysis. Polysaccharides from the recombinants
(20 mg), levan (20 mg), and dextran Type 100C (20 mg), were boiled
(1000C, 1 hour) under N2 in sealed glass ampules with 2 N HC1. Two
microliters samples of these reactions, along with 2 pl (fructose, sucrose,
and glucose) (1 M) as standards, were loaded onto a dry silica gel plate
which had been equilibrated overnight with the separation solvent
(chloroform-acetic acid-water, 3.0: 3.5: 0.5, v/v). After separation in the
solvent, the sugars were visualized by spraying the plate with 10 ml of 1%
diphenylamine (wt/v), 1% analine (v/v), acetone mixed with 1 ml 85%
phosphoric acid, and heating (1300C, 10 minutes) (DeStefanis & Ponte,
1968). Rf values of each spot were calculated as a ratio of the distance
between the origin and the center of the spot to the distance between the
origin and the solvent front. Crude recombinant lysates also were assayed
for GTF and FTF activity by incorporation of radioactivity from
differentially labeled sucrose (Robrish et al., 1972).

Determination of dextranase activity.

Dextranase activity was determined using a standard assay for
reducing sugars designed by Somogyi (1951) and Nelson (1944). Dextran
Type 100C (Sigma Chemical Co.) was incubated with the enzyme-
containing preparation at a final concentration of 0.25 mg ml-1, unless
specified otherwise. The reactions were stopped after 180 minutes. The
amount of reducing sugar released was determined by comparison with
glucose standards. One unit of enzyme activity (U) was defined as the
amount of dextranase catalyzing the release of 1 lmol glucose min-1 ml-1.









Each data point represents the mean (+/- SD) of triplicates from a
representative experiment.

Carbon source utilization by S. salivarius PC-1.

Growth curves were generated by inoculating triplicates of CDM or
CDM/substrate with a CDM/glucose (10 mM) starter culture at a 1:10
ratio. Substrates employed as carbon sources included glucose, sucrose,
dextran Type 100C, dextran Type 500C, dextran Type 2000C, isomaltose
(IM2), isomaltotriose (IM3), isomaltotetraose (IM4), isomaltopentaose
(IM5) (Sigma Chemical Co.), and glucan obtained from S. salivarius gtf
recombinant PG10, at final concentrations of 1 mg ml-1. Optical density
measurements were taken with a Klett-Summerson photoelectric
colorimeter (Filter No. 54) every 30 minutes until stationary phase was
reached. The pH of each culture was determined at the same time intervals
using ColorpHastT indicator sticks (MC/B Manufacturing Chemists, Inc.).

Substrate specificity of native and recombinant dextranases and product
analysis.

Tenfold concentrated S. salivarius PC-1 CDM/sucrose (10 mM)
culture supernatant and ammonium sulfate precipitated dex phage PD1
lysate were incubated with blue dextran, dextran Type 100C, dextran Type
500C, dextran Type 2000C, or glucan obtained from S. salivarius gtf
recombinant PG10 at final concentrations of 0.25 mg ml-1, for 180
minutes. Dextranase activity was determined as above. Duplicate reactions
were incubated overnight and analyzed by TLC, using the same solvent
system as described previously, to determine the products of native and









recombinant dextranase activity. Standards used were glucose and the
oligosaccharides of the isomaltose series (IM2, IM3, IM4, IM5).

Sucrose-mediated release of dextranase inhibition in PC-1 cell-free
superatant.

To compare the levels of dextranase activity in mid-exponential S.
salivarius PC-1 cultures grown with or without sucrose, tenfold
concentrated PC-1 culture supematants (CDM-galactose or sucrose, 10
mM) were incubated with dextran Type 100C, at a final concentration of
0.25 mg ml-1 and assayed for dextranase activity at 15 minute intervals.
To confirm that the increase in dextranase activity in response to sucrose
occurred after dextranase entered the extracellular environment, sucrose at
several concentrations (0-14 mM), was added to sterile, concentrated
culture supernatant from S. salivarius PC-1 cells grown in CDM with 10
mM galactose and incubated at 370C for 1 hour. Dextranase activity in this
supernatant was quantitated by measuring the amount of reducing sugar
released from dextran Type 100C.

Effects of untreated and sucrose-treated cell-free S. salivarius supematants
on recombinant dextranase activity.

Dialyzed, concentrated PC-1 CDM/galactose (10 mM) culture
supernatant was prepared as above. Sucrose (10 mM) was added to an
aliquot of the CDM/galactose supernatant and incubated at 370C for 1
hour. These supematants, CDM/galactose and CDM/galactose plus sucrose,
were added in 0, 10, 20, 30, 40, and 50% proportions to a constant
amount (25 gl) of ammonium sulfate precipitated recombinant dex lysate
(PD1). Controls with 100% supernatant and no recombinant enzyme were









included to provide estimates of background activity. Dextran Type 100C
was added to a final concentration of 0.25 mg ml-1 and incubated for 180
minutes at 370C. Dextranase activity was determined by measuring the
amount of reducing sugar released. Substrate concentration and enzyme
activity (U) were such that the limiting component was substrate.
Therefore, whether the entire reaction mixture (50 l.1) consisted of the
enzyme preparation (S. salivarius supernatant or PD1 lysate) or half the
reaction mixture (25 pl enzyme preparation, 25 pl 50 mM potassium
phosphate buffer, pH 6.35), measurable dextranase activity was the same
due to limiting dextran.

Results

Construction and screening of the S. salivarius genomic library.

Streptococcus salivarius DNA was cloned into Lambda ZAP II after
partial digestion with the restriction endonuclease, Eco R1, and ligation
with T4 DNA ligase. The phage library was used to infect E. coli XL1-
Blue which was plated on M9 agar plates containing sucrose. Clones
expressing sucrase activity appeared as rings of E. coli growth surrounding
individual plaques and occurred at a frequency of 3.3 x 10-3. Upon
further incubation, 33% of these sucrolytic clones were able to form domes
of polysaccharide from the sucrose (Figure 3-la). Ten polysaccharide-
positive clones (PG1 to PG10) expressed GTF, but not FTF, activity when
assayed with differentially labeled sucrose in the standard
glycosyltransferase assay (Lawman & Bleiweis, unpublished results). In
addition, the polysaccharides produced by these clones were hydrolyzed
with 2N HC1 and analyzed by TLC, and found to contain only glucose









(Lawman & Bleiweis, unpublished results). The glucan from PG10 was
used later in this study to test the substrate specificity of native and
recombinant dextranases. The S. salivarius library transduced into XL1-
Blue also was plated on blue dextran (Figure 3-lb). Clones able to clear
zones by hydrolyzing the blue dextran appeared at a similar frequency as
GTF-expressing clones (1.2 x 10 -3). One clone expressing dextranase
activity, PD1, was chosen for further investigation and was used to
characterize the dex gene and its product. PD1 was found to carry a 6.3 kb
Eco R1 fragment encoding the dex gene and a promoter, as the clone was
capable of expressing dextranase activity in the absence of IPTG. A more
complete characterization of the dex gene is the subject of Chapter 4. This
chapter focuses on the characteristics of the dex gene product.
Three polypeptides of molecular weights 190, 90, and 70 KD in
unconcentrated PD1 lysates were shown to be responsible for the
dextranase activity of PD1 by SDS-PAGE/electroblot, and blue dextran
overlays (Figure 3-2, lane 3). Upon precipitation with ammonium sulfate,
only the two smaller molecular weight species could be visualized. This
may be the result of proteolysis during the concentration process (Figure
3-2, lane 4). The native dextranase had an apparent molecular weight of
110 KD by this procedure regardless of the method of concentration
(Figure 3-2, lanes 1 and 2). This assay was not sensitive enough to detect
dextranase activity in unconcentrated S. salivarius supematants.

Carbon source utilization.

It was of interest to determine whether S. salivarius could metabolize
either large molecular weight dextrans or the products of endodextranase
hydrolysis, oligoisomaltosaccharides, since it has been postulated that









stored dextran might serve as an energy reservoir for this and other
organisms (Parker & Creamer, 1971; Hamada et al., 1975; Schachtele et
al.,1975b; Ellis & Miller, 1977). Streptococcus salivarius PC-1 was unable
to utilize any of the isomaltosaccharides or dextrans as measured by growth
or shifts in pH (Table 3-1). In fact, PG10 glucan appeared to inhibit S.
salivarius growth in sucrose as evidenced by the longer generation time
(from 57 to 69 minutes) and the lower absorbance at maximal growth (125
to 75 Klett units) when glucan was added to sucrose at 1 mg ml-1.
Melibiose also was tested as a growth substrate. If a melibiose operon were
present in S. salivarius as in Streptococcus mutans (Tao et al., 1990),
melibiose might induce the uptake of oligosaccharides and their utilization
as a fermentable carbon source. Melibiose did not serve as a carbon source
for S. salivarius PC-1, nor did it facilitate the utilization of dextran or
isomaltosaccharides. However, it did extend the lag-time of cells grown in
sucrose and in sucrose/PG10 glucan from 90 to 150 minutes and the lag-
time of cells grown in glucose from 90 to 240 minutes, and decreased the
density at which growth peaked (Table 3-1).

Substrate specificity of native and recombinant dextranases and product
analysis.

Dextranase is thought to hydrolyze accumulated dextrans in dental
plaque and release hexose for consumption by oral microorganisms.
Therefore, it was important to determine the extent to which various
dextrans might serve as substrates for this enzyme and the products of such
reactions. It also was of interest to compare the substrate specificity and
resultant reaction products of the native and recombinant dextranases.
Tenfold concentrated and dialyzed S. salivarius CDM/sucrose culture









supernatant and ammonium sulfate-precipitated PD1 lysate were incubated
with dextrans from various sources and of different molecular weights (S.
salivarius was grown in CDM/sucrose so that the native dextranase would
be in its active form). All dextrans tested served as substrates for both the
native and recombinant enzymes, but to varying degrees (Figure 3-3). The
native dextranase in CDM/sucrose culture supernatant released reducing
sugars from the native PG10 glucan to greater extents, ranging from 18 to
41% than from the other substrates (Figure 3-3a). The recombinant
enzyme, however, was far less active on the native glucan (50%) than was
the native dextranase (Figure 3-3b), although the dex gene product showed
equivalent or greater degrees of activity on the commercial dextrans
(Figure 3-3b). Relative dextranase specific activities (native: recombinant)
were as follows: PG10, 2.22; Blue dextran, 0.99; T100C, 0.52; T500C,
0.38; T2000C, 0.75)
TLC analysis of the reaction products of dextran degradation
revealed that the native form of dextranase was not able to hydrolyze any
of the glucan substrates to the smaller isomaltosaccharides released by the
recombinant enzyme (Figure 3-4). Because TLC was unable to resolve the
oligosaccharides produced by the native enzyme, it is presumed these
products were greater than 7 glucosyl residues in length. Neither form of
dextranase released free glucose.

Sucrose-mediated release of dextranase inhibition in S. salivarius PC-1
cell-free supernatants.

In order to demonstrate further the disparity of dextranase activity
between S. salivarius grown in CDM/galactose vs. CDM/sucrose
(Townsend-Lawman & Bleiweis, 1991), supernatants from mid-exponential









cultures were concentrated, dialyzed, and assayed for dextranase activity at
15 minute intervals. When S. salivarius was grown in CDM with galactose,
dextranase activity was minimal (Figure 3-5a), however, when cells were
grown in the presence of sucrose, the dextranase was active (Figure 3-5a),
even after the putative "activator" (sucrose) had been removed by extensive
dialysis. Maximal activity required 180 minutes incubation with substrate,
although significant activity was measurable earlier. To confirm that
sucrose-dependent increases in dextranase activity were not cell-mediated,
sucrose, at several concentrations was added to sterile, concentrated S.
salivarius CDM/galactose culture supernatant, incubated for 1 hour, and
assayed for dextranase activity. Sucrose was able to "activate" the
otherwise inactive dextranase in a concentration-dependent manner, with an
optimum at 10 mM disaccharide (Compare Figure 3-5b with 3-5a). This
provided further evidence that the sucrose effect was initiated
postgenetically.

Effects of untreated and sucrose-treated cell-free S. salivarius supernatants
on recombinant dextranase activity.

To determine whether the increase in dextranase activity upon the
addition of sucrose was due to activation or release from inhibition, cell-
free S. salivarius CDM/galactose supernatant, or CDM/galactose
supernatant plus sucrose was added, at various concentrations, to PD1
lysate. CDM/galactose supernatant inhibited the recombinant dextranase
activity in a concentration-dependent manner (Figure 3-6a). This
confirmed the presence of a dextranase inhibitor in the CDM/galactose
supernatant and implied that it was present in excess of the native
dextranase, since 37% was the maximum inhibition obtained using 50%









CDM/galactose supernatant. On the other hand, the dextranase activity in
CDM/galactose supernatant which had been treated with 10 mM sucrose
was additive to recombinant dextranase activity (Figure 3-6b), suggesting
that both native and recombinant forms of the enzyme were able to utilize
the same substrate molecules.

Discussion

All indications are that dextranase has the ability to participate in
dental plaque formation and modification through its effects (degradation
and/or synthesis) (Staat & Schachtele, 1974; Hamada et al., 1975) on the
capsular polysaccharides, which are thought to be important in bacterial
aggregation and adherence (Gibbons & Van Houte, 1975; Schachtele et al.,
1975a). However, the biological significance of this enzyme and its role(s)
in sucrose metabolism have not been firmly established. Dextranase may
act directly in conjunction with GTFs in the synthesis and modification of
dextran polymers, by providing primer and/or branchpoints, and thus may
be involved in determining the degree of dextran solubility (Walker, 1972;
Germaine et al., 1977), a property shown to be an important factor in
colonization, plaque formation and caries development (Walker & Jacques,
1987). Alternatively, dextranase may break down the accumulated dextran
in plaque, destabilizing the plaque matrix, and at the same time provide
fermentable hexose for consumption (Parker & Creamer, 1971; Hamada et
al., 1975; Schachtele et al., 1975b Ellis & Miller, 1977) and acid
production (Wood, 1969) by plaque organisms. In this capacity
extracellular endodextranase may act at the beginning of a catabolic
pathway in a synergetic manner with an intracellular exodextranase, as









found in S. mutans (Burne et al., 1986; Russell & Ferretti, 1990), to
hydrolyze short term dextran reserves rapidly to glucose (Dewar &
Walker, 1975). Studies were initiated, therefore, to determine the major
function of endodextranase, either anabolic or catabolic, and its
contribution to the physiology of S. salivarius and potential contribution to
other inhabitants of the oral cavity.
Most information to date on dextranase in oral streptococci has been
obtained from Streptococcus sobrinus (Germaine & Schachtele, 1976; Ellis
& Miller, 1977; Walker et al., 1981; Barrett et al., 1987). Purification of
this enzyme has proven difficult due to multiple molecular weight forms
attributable to enzyme aggregation and/or protease modification. Barrett
et al. (1987) developed a purification scheme for dextranase from S.
sobrinus culture supernatant fluids in which the majority of dextranase was
recovered in two forms, molecular weights 175 KD and 160 KD. Lower
molecular weight forms, ranging from 160 KD to 125 KD, were thought
to be proteolytic breakdown products of the 175 KD dextranase. When
these investigators screened S. sobrinus gene libraries for dextranase
activity, three phenotypes of recombinant clones were identified. The
authors believe that it was unlikely that the variation in molecular weight
was due to the cloning of two separate genes since genetic characterizations
of S. sobrinus dextranase mutants suggested a single dex gene.
Reported in this chapter is the cloning of the endodextranase from S.
salivarius PC-1. Dextranase-positive clones were recovered at a frequency
of 1.2 x 10-3. One such clone, designated PD1, was chosen for further
study and was found to carry a 6.3 kb Eco RI fragment encoding the dex
gene. It was deduced that this insert contained a promoter since the clone









was capable of expressing dextranase activity in the absence of IPTG, and
enough of the dex structural gene to produce a protein with dextranase
activity of at least 190 KD (see Figure 3-2). This protein was degraded
into lower molecular weight species (90 and 70 KD) which retained the
ability to hydrolyze blue dextran as did the S. sobrinus dextranase (Barrett
et al., 1987). The native dextranase; however, was recovered as a single
110 KD polypeptide. The fact that the native protein appeared to be
smaller than the recombinant, may be due to proteolytic degradation by
streptococcal enzymes or may represent a monomeric form of this enzyme.
At this point the possibility that the recombinant enzyme may be a fusion
protein cannot be ruled out.
Preliminary results of studies on the regulation of sucrose
metabolism in S. salivarius were presented above. In these studies it was
shown that extracellular dextranase activity was higher in sucrose-grown S.
salivarius PC-1 cells compared to cells grown in glucose, fructose, or
galactose; that dextranase activity increased 100-fold when sucrose was
added to cells growing in galactose; and that this immediate increase was
not affected by transcriptional or translational inhibitors. Ellis and Miller
(1977) found a similar sucrose effect on dextranase activity in S. mutans
6715 (S. sobrinus) supernatants. These results suggested that the increase
in the activity of native dextranase in response to sucrose may involve the
displacement of a dextranase inhibitor and implied that de novo synthesis
was not required for the production of dextranase.
Hamelik and McCabe (1982) concluded that the presence of an
inhibitor in batch-grown culture fluids of S. mutans accounted for the
absence of endodextranase activity in strains known to produce this









enzyme. To investigate further the regulation of dextranase by sucrose and
the presence of a dextranase inhibitor in S. salivarius, CDM/galactose and
CDM/sucrose culture fluids were assayed for their ability to release
reducing sugars from dextran. The CDM/galactose supernatant showed
little dextranase activity while the CDM/sucrose supernatant showed
increasing activity over time (see Figure 3-5a). "Activation" was a
posttranslational effect since dextranase activity increased in a
concentration-dependent manner when sucrose was added to cell-free S.
salivarius CDM/galactose supernatant (see Figure 3-5b).
It is clear from Figure 3-2 that native dextranase was active in the
absence of sucrose when separated from other proteins in S. salivarius
supernatants. Also, the recombinant dextranase was active in the absence
of sucrose (Figures 3-1, 3-2), which argues against the need for an
activator. To distinguish between activation and relief of inhibition,
CDM/galactose supernatant was added in increasing concentrations to
recombinant dextranase preparations. Figure 3-6a demonstrates the
presence of a factor in S. salivarius CDM/galactose supernatant which
inhibited recombinant dextranase activity. This inhibitor apparently was in
excess of a 1:1 molar ratio to the native dextranase, since it was available
for inhibition of the recombinant enzyme. If sucrose acts to relieve this
inhibition, presumably by replacing dextranase as the ligand for the
dextranse inhibitor, one would expect sucrose-treated supernatants to be
non-inhibitory to the recombinant dextranase. Not only was the effect non-
inhibitory (see Figure 3-6b), there was a synergetic effect, i.e., dextranase
activity, in the presence of a constant amount of substrate, increased to
twice that of either native or recombinant alone. This implied that both









forms of dextranase (native and recombinant) were able to release reducing
sugars from the same substrate molecules. It was not clear; however, if the
two enzymes released similar or different end-products when assayed with
dextran substrates.
To compare enzyme activities, S. salivarius CDM/sucrose culture
supernatant and ammonium sulfate-precipitated PD1 lysate were incubated
with various dextrans. The native and recombinant enzymes were able to
recognize all the dextrans tested, but varied in their recognition and/or
hydrolysis of dextran substrates. The native dextranase hydrolyzed
homologous glucan more successfully than dextrans from other sources,
while the recombinant enzyme hydrolyzed heterologous (commercial)
glucans to a greater degree than streptococcal capsular dextran (see Figure
3-3). The recombinant enzyme released the products expected from an
endodextranase (from isomaltose to larger isomaltosaccharides, see Figure
3-4). On the other hand, the S. salivarius supernatant, which had been
shown to release comparable levels of reducing activity (see Figure 3-3),
did not hydrolyze any of the dextran substrates to small oligosaccharides or
glucose (see Figure 3-4). It is thus likely that there was a factor present in
the S. salivarius supernatant which regulated or directed the recognition,
specificity and/or function of the native dextranase beyond its regulation
by the dextranase inhibitor. Perhaps this "controlling" factor, possibly
GTF itself, binds or otherwise modifies dextranase to limit and/or direct
the degradation of the dextran to allow production of appropriately-sized
primers or to pin-point insertion of branchpoints. Construction of
isogenic dex- mutants would allow clarification of this vital functional
activity.









Since S. salivarius PC-1 is unable to utilize dextrans,
isomaltosaccharides, or melibiose as sole carbon sources (see Table 3-1),
and since these substances may in fact inhibit growth, this organism
probably does not utilize a synergetic pathway involving extracellular
endodextranase, isomaltosaccharide transport factors, and intracellular
exodextranase to metabolize stored capsular polysaccharides for energy.
Ellis and Miller (1977) made similar conclusions from their work with S.
mutans 6715. Induction or activation of dextranase by sucrose, but not
dextran, would be an inefficient system if dextran was unable to serve as a
fermentable carbohydrate. It also could be suggested that the melibiose
transport operon found in S. mutans, enabling the utilization of melibiose
and isomaltosaccharides may be missing in this strain of S. salivarius.
Therefore, the major biological role of endodextranase in this organism
appears to be to augment the activity of GTF(s) in a synthetic capacity.
It has been shown through genetic and biochemical means that the
endodextranase of S. salivarius PC-1 acts as a component of the synthetic
machinery designed to construct extracellular polysaccharides, and that its
activity is directed by some factor present in the extracellular environment,
perhaps GTF itself. This makes the positive regulation of dextranase
activity by sucrose, the substrate for dextran formation by GTF(s),
understandable. These findings suggest that the regulation of sucrose
metabolism and the enzymes involved is more complex than once imagined.
Implicit in these conclusions is the importance of immediate availability of
critical enzymes on the cell's exterior to meet the extremely erratic signals
from the environment (e.g. nutrient availability/deprivation). The
extracellular availability of dextranase and certain glycosyltransferases may






60


be controlled to a great extent by posttranslational means rather than
genetic regulation in this organism.



























b












Figure 3-1. Screening the S. salivarius genomic library for dex and gtf
recombinants.
(a) Recombinants from the S. salivarius library were selected for their
ability to produce polysaccharide on M9 agar containing 1% sucrose. A
plaque-purified gtf recombinant (PG10) exhibits the characteristic
production of glucan from sucrose.
(b) Aliquots of the S. salivarius genomic library were mixed with NZY
soft agar containing 1% blue dextran and overlaid onto M9 agar.
Dextranase-positive (dex) clones were identified as plaques surrounded by
a zone of clearing and plaque-purified for further study.
















-.-190


-11 0


--".~I1
i:IilB I
!as S s a a! ,af.S..: ..: .. ... ... .. ilL i^B '


Figure 3-2. Detection of electroblotted dextranase activity on blue
dextran-agarose.
Proteins were separated on a 7.5% SDS-polyacrylamide gel and
transferred onto nitrocellulose membranes. The blots were laid over a 1%
agarose gel matrix containing 0.4% blue dextran and 50 mM potassium
phosphate buffer, pH 6.35 and incubated overnight at 370C. Dextranase
activity was visualized as clear bands on the nitrocellulose membranes
surrounded by blue color left by the infiltration of blue dextran from the
gel. Lanes: 1) Centriprep-concentrated PC-1 CDM/sucrose culture
supernatant, 2) ammonium sulfate-precipitated PC-1 CDM/sucrose culture
supernatant, 3) unconcentrated dex phage lysate (PD1), 4) ammonium
sulfate-precipitated dex phage lysate (PD1).







63

a


23



4 2




0
I -Bo






z i Substrate

b --
x














E 0
0 0



















S- Substrate
0 0 0




4.

U

4-


x -











Figure 3-3. Substrate specificity of native and recombinant dextranases.
E 0 -, -




(a) Tenfold concentrated PC- CDM/sucrose (10 mM) culture
supernatant and (b) ammonium sulfate-precipitated dex phage lysate were
incubated with glucan obtained from S. salivarius recombinant PG0,
.,- I- 0
a ~I--


Figure 3-3. Substrate specificity of native and recombinant dextranases.
(a) Tenfold concentrated PC-1 CDM/sucrose (10 mM) culture
supernatant and (b) ammonium sulfate-precipitated dex phage lysate were
incubated with glucan obtained from S. salivarius gtf recombinant PG10,
blue dextran, dextran Type 100C, dextran Type 500C, and dextran Type
2000C at final concentrations of 0.25 mg ml-1. Dextranase activity was
determined by the amount of reducing sugar released. One unit of enzyme
activity (U) was defined as the amount of dextranase catalyzing the release
of 1 gunol glucose min-1 ml-1.

















-A





Ii













Figure 3-4. Product analysis of native and recombinant dextranases.
The products of the following reactions were separated by thin layer
chromatography. Dialyzed, tenfold concentrated PC-1 CDM/sucrose (10
mM) culture supernatant (Native) and ammonium sulfate-precipitated PD1
dex phage lysate (Recombinant) were incubated with (1) no substrate, (2)
glucan obtained from S. salivarius PC-1 gtf recombinant PG10, (3) blue
dextran, (4) dextran Type 100C, (5) dextran Type 500C, and (6) dextran
Type 2000C (at a final concentration of 10 mg ml-1) overnight at 370C.
Standards include: G-glucose; IM2-isomaltose; IM3-isomaltotriose; IM4-
isomaltotetraose; IM5-isomaltopentaose.







65
a


12

10
E
.. 8 -

46



o 2-

0 0 1 T I
0 30 60 90 120 150 180 210
Time (Minutes)
b


8


6


< 4


0
2



0 2 4 6 8 10 12 14
Sucrose (mM)
Figure 3-5. Sucrose-mediated release of dextranase inhibition in PC-1 cell-
free supernatant.
(a) Tenfold concentrated culture supernatants from PC-1 grown in
CDM with galactose (closed diamonds) or sucrose (open squares) were
assayed for dextranase activity at 15 minute intervals. (b) Sucrose (0-14
mM) was added to sterile, concentrated culture supernatant from S.
salivarius PC-1 cells grown in CDM with 10 mM galactose and incubated
at 370C for 1 hour. Dextranase activity was quantitated by measuring the
amount of reducing sugar released from dextran Type 100C. One unit of
enzyme activity (U) was defined as the amount of dextranase catalyzing the
release of 1 p.mol reducing sugar min-1 ml-1. Assays were performed in
triplicate.








a

6

5
4
o
3

: 2

x
0
o 0
100 50 40 30 20 10 0
CDM/Galactose Supernatant (%)
b


10

8

*o 6

e 4



01
100 50 40 30 20 10 0
CDM/Galactose Supernatant + Sucrose (%)

Figure 3-6. Effects of CDM/galactose and CDM/galactose plus sucrose
cell-free S. salivarius PC-1 supernatants on recombinant dextranase
activity.
(a) Tenfold concentrated PC-1 CDM/galactose (10 mM) culture
supernatant and (b) tenfold concentrated PC-1 CDM/galactose (10 mM)
culture supernatant incubated for 1 hour with sucrose (10 mM) were added
in 0, 10, 20, 30, 40, and 50% proportions to ammonium sulfate-
precipitated dex phage lysate. One hundred percent culture supernatants
indicated the background levels of native enzyme in "inactive"
(CDM/galactose) and "active" (CDM/galactose plus sucrose) supernatants.
Dextran Type 100C was added to final a concentration of 0.25 mg ml-1
and incubated for 180 minutes at 370C. Dextranase activity was
determined by the amount of reducing sugar released. One unit of enzyme
activity (U) was defined as the amount of dextranase catalyzing the release
of 1 gmol glucose min-lml-1.













Table 3-1. Carbon source utilization


I blnr n ma


Lag
time
r i n


Generation
time
t(mrti


Peak
Absorbance
(T Kltt un;tc


Glucose
Sucrose
Melibiose
Melibiose + Glucose
Melibiose + Sucrose
PG10
PG10 + Sucrose
PG10 + Melibiose
PG10 + Suc + Mel
*Dextrans or IMs
(+/- Melibiose)


110
125


90
90
no growth
240
150
no growth
90
no growth
150
no growth


Streptococcus salivarius PC-1 cultures containing various carbon sources (1
mg ml-1) or combinations thereof (1 mg ml-1 each) were monitored by
Klett-Summerson colorimetric analysis for growth. Culture pH also was
measured.

Glucan from the S. salivarius gtf recombinant PG10
* Blue dextran, dextran Type 100C, dextran Type 500C, dextran Type
2000C, IMs= isomaltosaccharides; IM2-isomaltose; IM3-isomaltotriose;
IM4-isomaltotetraose; IM5 -isomaltopentaose


Final
nH-I


by S. salivarius PC-1.


ra 0on sorc mI nv ILp1













CHAPTER 4
ANALYSIS OF THE EXTRACELLULAR ENDODEXTRANASE GENE
OF STREPTOCOCCUS SALIVARIUS

Introduction

Sucrose metabolism in the oral streptococci has been the subject of
numerous studies, primarily because it is believed to be the major
contributory factor in plaque formation and the subsequent development of
dental caries (Drummer & Green, 1980; Hamada & Slade, 1980; Drucker
et al., 1984). The metabolism of sucrose, in its most basic form, involves
the translocation of sucrose or its component hexoses (Slee & Tanzer,
1979), concomitant phosphorylation, hydrolysis, and utilization for energy
(Saier, 1989). In a broad sense, however, it also includes the synthesis of
extracellular polysaccharides, both glucans and fructans, catalyzed by
glucosyltransferase (GTF) and fructosyltransferase (FTF) enzymes,
respectively. These polymers are, in turn, degraded by dextranase (a-1,6-
glucan hydrolase, EC 3.2.1.11), which has been shown to behave as an
endoenzyme, breaking glucans down to isomaltosaccharides (Walker, 1972;
Lawman & Bleiweis, submitted, Journal of General Microbiology, 1991),
and fructanase (3-D-fructan fructohydrolase, EC 3.2.1.80), which
hydrolyzes fructans to fructose (Burne et al., 1987).
While sucrose metabolism has been the topic of much discussion as it
relates to the potential virulence of oral streptococci, little attention has
been focused on its potential to serve as a model for studying the complex









regulatory network of enzymes necessary for cellular response to irregular
signals from the environment. Streptococcus salivarius, an unremarkable
pathogen is; however, an adept colonizer of the human mouth and can
constitute up to 60% of the streptococci in saliva and on the dorsal side of
the tongue (Hamada & Slade, 1980). It possesses the entire array of
enzymes, eg. glycosyltransferases, fructanase, and dextranase (Chassy et
al., 1976; Takahashi et al., 1983; Houck et al., 1987; Townsend-Lawman &
Bleiweis, 1991), necessary for the production and degradation of the
extracellular polysaccharides derived from sucrose and has been shown to
regulate these enzymes in an extremely complex fashion (Townsend-
Lawman & Bleiweis, 1991; Lawman & Bleiweis, submitted, Journal of
General Microbiology, 1991). For these reasons, S. salivarius is the ideal
species on which to base a model for the control of extracelluar enzymes in
Gram positive organisms. Evidence that the availability of dextranase and
certain glycosyltransferases is controlled to a great extent by
posttranslational, rather than genetic regulation (Townsend-Lawman &
Bleiweis, 1991; Lawman & Bleiweis, submitted, Journal of General
Microbiology, 1991), lays the foundation for further studies and calls for
indepth genetic studies of each enzyme involved.
The present chapter focuses on the analysis of the endodextranase of
S. salivarius at a molecular level and includes the confirmation of the
cloned dextranase activity, its molecular weight, the uniqueness of this
sequence in the S. salivarius genome and a molecular map of the DNA
which encodes it.








Methods

Bacterial strains and growth media.


Streptococcus salivarius PC-1, a fresh isolate (Townsend-Lawman &
Bleiweis, 1991) was grown in chemically defined medium (CDM)
(Terleckyj et al., 1975) and stored at -700 in 25% (v/v) glycerol. A
genomic library was constructed as described previously (Lawman &
Bleiweis, submitted, Journal of General Microbiology, 1991). Briefly, S.
salivarius PC-1 chromosomal DNA was partially digested with Eco R1
(Promega) and ligated to Eco Rl-cut and dephosphorylated Lambda ZAP
II vector arms (Stratagene) using T4 DNA ligase (IBI). The packaged
recombinant bacteriophage particles were transfected into Escherichia coli
XL1-Blue cells : endAI, hsdR17 (rk-, mk+), supE44, thi-1, lambda-,
recAl, gyrA96, relAl, (lac-) [F', proAB, laclZA M15, TnlO (tetR)]
(Stratagene). XL1-Blue was maintained on Luria-Bertani (LB)/tetracycline
(12.5 gg ml-1) (Sigma Chemical Co.) agar plates and routinely grown in
LB medium (Maniatis et al., 1982). Dextranase clones were identified as
plaques surrounded by zones of clearing when plated in top agar containing
blue dextran (Barrett et al., 1987). A recombinant clone expressing
dextranase activity, PD1, was selected for further study and stored in SM
buffer (0.58% NaC1, 0.2% MgSO4, 50 mM Tris, ph 7.5, and 0.1% gelatin)
and chloroform. Clone PD1, was shown to carry a promoter and express a
190 KD polypeptide which exhibited dextranase activity by SDS-
PAGE/electroblot, and blue dextran overlay (Lawman & Bleiweis,
submitted, Journal of General Microbiology, 1991).








In vitro translation of PD1.

In vitro translation experiments were conducted using the
prokaryotic DNA-directed translation kit from Amersham and L-[35S]
methionine (9.4 x 10-4 mM, 0.75 gCi gl-1) (Amersham). This kit was used
according to manufacturer's specifications and involved the addition of E.
coli RNA polymerase, a mixture of unlabeled amino acids (without
methionine), L-[35S]-methionine, template DNA (either Lambda ZAP II
DNA or PD1 DNA) and an E. coli cell extract. Proteins were separated by
sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis on 10%,
0.77 mm gels, at 25 mA using the discontinuous buffer system of Laemmli
(1970). High range prestained SDS-PAGE standards (Bio-Rad
Laboratories) were run simultaneously in order to determine the
approximate molecular weights) of the recombinant protein(s). The gel
was then fixed in 25% methanol (v/v) and 10% acetic acid (v/v), washed in
dH20 for 20 minutes, then in 1 M sodium salicylate for 20 minutes. After
a brief rinse in dH2 0 the gel was wrapped in plastic wrap and overlayed
with X-ray film for 48 hours at room temperature before developing.

DNA manipulations, chemicals and enzymes.

Recombinant phage DNA of clone PD1 was prepared by a protocol
for the large-scale preparation of bacteriophage lambda and subsequent
extraction of DNA as described by Maniatis et al. (1982). DNA analysis
was performed by horizontal agarose (SeaKem) gel (0.7% wt/v)
electrophoresis in TAE buffer, pH 8.6 (40 mM Tris acetate, 1 mM EDTA)
at 15V cm-1 of gel and visualized by the fluorescence of ethidium bromide.
The restriction endonucleases; Ssp I, Xmn I, Xba I, Not I, Aat II, Sac I,









Sau I, Sac II, Pvu II, Bal I, Bgl I, and 10X universal restriction
endonuclease buffer were obtained from Stratagene. T4 DNA ligase and
the restriction endonucleases; Tth III 1, Bam HI, Pst I, and Xho I were
obtained from IBI. Pvu I, Hinc II, Nar I, the nick translation kit, and
PhotoGene nucleic acid detection systems were purchased from BRL,
Dra I, Sty I, Nhe I, Hind III, Nsi I, Eco RI, and Eco RV were obtained
from Promega. Biotin-labeled dATP was incorporated by nick translation
into DNAs used as probes in the Southern hybridization analysis (either
pPD13 or the 3.7 kb Xho I fragment of PD1) and were separated on PD-10
Sephadex G-25 M columns (Pharmacia). Streptococcus salivarius
chromosomal DNA was extracted as previously described (Lawman &
Bleiweis, submitted, Journal of General Microbiology, 1991), digested with
restriction endonucleases, and separated by agarose gel electrophoresis, see
above. After depurination in 0.25 M HC1 (15 minutes), denaturation in 1.5
M NaCl and 0.5 M NaOH (twice for 20 minutes), and neutralization in 1.5
M NaCl and 1.0 M Tris-HCL, pH 7.5 (20 minutes), these DNAs were
electroblotted onto PhotoGeneT nylon membranes (BRL) for 1 hour at
200 mA in 0.1X TAE buffer, pH 8.6. Biotinylated Hind III-cut lambda
molecular weight markers (BRL) were run simultaneously. The Southern
blots were prehybridized, hybridized with biotinylated probe DNA, and
developed with PhotoGeneT nonradioactive detection system according to
specifications from BRL. The blots were exposed to the PhotoGeneT
substrate (15-60 minutes) and autoradiographed at room temperature (10
secconds-7 minutes) to obtain the desired results. All DNA manipulations
using these products were conducted under the conditions suggested by the
manufacturers.








Subcloning the dex gene.

In vivo excision and recircularization of the cloned inserts in the
pBluescript plasmid were performed using R408 or VCSM13 as the fl
helper phages. Dextranase-positive plaques were cored from the blue
dextran agar plate and transferred to a sterile microfuge tube containing
500 pl SM buffer and 20 pl chloroform. Each phage stock (200 pl) was
incubated with 200 p1l XL1-Blue cells and 1 L.1 helper phage (1 x 1011 pfu
ml-1). Five milliliters of 2X YT broth (1% NaCI, 1% yeast extract, and
1.6% bacto-tryptone) was added and incubated 3 hours at 370C with
shaking. The tube was heated at 700C for 20 minutes. An aliquot of the
rescued phagemid stock (10 p.1) was incubated with 200 p XL1-Blue cells
at 370C for 15 minutes. Infected XL1-Blue cells (50 p ) were plated on
LB/ampicillin (50 gpg ml-1) (Sigma Chemical Co.) and incubated overnight
at 370C. Broth cultures of single colonies from the R408 excisions were
tested for dextranase activity by placing samples in wells cut in a 1%
agarose gel matrix containing 0.2% blue dextran and 50 mM potassium
phosphate buffer, pH 6.35. The VCSM13 excisions produced no colonies.
The 6.3 kb insert of PD1 was excised with Eco RI, isolated by
agarose gel electrophoresis, recovered by freezing and centrifugation in
Microfilterfuge"T tubes with 0.45 pgm nylon membrane filters (Rainin
Instrument Co.), and ligated into Eco RI-cut pDL278 vector with T4 DNA
ligase. The resultant recombinant plasmids were transformed into
competent XL1-Blue cells following the procedure outlined by Maniatis et
al. (1982). The transformation mixture was plated onto LB agar
containing spectinomycin (50 p.g ml-1), 0.05 M isopropyl-P-thio-
galactopyranoside (IPTG), and 5-bromo-4-chloro-3-indoyl-3-d-









galactopyranoside (X-gal), in N,N-dimethylformamide (DMF) (75 jig ml-
1). Plasmid DNA was extracted from the resulting white colonies by the
boiling method supplied by Stratagene, digested with Xba I, and analyzed
by agarose gel electrophoresis.
The 6.3 kb insert of PD1 containing the dex gene also was excised
with Xho I and Not I, producing a 2.6 kb Xho I/Not I fragment and a 3.7
kb Xho I fragment. These fragments were subcloned into Xho I/Not I and
Xho I-cut pBluescript vector (Stratagene), respectively. The ligation,
transformation, and DNA extraction procedures used were as described
above, except that ampicillin (50 pgg ml-1) was used for the selection of
pBluescript. Large plasmid isolations of the successfully subcloned 2.6 kb
Xho I/Not I fragment, pPD13, were prepared according to Maniatis et al.
(1982) and purified by cesium chloride-ethidium bromide gradient-
centrifugation in a VTi50 rotor at 45,000 rpm for 48 hours in a Beckman
L7-55 Ultracentrifuge.

Results

Analysis of the dex gene product.

Previously (Lawman & Beiweis, submitted, Journal of General
Microbiology, 1991), three polypeptides of molecular weights 190, 90, and
70 KD were shown to be responsible for the dextranase activity of PD1 by
SDS-PAGE/electroblot and blue dextran overlays. Since it was not known
whether the two lower molecular weight species were products of
translation or proteolytic breakdown, an in vitro DNA-directed translation
of PD1 was performed. The proteins encoded by PD1 were labeled with
L-[35S] methionine, analyzed by SDS-PAGE, and compared to similarly-









labeled Lambda ZAP II proteins. A major protein, 190 KD, was the
immediate translation product synthesized by the recombinant (Figure 4-1).
Neither the 90 nor the 70 KD polypeptide could be visualized in the in
vitro translation of PD1. This experiment was performed in the absence
of IPTG.

Subcloning the dex gene.

The Lambda ZAP II vector was designed to allow in vivo excision
and recircularization to form a phagemid containing the pBluescript
plasmid and the cloned insert. Broth cultures of single colonies were tested
for dextranase activity (Figure 4-2b) and compared with activity in the
original phage lysate before excision (Figure 4-2a). Dextranase activity
was visualized by a zone of clearing surrounding the well. Unfortunately,
the R408 helper phage could not be removed (Lawman & Bleiweis,
unpublished results). When VCSM13 was used as the helper phage,
excision was complete, but the insert containing the dex gene proved to be
lethal to the E. coli host. Effort was made to subclone the entire 6.3 kb
fragment into pDL278, an E. coli/Streptococcus shuttle vector (Dunny et
al., in press, Genetics and Molecular Biology of Streptococci, Enterococci
and Lactococci, 1991) which was kindly provided by Dr. Don LeBlanc.
However, use of this vector was unsuccessful. Attempts to subclone the 3.7
kb Xho I PD1 fragment also were unsuccessful, presumably resulting in
lethality to E. coli. The 2.6 kb Xho I/Not I fragment; however, was
subcloned into Xho I/Not I-cut pBluescript, where it was maintained stably.
This construct, pPD13, was digested with numerous restriction
endonucleases; the resultant map is shown in Figure 4-3. A deduced
restriction map of PD1 is shown in Figure 4-4. Restriction endonucleases









which failed to cut within the 2.6 kb Xho I/Not I fragment were: Eco RI,
Bal I, Aat II, Sac II, Xmn I, Dra III, Bgl I, Xba I, Nar I, Nsi I, Tth 1 1,
Sau I, and Not I.

Southern blot analysis.

Streptococcus salivarius PC-1 chromosomal DNA was completely
digested with Xho I, Xho I and Eco RI, or Xba I (Figure 4-5a, lanes 2, 3,
and 4, respectively). These digested DNAs were separated by agarose gel
electrophoresis along with the appropriate controls; Xho I/Not I-cut pPD13
(Figure 4-5a, lane 5), Xho I/Not I-PD1 (Figure 4-5a, lane 6), and Eco RI-
cut PD1 (Figure 4-5a, lane 7). These DNAs were blotted onto nylon
membranes and probed with either biotin-labeled pPD13 (Figure 4-5b) or
the 3.7 kb Xho I fragment from PD1 which had been gel-purified and nick
translated to incorporate biotinylated dATP (Figure 4-5c). The pPD13
probe hybridized to a single 10.4 kb Xho I chromosomal fragment (Figure
4-5b, lane 2); a single 2.6 kb Xho I/Eco RI chromosomal fragment (Figure
4-5b, lane 3); a single 10.4 kb Xba I chromosomal fragment (Figure 4-5b,
lane 4); both the 2.6 kb insert and the 3.0 kb pBluescript bands of pPD13
(Figure 4-5b, lane 5); the 2.6 kb fragment of Xho I/Eco RI-cut PD1
(Figure 4-5b, lane 6) from which it was subcloned; and a 6.3 kb fragment
of the original PD1 clone (Figure 4-5b, lane 7). The 3.7 kb probe
hybridized to a single 5.2 kb Xho I chromosomal fragment (Figure 4-5c,
lane 2); a single 3.7 kb Xho I/Eco RI chromosomal fragment (Figure 4-5c,
lane 3); a single 10.4 kb Xba I chromosomal fragment (Figure 4-5c, lane
4); the 3.7 kb Xho I/Not I fragment of PD1 (Figure 4-5c, lane 6); and the
6.3 kb Eco RI fragment of PD1 (Figure 4-5c, lane 7). This probe did not









hybridize to the pPD13 subclone to a significant degree (Figure 4-5c, lane
5).

Discussion

The extracellular endodextranase of S. salivarius has been shown to
have an apparent molecular weight of 110 KD (Lawman & Bleiweis,
submitted, Journal of General Microbiology, 1991). The corresponding
recombinant form was expressed in E. coli as a 190 KD protein along with
two smaller polypeptides (90 and 70 KD) which retained their ability to
hydrolyze high molecular-weight blue dextran. It was not known if the
190 KD protein was the immediate product of translation, a conclusion that
now can be made since a 190 KD protein results from the in vitro DNA-
directed translation of PD1. Furthermore, it can be concluded that the 90
and 70 KD polypeptides were the result of E. coli proteolysis. The
possibility remains; however, that the 190 KD protein may be a fusion
product.
Attempts to subclone the 6.3 kb Eco RI fragment from PD1 and the
3.7 kb Xho I/Eco RI portion of the insert were unsuccessful, resulting in
multiple rearrangements and/or lethality to the E. coli host. Others have
been unable to isolate the RNA-polymerase sense strand of the gtfB gene
(Shiroza et al., 1987), theftf gene (Shiroza & Kuramitsu, 1988), and the
scr B gene (Sato & Kuramitsu, 1988) of S. mutans for sequencing. Each of
these genes contained promoter sequences directly upstream from the
structural gene and were expressed at high levels in E. coli. Clones
containing the dex gene of S. sobrinus also are reported to be unstable (R.
Curtiss III, personal communication). Perhaps the dex gene of S. salivarius









exhibits similar properties since it carries a promoter and can be expressed
at high levels (see Figure 4-1). This also may account for the difficulty in
obtaining ftf clones from the S. salivarius/Lambda ZAP II genomic library
(Lawman & Bleiweis, unpublished results). The hope was that lethality
might be controlled if the dex gene could be subcloned into the pDL278
shuttle vector which had been constructed to minimize the transcription of
DNA inserts by surrounding the multiple cloning site with transcriptional
terminators. This, however, did not seem to rectify the situation.
The 2.6 kb Xho I/Not I fragment was subcloned stably into
pBluescript (see Figure 4-3). Since the S. salivarius insert in PD1 encodes
a 190 KD dextranase, the expected composition of the dex gene should be
approximately 5,100 bp, leaving at least 1,400 bp of the gene residing in
the subclone, pPD13. It will be interesting to see if pPD13 contains the 3'
or 5' sequence of the dex gene.
It has been postulated, based on mutational analyses, that S. sobrinus
carries a single dextranase gene (Curtiss, 1985). To see if the same holds
true for S. salivarius PC-1, chromosomal DNA was digested to completion
with either Xho I, Xho I/Eco RI, or Xba I, and analyzed by Southern
hybridization with both biotin-labeled pPD13 (carrying the 2.6 kb Xho
I/Not I fragment of PD1) and the remaining 3.7 kb Xho I fragment (see
Figure 4-5). It was known that the 6.3 kb Eco RI fragment of PD1 did not
contain an Xba I restriction site (Lawman & Bleiweis, unpublished results);
therefore, a single Xba I fragment of indeterminant size should carry the
entire dex gene. This is the case, since both probes hybridized with a
single fragment of 10.4 kb (see Figure 4-5b, lane 4 and 4-5c, lane 4). The
pPD13 probe hybridized with a single 2.6 kb Xho I/Eco RI chromosomal
fragment (see Figure 4-5b, lane 3), as predicted, and the 3.7 kb probe









hybridized with a single 3.7 kb Xho I/Eco RI chromosomal fragment (see
Figure 4-5c, lane 3). Each probe hybridized to a single Xho I
chromosomal fragment, each of unpredictable length (see lane 2 of Figure
4-5b and c). From these data it can be concluded that the only gene
encoding the endodextranase of S. salivarius PC-1 has been cloned. It also
is useful to know that genes linked immediately upstream and downstream
would be readily available from an Xho I library, since a 10.4 kb Xho I
clone could be selected by hybridization with pPD13 and would contain 7.8
kb of flanking DNA. Concomitantly, a 5.2 kb Xho I clone, selected by
hybridization with the 3.7 kb fragment of PD1, would carry 1.5 kb of
DNA flanking the other end of the dex gene.









































Figure 4-1. Autoradiographs of L-[35S] methionine-labeled proteins from
in vitro translations of Lambda ZAP II and PD1.
Positions of the molecular weight markers are shown on the left. A
major protein band (190 KD) not expressed by the vector Lambda ZAP II
was synthesized by the recombinant.









































Figure 4-2. Excision of pBluescript and dex insert.
XL1-Blue cells were coinfected with PD1 and fl helper phage R408
to produce phagmid constructs. XL1-Blue cells were infected with the
packaged phagemid and plated on LB plates with ampicillin (50 pg ml-1) to
recover the excised plasmid. Broth cultures of single colonies were tested
for dextranase activity (b) and compared with activity in the original PD1
phage lysates before excision (a). Samples were placed in wells cut in a 1%
agarose gel matrix containing 0.2% blue dextran and 50 mM KPO4 buffer,
pH 6.35. Well 1 contains Lambda ZAP II/ no insert; wells 2 and 3, PD1.






82













sty I
Sac 1
Sty I pPD13
5.6 kb













Figure 4-3. Partial restriction map of pPD13 containing the 2.6 kb Xho
I/Not I fragment of PD1.






83






o S. salivarius dex


Terminator


Initiator


_

III I I
-II C4


04 -
U

I c
I i i I
3: vl a Mi

4- M------


11L


S2.6 kb 3.7kb

I I I I3 I I
0 1 2 3 4 5 6 6.3


Figure 4-4. Construction of the original PD1 clone and deduced restriction
map of the 6.3 kb fragment containing the dex gene.
The majority of restriction sites were mapped from the 2.6 kb Xho
I/Not I fragment subcloned in pPD13.


T3
-810-


T7
T7


PD1
46.3 kb


I
I


k

kb







































Figure 4-5. Southern blot analysis of complete restriction digests of the S.
salivarius PC-1 chromosome.
Panel (a) represents the original agarose gel containing the following
DNAs: lane 1, biotinylated Hind III-cut lambda molecular weight markers
(the molecular weight of each marker is indicated to the left of panel (a);
lane 2, Xho I-cut S. salivarius PC-1 chromosomal DNA; lane 3, Xho I/Eco
RI-cut S. salivarius PC-1 chromosomal DNA; lane 4, Xba I-cut S.
salivarius PC-1 chromosomal DNA; lane 5, Xho I/Not I-cut pPD13; lane 6,
Xho I/Not I-cut PD1; and lane 7, Eco RI-cut PD1. Panels (b) and (c)
represent the same DNAs blotted onto nylon membranes and probed with
biotin-labeled pPD13 and the 3.7 kb Xho I fragment from PD1,
respectively.














CHAPTER 5
SUMMARY

The dynamics of host-microbe interactions require adaptive
responses on the part of the bacterium to diverse environmental conditions
of different locations within the host, as well as the transition to and from
an external reservoir (Miller et al., 1989). This is especially true of S.
salivarius, since this organism has developed the ability to colonize a wide
range of mammalian species, can survive in the digestive tract and fecal
matter, and can not only survive in the oral cavity, but outcompete many
other species (Hardie, 1986). To survive encounters with such extremes in
the immediate environment requires a constant sensing of the environment
and a timely set of appropriate responses (Miller et al., 1989).
Gram positive bacteria, including S. salivarius, must by necessity
regulate major carbohydrate pathways differently from Gram negative
organisms, since they have a much thicker cell wall, no outer membrane,
and therefore, no periplasmic space (Volk et al., 1986). Communication
between the inside of the cell and the environmentally exposed exterior
may be more difficult for Gram positive organisms. Therefore, it would
be logical to hypothesize that some proteins destined for the outer surface
of the cell wall might be produced at a baseline level, exported to the
surface, and regulated at the posttranslational level. Such proteins might
include large molecules, anchored to the wall, serving as scaffolds for
extracellular enzyme networks. Streptococcus salivarius certainly has no









paucity of surface appendages (Gibbons et al., 1972) and its extracellular
proteins are difficult to separate from each other and from other structures
such as lipids, teichoic acids, and peptidoglycan (Montville et al., 1977).
Both of these observations would support such a theory.
Sucrose can be considered an external stimulus to which inhabitants
of the oral cavity must respond. Streptococcus salivarius has a number of
extracellular and intracellular enzymes which are designed to metabolize
sucrose, its constituents, and its byproducts (see Figure 1-1). To believe
that these enzymes are not regulated in some manner seems to be untenable,
and yet this was the accepted dogma until the present study began.
Townsend-Lawman and Bleiweis (1991) were the first to show that
sucrose-metabolizing enzymes in this oral Streptococcus are regulated, both
genetically and posttranslationally. That these enzymes, e.g., GTF, FTF,
fructanase, and dextranase are in fact regulated, was an easy hypothesis to
make and yet quite a difficult one to test, since classical genetic techniques
were and are largely unavailable for streptococci.
Since the overall objective of the preliminary physiological study
was to determine whether or not one or more of these enzymes was
differentially produced upon the addition of sucrose, appropriate culture
conditions were developed first. Extensive growth curves were completed
to determine the optimum stage of growth, where cells would be growing
at a constant rate for a period of time, but not long enough to exhaust any
nutrient. In this way growth rate would not be a variable factor in the
production of these enzymes and repetitions of the experiments could be
made. These curves were compared as the sole carbon source was varied
(glucose, fructose, sucrose, galactose, or glucosamine). With this
information, the growth conditions and cell preparations could be









normalized. The cells were filtered, washed, and reconstituted to a tenfold
concentration equivalent with KPO4 buffer. The supernatants were
concentrated, dialyzed, and reconstituted to the same concentration with the
same buffer. This gave cells at equivalent mass, growth phase, and
concentration. Comparisons could then be made between cell-associated
and extracellular enzymatic activities.
From the data presented in Chapter 1, it can be concluded that de
novo synthesis is required for the production of extracellular GTF activity
which, upon the addition of sucrose, becomes associated with the cell
surface. Cell-associated FTF activity appears to require genetic induction
for its production and cell-surface association, but requires sucrose for its
release from the surface framework. Extracellular fructanase was twofold
higher when cells were grown in sucrose than when they were grown in
the other sugars. The increase in fructanase activity occurred within 5
minutes, but was diminished by transcriptional and translational inhibitors.
Extracellular dextranase activity of cells grown in sucrose was tenfold
higher than that of cells grown in glucose, fructose, or galactose.
Dextranase activity increased 100-fold in less than 5 minutes following the
addition of sucrose to galactose-grown cells. The increase in dextranase
activity was affected by neither rifampicin nor chloramphenicol.
Therefore, the production of dextranase activity by S. salivarius PC-1 does
not require de novo synthesis.
Several lines of evidence suggest that dextranase is tightly controlled
by a dextranase inhibitor which can be displaced by sucrose, and by one
other factor, which appears to regulate and/or direct hydrolysis of dextran
substrates. For example, sucrose was necessary for native dextranase
activity and its effect was biochemically-mediated; supernatant from cells









grown in galactose inhibited recombinant dextranase activity, but when
treated with sucrose the native enzyme acted synergetically with the
recombinant enzyme; and recombinant dextranase released
isomaltosaccharides from dextran substrates, while the native enzyme
incompletely hydrolyzed the dextrans to large polymers.
Streptococcus salivarius PC-1 was able to utilize neither the
substrates nor the products of dextran hydrolysis for growth. Therefore,
the major biological role of the endodextranase appears to be the
augmentation of GTF(s) in a synthetic capacity as opposed to being a
catalytic convertor of stored dextrans.
Genetic analysis of the dex gene began as dex clones from a S.
salivarius/Lambda ZAP II genomic library were identified by their ability
to clear zones in blue dextran agar. A 6.3 kb Eco RI fragment from the
genomic clone, PD1, was shown to carry a promoter and a sufficient
portion of the dex structural gene to encode an active 190 KD recombinant
protein. However, the native dextranase was represented by a 110 KD
polypeptide. Russell and Ferretti (1990) found that the endodextranase
activity of S. mutans could be localized in two major bands (120 and 105
KD) with breakdown products as low as 70 KD retaining their dextranase
activity. Southern blot analysis using either end of the 6.3 kb Eco RI
fragment showed that no other S. salivarius PC-1 chromosomal fragment
had sufficient homology to the labeled probes to allow hybridization under
the conditions used. From this it could be concluded that there is a single
endodextranase gene in S. salivarius.
In order to sequence the dex gene it was necessary to amplify the
cloned DNA. Attempts to subclone the 6.3 kb Eco RI insert were
unsuccessful due to rearrangements and lethality to the E. coli host.