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Studies on the phosphoenolpyruvate-dependent phosphotransferase systems in Streptococcus mutans GS5

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Studies on the phosphoenolpyruvate-dependent phosphotransferase systems in Streptococcus mutans GS5
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Liberman, Ellen S., 1950-
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ix, 153 leaves : ill. ; 28 cm.

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Cell growth ( jstor )
Cell membranes ( jstor )
Enzymes ( jstor )
Membrane proteins ( jstor )
Membrane transport proteins ( jstor )
Metabolism ( jstor )
Phosphorylation ( jstor )
Reactants ( jstor )
Repression ( jstor )
Sugars ( jstor )
Dissertations, Academic -- Microbiology -- UF
Microbiology thesis Ph. D
Phosphorylation ( fast )
Phosphotransferases ( fast )
Streptococcus mutans ( fast )
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bibliography ( marcgt )
theses ( marcgt )
non-fiction ( marcgt )

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Thesis:
Thesis (Ph. D.)--University of Florida, 1982.
Bibliography:
Includes bibliographical references (leaves 145-152).
Additional Physical Form:
Also available online.
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Typescript.
General Note:
Vita.
Statement of Responsibility:
by Ellen S. Liberman.

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STUDIES ON THE PHOSPHOENOLPYRUVATE-DEPENDENT
PHOSPHOTRANSFERASE SYSTEMS IN
Streptococcus mutans GS5








BY

ELLEN S. LIBERMAN











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


UNIVERSITY OF FLORIDA 1982












ACKNOWLEDGMENTS


The author wishes to express her gratitude to the chairman of her

committee, Dr. Arnold S. Bleiweis, for his valuable guidance and support. She would also like to thank Drs. Richard Boyce and Francis Davis, for their counsel during these studies. She owes special thanks to Dr. Keelnatham Shanmugam whose help in obtaining mutants and with various

other aspects of this work was invaluable.

She would also like to show her appreciation to Steven F. Hurst for his assistance with gas liquid chromtographic analysis and for his excellent graphics and photography. Lastly, two of the author's co-workers, Dr. Julie Siegel and Ms. Patti Traube-Beede, are given special thanks for their support and encouragement.





















ii














TABLE OF CONTENTS


Page

ACKNOWLEDGMENTS . . . . . . ii


LIST OF TABLES . . . . . . . iv


LIST OF FIGURES . . . . . . vi


ABSTRACT . . . . . . . . Viii


INTRODUCTION AND LITERATURE REVIEW . . 1 MATERIALS AND METHODS . . . . . 39 RESULTS . . . . . . . . 53


CONCLUSIONS . . . . . . . 127


BIBLIOGRAPHY . . . . . . . 145


BIOGRAPHICAL SKETCH . . . . . 153













iii













LIST OF TABLES

Table Page

1 Phosphoenolpyruvate-dependent phosphotransferase systems
in S. mutans . . . . . . . . . . . 26
2 Lactose phosphotransferase system and phospho-6galactosidase enzyme activities as a function of buffer
composition and pH . . . . . . . . . . 59

3 Lactose phosphotransferase system and phospho-0galactosidase activities as a function of ionic strength
of buffer reagent . . . . . . . . . . 60

4 Determination of optimal amounts of solvents for decryptification of cells for phosphotransferase enzyme assays . 61

5 Comparison of sugar (glucose and fructose) transport and
phosphorylation by decryptified or untreated cells . . 63

6 Determination of phosphoenolpyruvate dependence for glucose
phosphotransferase system-mediated phosphorylation . . 67

7 Induction of phosphoenolpyruvate-dependent phosphotransferase systems as a function of carbon source in growthmedia (LDH/NADH-linked assay) . . . . . . . 71

8 Induction of phosphoenolpyruvate-dependent phosphotransferase systems as a function of carbon source in growthmedia (radioactive assay) . . . . . . . . 74

9 Competitive inhibition by unlabelled sugars of uptake of
radiolabelled sugars by S. mutans GS5 . . . . . 75

10 Glucose phosphotransferase system negative mutants:
Group I . . . . . . . . . . . . . 88

11 Glucose phosphotransferase system negative mutants:
Group II . . . . . . . . . . . . . 89

12 Glucose phosphotransferase system negative mutants:
Group III . . . . . . . . . . . . 90

13 Glucose phosphorylation by cell fractions of S. mutans GS5 93

iv









LIST OF TABLES (Continued)

Table Page
14 Phosphoenolpyruvate-dependent phosphorylation of glucose
by mutanolysin-prepared membranes of S. mutans GS5 . . 97

15 Phosphotransferase activities of mutanolysin-prepared
membranes . . . . . . . . . . . . 98

16 Inhibitory effect of competing sugars on the phosphorylation of D-[14C(U)]-glucose by the phosphoenolpyruvatedependent phosphotransferase system of decryptified cells
and mutanolysin-prepared membranes derived from
S. mutans GS5 . . . . . . . . . . . 99

17 Glucose-glucose-6-phosphate exchange reaction (transphosphorylation) catalyzed by mutanolysin-purified membranes
of S. mutans GS5 as a function of reactant concentrations. 101

18 Pyruvate-phosphoenolpyruvate exchange reaction as a probe
for Enzyme I: Distribution of activity in cell-free
extracts of S. mutans GS5 . . . . . . . . 107

19 The relative glucose and fructose phosphotransferase
activities in mutanolysin prepared membranes of mutant
and wild-type strains of S. mutans GS5 . . . . . 108

20 Transphosphorylation as a measure of Enzyme II in mutanolysin-prepared membranes of wild-type and mutant strains
of S. mutans GS5 . . . . . . . . . . 110

21 The distribution of Enzyme I as determined by the phosphoryl exchange reaction between pyruvate and phosphoenolpyruvate in cell-free extracts of wild-type and mutant
strains of S. mutans GS5 . . . . . . . . 112

22 The growth and sugar uptake by cells of S. mutans GS5
induced for lactose dissimilation in medium supplemented
with lactose and glucose . . . . . . . . 116










v












LIST OF FIGURES

Figure Page
1 Schematic of the phosphoenolpyruvate-dependent
phosphotransferase system . . . . . . . . 4

2 Proposed scheme for the regulation of carbohydrate
transport and metabolism by the phosphoenolpyruvatedependent phosphotransferase system in E. coli . . . 17

3 Lactose phosphotransferase system activity as a function
of cell concentration . . . . . . . . . 56

4 The pH optimum for the lactose phosphotransferase system 58

5 Lactose phosphotransferase system enzyme activities and
growth of S. mutans GS5 as a function of time . . . 67
6 Glucose phosphotransferase system activity as a function
of phosphoenolpyruvate (PEP) concentration . . . . 70
7 Kinetics of glucose-6-phosphate formation by the PEPdependent phosphotransferase system of S. mutans GS5 . 79
8 Kinetics of mannose-6-phosphate formation by the phosphoenolpyruvate-dependent phosphotransferase system of
S. mutans GS5 . . . . . . . . . . . 81

9 Kinetics of fructose-6-phosphate formation by the phosphoenolpyruvate-dependent phosphotransferase system of
S. mutans GS5 . . . . . . . . . . . 83

10 Growth of S. mutans GS5 in glucose, fructose, and mannose 85 11 Phosphorylation of glucose by cell-free membranes . . 95

12 Glucose-glucose-6-phosphate transphosphorylation by cellfree membranes . . . . . . . . . . 103

13 Phosphoenolpyruvate-dependent phosphotransferase component
El activity in cell-free membranes . . . . . . 106

14 Diauxic growth patterns by S. mutans GS5 using glucose
and lactose substrates . . . . . . . . . 115
15 Growth patterns displayed by S. mutans GS5 in fructose
plus lactose . . . .. . . . . . . . 119

vi









LIST OF FIGURES (Continued)

Figure Page

16 Growth of S. mutans GS5 (wild-type) and a glucose
phosphotransferase negative mutant in lactose plus
glucose . . . . . . . . . . . . 122

17 Lactose uptake by S. mutans GS5 (wild-type) and a glucose
phosphotransferase negative mutant as a function of cell
density . . . . . . . . . . . . 124



































vii













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

STUDIES ON THE PHOSPHOENOLPYRUVATE-DEPENDENT PHOSPHOTRANSFERASE SYSTEMS IN
Streptococcus mutans GS5

By

Ellen S. Liberman
December 1982

Chairman: Dr. Arnold S. Bleiweis Major Department: Microbiology and Cell Science

This dissertation is concerned with characterization of the glucose phosphoenolpyruvate (PEP)-dependent phosphotransferase system (glc PTS) in Streptococcus mutans GS5. The PTS is a mode of transport which involves the translocation of a phosphoryl group from PEP to an incoming sugar moiety. It is a complex system which requires up to four proteins, Enzyme I (El), HPr, Enzyme III (EIII), and Enzyme II (EII),to accomplish the transport and group translocation functions.

The objectives of the present study were (1) to obtain a general

overview of the glc PTS, (2) to obtain a more detailed picture of the glc PTS by studying isolated membranes derived from cells of S. mutans, and

(3) to study the hierarchy of sugar utilization in S. mutans.
The substrates of the glc PTS in order of declining affinities were

found to be glucose, mannose, and 2-deoxyglucose. The evidence for this finding comes from studies of the competitive effects exerted by the above sugars on the transport of the heterologous sugars, from kinetic studies, and from studies on glc PTS-negative mutants.

viii










Isolated membranes derived from cells grown in glucose were prepared using the muralytic enzyme, mutanolysin. These membranes were able to phosphorylate glucose and mannose when supplied with exogenous PEP. They were also able to phosphorylate glucose when the phosphoryl donor glucose-6-phosphate was used (transphosphorylation), thus demonstrating the presence of a functional EIIglc in cell-free membranes of the wildtype strain. The presence of El in these membrane preparations was demonstrated by the phosphoryl exchange reaction between PEP and pyruvate.

Glucose in the growth-medium prevented the induction by lactose of the lac PTS in S. mutans GS5. Thus, glucose is a preferred sugar. Glucose did not appear to repress the induction of the lac PTS in glc PTS-negative cells even though glucose was taken up by these cells. This finding indicates the necessity for a functioning glc PTS for the regulation of lactose uptake. The mutant cells contained wild-type levels of El but lacked a functioning EIIglc as shown by the two phosphoryl exchange reactions. These results suggest that the E11iglc is required for the regulation of sugar uptake in S. mutans GS5.















ix













INTRODUCTION AND LITERATURE REVIEW


Streptococcus mutans is the causative agent of dental caries, the most prevalent bacterial disease in humans. This species is divided into a number of serotypes (a-g), however serotype c is most commonly isolated from carious lesions. The growth of this organism is accompanied by the production of lactic acid which causes the demineralization of dental enamel. The resulting cavity is the clinical manifestation of this disease (13,14). Numerous investigators have been studying the physiology of sugar metabolism of S. mutans in order to deduce basic mechanisms of transport and dissimilation and to learn about the genetic basis of the pertinent enzymology. It is hoped that such knowledge could be applied to preventative therapy and thus aid in the elimination of this disease.

At this stage only limited knowledge exists concerning the transport of sugars into S. mutans. The purpose of this dissertation is to expand our knowledge of a major transport route, the phosphotransferase system, in serotype c strain GS5.
Mechanisms of transport. Carbohydrate transport in bacteria is

characterized by three general modes. The first is exemplified by the lactose system in Escherichia coli. The energy required for lactose uptake is derived from proton symport, i.e., lactose is co-transported with a proton, thus dissipating the proton gradient produced during electron transport. The second type of transport utilizes adenosine

1






2


5'-triphosphate (ATP) as an energy source and is exemplified by maltose uptake in E. coli. This type of transport is characterized by the requirement for a periplasmic binding protein. One way these two types of sugar accumulations are differentiated is by the loss of transport functions upon osmotic shock in the case of the latter transport system and the retention of such functions in the former system (7). The third mode of transport is characterized by glucose transport in E. coli and lactose transport in Staphylococcus aureus (21). It is termed the phosphoenolpyruvate (PEP)-dependent phosphotransferase system (PTS). This type depends on the energy inherent in the enol configuration of the phosphoryl group of PEP. Since the phosphoryl group is transferred, it falls into the general category of group translocation.

Components of the phosphotransferase system. The two systems

which have been studied in the most detail, and after which a standard scheme has been modeled, are the glucose system of E. coli and the lactose system of S. aureus. This model has been developed by the isolation of individual components and their reconstitution in vitro. A general scheme for PTS-mediated sugar transport is outlined in Fig. 1. The phosphoryl group is transferred from PEP to a small molecular-weight protein called HPr. This translocation is mediated by the enzyme termed Enzyme I (El). El itself forms a phosphorylated intermediate. From HPr, the phosphoryl group is transferred to a third protein referred to as Enzyme III (EIII). The final steps involve the concomitant transfer of the phosphoryl group from EIII to the sugar as it crosses the cytoplasmic membrane. The permease is referred to as Enzyme II (Eli). Both EIII and Eli are sugar specific (31,55,77).


















Fig. 1. Schematic of the phosphoenolpyruvate-dependent phosphotransferase system (55). I (Enzyme I) and HPr refer to the general phosphoryl carriers found in the cytosol. III (Enzyme III) refers to the sugar specific phosphoryl donor. II (Enzyme II) is the membranebound permease.














S-P
PEP I HPr-P I



pyruvate I-P HPr I-P / IN OUT






5


The first two proteins (HPr, El) are general proteins in that they are involved in the transport of all "PTS sugars" by a given cell (21, 25,78). HPr has been studied in S. aureus and E. coli. In both cases it is a small molecular weight protein; the E. coli HPr is 9600 daltons

(1) and the S. aureus HPr is 9000-9200 daltons (77). Its physiological activity is stable to heating at 100 C for several minutes (25,29). From studies of the rates of hydrolysis of phospho-HPr (HPr~ P) under acid and alkaline conditions, it has been determined that the phosphoryl group is transferred to the N-1 of the imidizole ring of histidine (1,76). This component has been shown to be non-specific since point mutations in the gene coding for HPr produces pleiotropic effects in terms of sugar transport (55,78). Attempts to cross species lines have met with little success, although the E. coli HPr allows low levels (5% of the homologous system) of phosphorylation of thiomethyl-S-Dgalactopyranoside (TMG) by the S. aureus PTS enzymes. However, the S. aureus HPr cannot substitute for E. coli HPr when c-methylglucoside (a-MG) is the substrate (77). Durham and Phibbs (9) were unable to demonstrate cross-complementation between Pseudomonas aeruginosa and E. coli. Simoni et al. (77) reported the block occurrred between HPr~ P and sugar phosphorylation when S. aureus HPr was substituted in the E. coli system. Cords and McKay (6) reported cross-complementation of crude extracts from S. aureus and S. lactis; however, the results are difficult to evluate since a quantitative analysis was not presented.
The El protein has been isolated from S. Aureus and E. coli. However it has been only partially purified and the failure to attain full purification of this protein has been attributed to its sensitivity to






6



oxidation (25,30). In S. aureus, the phosphorylated derivative of this enzyme migrates as a single band of 80,000 daltons in sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) (25). The molecular weight of this enzyme in E. coli and Salmonella typhimurium has been estimated to be 70,000 and 90,000 daltons, respectively (21).

The cytoplasmic localization of El and HPr was determined by the

complete separation of cytoplasmic and membrane fractions (31,77). In S. aureus this was accomplished by ultracentrifiguation (77). The biological activity of HPr and El has been elucidated mainly in the laboratory of Dr. S. Roseman from studies with E. coli and S. aureus. Kundig and Roseman (30) demonstrated that in the presence of 32p enolpyruvate, and Mg2+, the rate of the reaction involving the transfer of the phosphoryl group to HPr was directly proportional to El concentration, whereas HPr demonstrated saturability. This indicated
that the catalyst was El while HPr served as a phosphoryl carrier. The reaction is stoichiometric and dependent on the PEP concentration. Furthermore, determination of equilibrium constants using components isolated in S. aureus demonstrated that the energy inherent in the enolate bond of PEP is essentially maintained in HPr~ P (76). During

this transfer El forms a transitory phosphorylated intermediate (21, 25). With high concentrations of PEP and pyruvate, an abortive

complex is formed and thus the PTS reaction is inhibited. Saier et al.
(64) used this observation to develop a reaction to directly assay for the presence of El; that is, El catalyzes a phosphoryl exchange between PEP and pyruvate.






7


The sugar-specific component, EIII, shows more variation among
organisms and also among PT-systems within the same organism. This is best illustrated by three systems: the lactose PTS (lac PTS) of S. aureus and the two glucose phosphotransferase systems (glc PTS) of E. coli.
The EIII of the lac PTS in S. aureus was purified from the soluble fraction of an ultracentrifugation of cell extracts (22). Studies showed it to be a trimer composed of three identical sub-units. Various physical techniques such as analytical ultracentrifugationand SDSPAGE allowed a measurement of 35,700 daltons for the trimer (22). This protein is auto-catalytic; i.e., it catalyzes the transfer of the phosphoryl group from HPr to itself (22). Each sub-unit is capable of accepting a phosphoryl group on a histidyl residue. Hydrolysis
under acid and alkaline conditions allowed the determination of the phosphoryl bond to be at the N-3 of the imidizole ring (22,25). The phospho-EIII (EIII P) then is able to donate the phosphoryl group to lactose (76). Experiments with membrane-bound EIII have demonstrated

that the phosphorylated form of EIII exhibits lipophilic behavior, whereas the non-phosphorylated does not. Sub-unit exchange was readily demonstrated with the phosphorylated form but not the underivatized form. Also, dissociation of the phosphorylated form by lipophilic agents was observed. It was suggested that physiologically EIII exists as an
underivatized, non-phosphorylated trimer and a membrane-bound phosphorylated monomer. It is in the latter form that the EIII has been postulated to be able to intercalate in the membrane and to donate the phosphoryl group to the incoming sugar (25).





8


As stated, in E. coli there are two different glc PT-systems. One has been characterized as a high-affinity system and is able to phosphorylate the glucose analogue, a-MG, as well as glucose. This system, similar to the lactose system of S. aureus, features a "soluble" EIII. Unlike EIIIlac, the phosphoryl protein link is via an acyl group (21,25). Its apparent molecular weight is 20,000 (25). As will be described below, EIIIglc may have a regulatory role (73).

In contrast to the latter two systems is the low-affinity glucose system of E. coli. This system has been shown to transport glucose and its anomers, mannose and fructose. In addition, it transports the glucose analogue 2-deoxyglucose (2-DG) as well as being responsible for the accumulation of the acetyl and N-acetyl derivatives of glucosamine and mannosamine (30,55). This EIII is an integral membrane protein which is complexed with the translocating component, Eli. It has been isolated

by Kundig and Roseman (31) and shown by isoelectric focusing to be composed of three proteins each of which showed EIII activity, albeit each with differeing specificities. One band had EIII activity with mannose, a second was specific for fructose and the third was specific for glucose (31).
The soluble components have been isolated and well characterized.
Their biological activities have been established through reconstitution.
Furthermore, since the phosphoryl addition involves sequential steps, the order as well as the individual proteins involved have been established through the use of a 32P-probe and the isolation of the intermediates by electrophoresis (30) or gel filtration chromatography (76). The transport step and the concomitant phosphoryl transfer to






9


the sugar is more poorly understood. Transport requires an intact membrane and conversely membrane proteins require lipid matrices to function; thus dissection of this step is a more formidable task.
Various Eli's have been isolated and along with EIII~ P have been shown to complete the phosphoryl transfer to the sugar. Among the Eli's that have been isolated thus far is the Eli from the low-affinity glucose system in E. coli (31) and the EIIlac of S. aureus (77).

The E. coli EIIglc was isolated using a butanol/urea extraction technique. It was shown to have biological activity in the presence of phosphatidylglycerol and the requirement for this phospholipid was stringent (31). This observation was surprising since this is a minor lipid within the E. coli membrane (21).
EIIlac was purified by using a combination of agents including sodium deoxycholate, Tween 40 and Triton X-100 (77). Lipid-free EII could be obtained by sucrose-gradient centrifugation and biological activity was detected in the presence of the other PTS components if a lipophilic environment is created with a detergent such as Triton X-100

(25).
Identification of products of phosphotransferase system activity.
The definitive assignment of functional phosphotransferase systems involves isolating and identifying the derivatized sugar(s) as product(s). Reconstitution of the S. aureus lac PTS results in the formation of TMG-6-P (77), and lactose-P (78) where the phosphoryl group is esterified at the C6 position of the B--galactopyranosy1 moieties. The E. coli glucose systems form glucose-6-phosphate (30,55), a direct intermediate in glycolysis. Fructose is phosphorylated at Cl (55).






10


Other phosphotransferase systems for sugar uptake. Many systems for sugar transport have been investigated to date. Most of these systems conform to the model discussed in the preceding section. However, a few have been found to diverge from the accepted scheme. For illustrative purposes two will be discussed in this section.

The obligate aerobe, P. aeruginosa, contains an unusual PTS for

fructose utilization. Only two components have been identified in this system, one membrane bound and one soluble. The soluble protein appears to be similar to El in terms of molecular weight; however, unlike El, this may be a peripheral membrane protein since membranes alone have residual PTS activity (9). On the other hand, this residual activity may represent cytoplasmic contamination of the membrane preparation used in the study cited. The present study of S. mutans GS5 will present a similar finding with regard to the glc PTS (see RESULTS).

Another interesting divergence from a better known system is the inducibility of these two components in the pseudomonad. In E. coli, both HPr and El are constitutive. However, the relative levels are enhanced when glucose is present (53). The membrane components (Eli, EIII) vary also with the composition of the medium. However, the relative levels of the sugar-specific components allow for the conclusion that the E. coli low affinity glucose system is constitutive and the high affinity system is inducible (21). The sugar specific components (Eli and EIII) of the S. aureus lac PTS are inducible whereas the equivalent components for glucose transport are constitutive (21). A different pattern is observed in P. aeruginosa. Both factors are inducible (9). This may be related to the presumably secondary role the PTS plays in this organism.










The occurrence of the PTS is directly related to the metabolism of the organism. In general, strict aerobes do not have a PTS. There

are a number of exceptions such as P. aeruginosa. Also, heterofermentative genera do not transport sugars via a PTS. This is presumably due to the stoichiometric relationship between the fermented sugar and the generation of PEP (21,54). Again P. aeruginosa is an obvious exception to this generalization. Durham and Phibbs (9) have proposed that after being brought in by a PTS, a putative kinase is involved in phosphorylating fructose. Thus, fructose plus PEP results in fructose-l-phosphate which then accepts a second phosphoryl group from

ATP to form fructose-l1,6-diphosphate. Generally, organisms which have phosphotransferase systems are either anaerobes or facultative anaerobes. These organisms utilize the Embden-Myerhof pathway and thus generate two moles of PEP for every mole of sugar fermented (21,54). The advantage of this type of transport lies in the conservation of ATP (54).

In Spirochaeta aurantia, a facultative anaerobe, only mannitol is transported by a PTS. The product of this reaction is mannitol-lphosphate. Three proteins are required for this reaction. Two are

soluble and on the basis of physiochemical properties are analogous to HPr and EI. The third protein is a membrane-bound EII (61). The regulation of this system differs from the model systems. In these systems HPr and El are regulated on one operon, whereas the sugar speccific components are coordinately regulated as a separate operon. These operons map at distinct loci on the E. coli chromosome (55). In S. aurantia, the three PTS proteins are genetically regulated in a coordinate fashion. In addition, the enzyme responsible for the first






12


step in mannitol dissimilation, mannitol-l-phosphate dehydrogenase, is also regulated in the operon (55).

Phosphotransferase systems and the regulation of transport.

In addition to a transport function, the phosphotransferase systems (PT-systems) in the Gram negative enterics function in the control of non-PTS transport. This control is bilateral: the permeases are regulated directly through allosteric modulation and indirectly by regulating their synthesis (55).

In order to demonstrate a model of regulation, the control of

the glc PT-systems over the lac permease of E. coli will be described as an example. A brief review of the lac operon will aid in explaining this model. The controlling factors in transcription of the lac operon are two-fold: a negative modulator, the repressor, and a positive modulator cyclic adenosine 5'-monophosphate (cAMP).

In the absence of inducer (lactoseor one of its analogues) a

repressor molecule inhibits the transcription of lac RNA by binding to an operator site on the DNA. This operator site is flanked by the promoter site on one side and the structural genes on the other. In the presence of an inducer, a repressor-inducer complex is formed. The binding of the inducer causes the repressor to undergo an allosteric shift and therefore causes it to lose its affinity for the sequence of nucleotides to which it binds. It is evident that in the absence of intracellular inducer the lac permease cannot be expressed and thus by controlling the transport of inducer molecules, the cell is able to control the synthesisoftransport systems. The prevention of inducer entry is termed "inducer exclusion" (60).





13



RNA polymerase binding and initiation of transcription is dependent on the formation of an "open complex," that is, the conformation of the DNA is altered as to produce localized melting. The effect of this is to allow the polymerase to transcribe the lac genes (45).

The formation of the "open complex" is brought about by a second
modulating system composed of two elements: cAMP receptor protein (CRP) plus cAMP. This system is analogous to the repressor-inducer complex except that it exerts a positive control. CRP by itself does not exhibit an affinity to the DNA. However when complexed to the low molecular

weight effector molecule, cAMP, its affinity towards the DNA increases by virtue of an allosteric transition. The binding site of this complex is on the lac promotor in the region distal from the operator (45). Thus, it is obvious that controlling cAMP synthesis would control the level of expression of the lac permease, and therefore, lactose uptake. Synthesis of cAMP is accomplished by regulation of the allosteric enzyme, adenylate cyclase.

It has long been observed that growth of E. coli in a combination of lactose and glucose results in abiphasic growth pattern. This diauxic growth is a result of the repression of the expression of the lac genes

(60). Not only is the inducer (lactose) excluded from these cells but cAMP levels are low. It is during the lag period preceding the second burst of growth, following glucose exhaustion, that cAMP is synthesized and sufficient quantities of inducer are accumulated to induce the lac operon.
If cells are grown on a permissive growth substrate such as glycerol, where the expression of the lac genes occurs in the absence of inducer






14


and glucose is added, two effects are observed: (1) there is an immediate cessation of lac gene transcription followed by (2) a resumption of transcription at a repressed level. The immediate severe repression is called transient repression and the second type of repression is termed catabolite repression (38). Both these forms of inhibition occur even though the inducer is present intracellularly. On the other hand, both phenomena reflect the cAMP levels measured. That is, glycerol is a permissive substrate because it allows cAMP to be synthesized, whereas growth in glucose causes an inhibition of adenylate cyclase; thus cAMP levels are low in cells growing on glucose (39,46). The addition of glucose to cells growing on a permissive substrate results in a severe transient decrease in intracellular cAMP followed by a resumption of synthesis of this metabolite but at a lower level than that observed in the absence of glucose (38). The lowering of cAMP levels is due to an inhibition of adenylate cyclase (46), excretion is a function of intracellular levels not of growth substrates (45).
Catabolite repression is a misnomer since the effect does not
require metabolism of glucose. Non-metabolizable analogues such as a-MG produces the same effect (45,47). The glucose effects (i.e., inducer exclusion, transient repression, and catabolite repression) are exerted with a wide range of metabolizable sugars. Most of these systems have in common the following: they are inducible and they are transported by non-PT-systems, though their transport may be mediated by an ATPdependent or a proton motive force-dependent mechanism (45,55).
A unifying hypothesis to explain the two underlying mechanisms of repression, inducer exclusion and adenylate cyclase inhibition, has been






15


proposed by Saier (55). This is diagrammed in Fig. 2. The overall mechanism is a phosphorylation-dephosphorylation modulation of competing functions which are all dependent on a central protein. In the outline, the phosphorylation reaction is mediated by the general PTS proteins, El and HPr. In the presence of glucose, the equilibrium of the phosphoryl donation lies in the direction of glucose phosphorylation; however in the absence of glucose, the phosphorylated HPr is free to donate

the phosphoryl group to a hypothetical protein termed RPr (regulatory protein). In the phosphorylatedform, this protein becomes a positive effector for adenylate cyclase, thus allowing cAMP to be synthesized thereby negating catabolite and transient repressions. In the nonphosphorylated form (i.e., when glucose is present), RPr has an affinity for the non-PTS permeases. The proposal is that these permeases are allosteric proteins and that RPr is a negative effector. Therefore, RPr binding inhibits transport of non-PTS sugars and is responsible for inducer exclusion. When phosphorylated, RPr loses its affinity for the permease and the unmodified permeases can then function.
Much evidence has been accumulated to support this model. A selected

amount will be summarized here.

Using S. typhimuriumand E. coli, it was found that mutations in the genes coding for El or HPr had a surprising effect on the metabolism of non-PTS sugars such as melibiose and maltose. Tight mutants could not grow on these substrates, whereas the growth pattern of leaky mutants were phenotypically indistinguishable from wild-type. However, if a-MG is added to induced cells, the leaky mutants exhibit a profound increase

in sensitivity to the repressive effects of this analogue as























Fig. 2. Proposed scheme for the regulation of carbohydrate transport and metabolism by the phosphoenolpyruvate-dependent phosphotransferase system in E. coli (55). Abbreviations are as follows: PEP, phosphoenolpyruvate; I, Enzyme I: II, Enzyme II; S1, sugar; S1-P sugarl-phosphate; RPr, regulatory protein: A.C., adenylate cyclase. The permease refers to a non-PTS permease; i.e., ATP- or proton motive .force-dependent transport.


















IN OUT


St-P RPr P (Inactive t rPermease)




PEP 8 r + P (Active SPermCyclase)


pyruvate P + activee Adenylate
Cyclase)

P (Active Adenylate
(AC) Cyclase)






18


compared to wild-type. This repressive effect is evident in terms of growth and enzyme synthesis (66). The degree of this repression is related to the extent of induction. Thus, fully induced cultures are more refractory towards the repressive effects of a-MG. Also, noninduced cells cannot grow if transferred to a medium containing this glucose analogue and the non-PTS sugar, melibiose (60,63). This indicates that the primary effect being investigated in these studies is inducer exclusion (63). Furthermore, the Eli specific for the PTS sugar must be present for the sugar to cause this hypersensitivity (63, 66), substantiating the involvement of PTS-mediated transport.

A second mutation described by Saier and Roseman (62) mapping at a

site co-transducible with the pts operon (genes for El and HPr) but not part of the operon was found to suppress mutationswithin the pts operon in terms of the hypersensitivity and total repression described above. The distinguishing phenotypic characteristic resulting from these suppressor mutations was found to be depressed levels of the EIII for glucose; i.e., the soluble factor which is part of the high affinity glc PTS. These mutants were termed catabolite repression resistant and the gene responsible was termed crr. It is postulated that this protein (EIIIgc) may indeed be RPr (60).

A second group of mutants were isolated by Saier et al. (67) which also relieved PTS-mediated repression; i.e., repression due to an El or HPr mutation. The characteristic that distinguished this second group from the crr mutants, is the specificity of the relief. The crr mutation was general in that both transport and metabolism of all the affected non-PTS sugars were relieved of repression, whereas in this second






19



category, a number of unique mutants were isolated which exhibited a refractory response when a specific non-PTS sugar was tested. That is, there was a double mutant in which maltose transport and dissimilation

was released from repression, one which melibiose transport was released, etc. The mutations responsible for this release mapped in or near the genes coding for the individual transport proteins. These results suggest that an altered permease may be responsible for this relief and thus indicate an allosteric interaction between a PTS component and the permeases (60,67).

Lastly, increased levels of PEP was shown to partially overcome the inhibitory effect of a-MG-induced repression of glycerol transport in wild-type cells. This indicated phosphorylation involving PEP was utilized by these cells to regulate sugar uptake (58).

All these results relate to the transport of non-PTS sugars and thus the regulation of non-PTS-mediated transport. An interaction was also observed between PTS components and adenylate cyclase. However, the effects of the various genetic lesions were not completely analogous. The following observations are relevant to the discussion on adenylate cyclase: (1) Mutations affecting the low affinity Eli proteins render the adenylate cyclase of E. coli insensitive to glucose and its analogues (i.e., 2-DG) but not a-MG, whereas mutations resulting in a disfunctional EII of the high affinity system render the enzyme insensitive to a-MGinduced but not 2-DG-induced inhibition (19). These mutations affect the regulation not the activity of the enzymes and these data indicate a functional PTS is required for adenylate cyclase regulation.






20


(2) In vitro assays of wild-type adenylate cyclase fail to show

glucose-mediated inhibition of this enzyme, again pointing to the need for transport activity (47).

(3) El deletion mutants show low levels of adenylate cyclase activity. Leaky El mutants are "hypersensitive" to the inhibitory effects of glucose on adenylate cyclase activity and this "hypersensitivity" could be partially overcome in permeabilized cells by the addition of PEP (48,57).

(4) A mutation in the crr gene results in depressed levels of cAMP in cells which have wild-type levels of El and HPr (48).

(5) Kinetic analysis revealed that adenylate cyclase was still
functioning in vivo albeit at depressed levels in the presence of glucose. This tends to indicate positive regulation is required and taken with other data, glucose prevents the positive regulation of adenylate cyclase. These data also suggest that the crr gene product (EIIIglc) may be involved in regulation. The effect of temperature on the acivity and regulation of adenylate cyclase suggests that a stable complex, i.e., an allosteric interaction, is a prerequisite for activation (48).
The isolation of the crr mutation along with the observation that pre-growth on glucose enhanced the inhibition of transport suggests that
a component of the inducible high-affinity glucose system is also involved in inducer exclusion. In support of this, Scholte et al. (73) have shown the identity of the crr gene product to be EIII glc using crossimmunoelectrophoretic techniques.
Other phosphotransferase functions. There is evidence that the PTS functions in the regulation of motility (55). The responsible factor appears to be the sugar-specific Eli protein (36). In addition, the Eli






21


protein may play a role in the fermentation of the sugar they are transporting (36). Lastly, the various Eli's are able to catalyze the transphosphorylation reaction: the transfer of a phosphoryl group from a derivatized sugar to its non-derivatized counterpart (36,59). The physiological significance of this latter reaction is unknown; however, there has been speculation that it may have a regulatory function (60).

Characterization of the phosphoenolpyruvate-dependent phosphotransferase systems in the strptococci. The literature concerning the streptococci lacks the detail outlined above. As demonstrated in the above discussion, the model system is based on data collected from studies in E. coli and S. aureus. Studies on the streptococci do not involve the molecular dissection of the various PT-systems. Identification of PTS transport in these organisms is based on one or more of the following criteria: stimulation of transport by intracellular PEP reserves and/or PEP-dependent phosphorylation of a sugar, the demonstration that ATP cannot substitute for PEP for either of these two functions, and the identification of the transport product of a sugar and/or its non-metabolizable analogue as a phosphorylated derivative.

The most extensive studies of the lactic streptotocci have been on
the lac PTS in Streptococcus lactis. McKay et al. (42) demonstrated that NaF, an inhibitor of enolase which catalyzes the glycolytic step involved in the conversion of 2-phosphoglycerate to phosphoenolpyruvate inhibits the transport of TMG, a lactose analogue. The intracellular product was identified as TMG-P in this and other studies (42,86). Thompson (86) measured PEP utilization along with TMG accumulation and found there was a stoichiometric relationship between these two parameters.






22



Furthermore, the conditions used in this study did not permit the ATPdependent accumulation of amino acids; thereby demonstrating that PEP was not being converted to ATP by pyruvate kinase.

In this organism, lactose metabolic enzymes are plasmid coded (32). In a strain devoid of its lac plasmid, both lac PTS and phospho-a-galactosidase activity are absent, suggesting a close linkage between these lac genes. However, regulation of these enzyme activities does not appear to be coordinated. In wild-type, the phospho-6-galactosidase is present under growth conditions which repress the expression of the lac PTS (32). This is an interesting observation in light of other studies on Gram positive bacteria suggesting that the inducer for the lac PT-systems is galactose-6-phosphate. Morse et al. (43) isolated phospho-a-galactosidase negative mutants from S. aureus and found that these cells could not transport lactose when grown under conditions which induce the lac PT-system in wild-type. However, galactose-6-phosphate was able to induce this system in such mutants. As stated in a previous section, incoming lactose is phosphorylated at the C6 of the galactose moiety (78). Also, early reports indicate the constitutive nature of the phospho-a-galactosidase of S. aureus (41).

In strains of S. lactis, galactose is a more potent inducer of the lac PTS than lactose itself (6,32). Galactose has two transport systems in these cells, one which requires ATP and a second which requires PEP. In an elegant series of experiments, Thompson demonstrated the existence of two systems (88). He accomplished this by comparing galactose dissimilation in the presence of glycolytic inhibitors such as iodoacetate or

the presence of ionophores. The latter he termed the Gal P system.






23



He confirmed his findings by using an ATP-generating substrate, arginine, which allowed galactose transport but not PEP-mediated lactose transport. Through competition studies, it was shown that PEP-dependent galactose transport was mediated by the lac PT-system. This is in agreement with LeBlanc et al. (32) who were able to demonstrate the coincident disappearance of the lac and gal PT-systems upon curing a strain of S. lactis of one of its plasmids. In another study, Cords and McKay (6) isolated a revertant of a lac PTS strain. However, galactose could not induce the lac-PTS in these cells and upon closer examination, it was shown that these cells only possessed the gal P system.

Transport via these two different systems results in the dissimilation of galactose by two different routes. If galactose is translocated by the gal P system, it enters the cell cytosol as a free sugar. Subsequently, it is phosphorylated by galactokinase and ATP (6), and is metabolized via the Leloir pathway (88) to glucose-l-phosphate and consequently by the Embden-Myerhof pathway upon isomerization to glucose-6phosphate (90). Galactose-6-phosphate is the product of PTS transport. This compound is metabolized through the tagatose pathway (43,88). This biochemical sequence involves the conversion of galactose-6-phosphate to tagatose-6-phosphate which is phosphorylated by ATP to tagatose-l,6diphosphate and finally to dihydroxyacetone phosphate which can feed into the Embden-Myerhof path (87). It should be noted that in these cells, lactose is phosphorylated at the C6 of the galactose moiety. Thus it also must be metabolized via the tagatose pathway with the free
glucose produced by the action of phospho-B-galactosidase being processed through the Embden-Myerhof pathway (87).






24



There appears to be some controversy as to which galactose transport system is the predominant mode of transport. Based on their biochemical analysis of mutants, Cords and McKay (6) speculated that galactose-6phosphate repressed the gal P system. However, Thompson (88) found that the gal P had a 10-fold lower K for galactose than the lac PTS. In addition, he found growing cells contained low levels of tagatose phosphate but high levels of fructose phosphate which indicated the Leloir path was operating. He interpreted these results to mean that the Leloir path is the predominant route of galactose degradation in relatively low levels of sugar and thus represents the major route.

In addition to these two types of galactose transport systems, a second gal PTS appears to exist (44,88). This system is specific for galactose and thus gal+ lac- cells can only grow on galactose. Other

phenotypic traits of this strain are the inability to accumulate TMG and the production of galactose-l-phosphate and galactose-6-phosphate. As already discussed, these are the first intermediates in galactose metabolism when galactose is transported by the gal P and the gal PTS, respectively. Thus, these cells which are lac PTS deficient are still able to transport galactose through a PTS in addition to a proton-driven permease. It is apparent that galactose transport in this organism is

compl i cated.
A second PEP-mediated transport system in S. lactis has been defined for glucose. Thompson (86) has shown that there is a stoichiometric relationship between PEP depletion and 2-DG uptake. He also demonstrated

that the selective use of inhibitors of glycolysis such as P-chloromecuribenzoate (target: glyceraldehyde-3-phosphate dehydrogenase) leads to






25



inhibition of 2-DG uptake in metabolizing cells. Based on the following criteria this glc PTS was classified as being similar to the low affinity E. coli system: (1) it was refractory to the inhibitory

effects of sulfhydryl reagents, (2) the following glucose analogues appeared to compete with 14C-glucose: 2-DG, mannose, and N-acetylglucosamine, while the substrate of the high affinity system, e-MG, was without effect, and (3) it is a constitutive system (86).

It appears that the predominant transport system for sugars in the cariogenic streptococcus, S. mutans is of the PEP-dependent phosphotransferase type. This is not surprising in view of the fact that this is an aerotolerant anaerobic organism (85), and, therefore, must conserve its energy reserves. The PT-systems studied to date in this organism are outlined in Table 1. The literature contains an extensive

survey of the various transport systems found in the species; however, details of the individual systems are lacking.

The first system to be defined was the glc PTS. Schachtele and

Mayo (71) demonstrated a PEP-dependent phosphorylation reaction involving 2-DG in permeabilized cells of S. mutans. ATP could not replace PEP in this reaction. Supporting evidence for a PEP dependent transport system came from the observation that NaF inhibited the uptake of 2-DG. Based on the failure of 6-DG to inhibit phosphorylation of 2-DG, they concluded that this analogue is phosphorylated at C6. These investigators extrapolated their findings to the natural substrate, glucose. Even though it is an obvious assumption that 2-DG is transported and phosphorylated by the same proteins involved in glucose uptake, competitive inhibition studies to prove this point have never been done.






26







Table 1. Phosphoenolpyruvate-dependent phosphotransferase systems
in S. mutans.


Glucose (Schachtele and Mayo, 1973)
Mannitol (Maryanski and Wittenberger, 1975) Sorbitol (Maryanski and Wittenberger, 1975)

Lactose (Calmes, 1978)

Sucrose (St. Martin and Wittenberger, 1979)






27


Two other monosaccharides studied in this organism are the hexitols: sorbitol and mannitol. Both of these sugars are substrates of a PEPdependent phosphorylation reaction. These PT-systems are inducible and therefore are evident in cells only when grown on the respective sugars. The enzymes, mannitol-l-phosphate dehydrogenase and sorbitol-6-phosphate dehydrogenase are coinduced indicating a regulon codes for these PTsystems (40).
The most thoroughly investigated PTS in this organism is the sucrose (suc)PTS. The reason for this is the medical implication of sucrose metabolism in this species. This organism is able to synthesize a dextran capsule through the action of a glucosyltransferase and it is this capsule that allows adhesion to tooth surfaces (13,14).

Slee and Tanzer (80,81) described a unique PTS for sucrose in this species. Using a variety of strains, they demonstrated a PEP-dependent

phosphorylation of sucrose to sucrose-6-phosphate which appeared inducible; i.e., it was not present in glucose adapted cells. St. Martin and Wittenberger (83,84) verified the inducibility of the suc PTS by employing a greater variety of growth substrates.

Upon translocation of sucrose, the dissacharide is phosphorylated

at the C6 position of the glucose moiety (83,84). Hydrolysis of sucrose6-phosphate requires a unique invertase. A sucrose-6-phosphate hydrolase was described in S. mutans (5,84). This enzyme had a high affinity for

its substrate and thus could only be detected directly by using sucrose6-phosphate. It is synthesized constitutively, unlike the suc PTS (84). It is interesting to compare this system to the lactose system in S. aureus. In this system galactose-6-phosphate is the inducer (43) and this compound






28



is the product of a constitutive hydrolase (41). It may be that sucrose6-phosphate hydrolase activity results in the production of the suc PTS inducer. Indeed, sucrose-6-phosphate hydrolase negative cells which

have an evident suc PTS are isolated as PTS constitutive (84). However, using the S. aureus analogy, it seems unlikely that glucose-6-phosphate is the inducer since glucose-grown cells have repressed levels of suc PTS activity (81,83,84). The other product resulting from the action of this hydrolase is fructose. Growth of cells on fructose results in lower levels of the suc PTS than growth on glucose (83). Further experiments must be performed to clarify these contradictory data.

Thompson and Chassy (89) have recently described a suc PTS in S.

lactis. It is interesting that in this organismthey found the sucrose-6phosphate hydrolase to be inducible. They also found a fructokinase

which was induced in sucrose-grown cells. The identification of this latter enzyme in cells metabolizing sucrose along with the finding that the cytosol of these cells contains high levels of free fructose, lead these investigators to conclude that the fate of the fructose resulting from the hydrolysis of sucrose-6-phosphate was phosphorylation by an ATP-dependent kinase.

Recently a suc PTS has been reported in E. coli (35,72). This is an interesting finding since it had been previously thought that in this organism the PTS was used exclusively for the transport of monosaccharides

(55). The ability to utilize sucrose is a plasmid-borne function, the origin of which appears to be Klebsiella pneumoniae (35). This plasmid bears two genes coding for sucrose metabolic enzymes, Elisuc and sucrose6-phosphate hydrolase (72). There are some interesting features of this






29



system. First, there is a coordinate induction of these two proteins and genetic studies suggest that the inducer is fructose or its phosphorylated derivative. This is different from S. mutans where fructose appears to be a repressive sugar for the suc PTS (84). In addition the genes for these two proteins do not appear to be transcribed as a regulon in S. mutans. Additional evidence that a regulon exists in E. coli is the isolation of a mutant constitutive for both proteins. Another unusual feature is the apparent requirement for the EIIIg9lc. This was demonstrated by transducing the suc PTS into EIII negative strains of E. coli and was confirmed using in vitro reconstitution of the individual protein (35).

Using the criteria of PEP dependency, NaF inhibition, and product isolation, Cal-nes (3) concluded that lactose uptake in S. mutans is mediated by a group translocation mechanism. He used a variety of assays to study this system including the hydrolysis by permeabilized cells

of the lactose analogue, o-nitrophenyl-6-galactopyranoside (ONPG), in the presence of PEP and the transport of TMG into intact cells followed by the extraction and identification of its phosphorylated derivative.

A phospho-6- galactosidase has also been identified (16,17) and

characterized (4) in this species. Conflicting data as to the regulation

of this enzyme have been reported. Results published by Calmes and Brown (4) suggest that this is an inducible enzyme. Furthermore, the preferred inducer is lactose. Galactose is also an inducer but the amount of the enzyme synthesized in the presence of this monosaccharide is less than that obtained upon lactose induction.






30


These results parallel those obtained by Calmes (3) when investigating the lac PTS; namely, both lactose and galactose are inducers with

the dissacharide more active in this regard. These findings are in direct disagreement with the induction pattern of the lac PTS postulated to occur in S. aureus. The results of Hamilton and Lo (17), however, suggest a similarity between the mechanisms of induction of the lac PTS in S. mutans with that in S. aureus. That is, in S. mutans the levels of phospho-B-galactosidase vary to some extent with the growth condition; however, this enzyme is always present. Also the levels are highest when cells are grown in galactose even though with some strains tested the relative levels were close. The reasons for the discrepancies between the authors may be due to the different strains used in the respective studies. Finally, in a later report published by Hamilton and Lebtag

(16), muchmore dramatic increase in the levels of phospho-a- galactosidase are shown in cells grown in lactose and galactose when compared with glucose. For both the lac PTS and phospho-B-galactosidase, galactose is a better inducer than lactose. They interpret their data to mean a co-regulation of the lac PTS and phospho-6- galactosidase genes.

Enzymes of the tagatose pathway have been demonstrated in S. mutans

(16). The levels of these enzymes increase, though not as dramatically in all strains, when lactose is the carbon source. Furthermore, this increase is evident only when lactose, not galactose, is included in the

growth medium. Growth in galactose induces enzymes of the Leloir pathway

(16). Thus in this respect, lactose metabolism follows the model defined in S. lactis.






31


Most of the work on transport in the streptococci has been physiological in nature, The biochemistry of PT-systems in these organisms has not been extensively investigated. One exception is the study of the glc PTS of S. faecalis. Interestingly, the PTS appears to be similar to that of the low affinity of E. coli in that the EIII has been characterized to be membrane bound and complexed to EII (27).

Two attempts at dissecting the PTS in S. mutans are noteworthy.

Schachtele (70) isolated membranes from glucose-grown cells of S. mutans. These membranes alone were able to carryout a PEP-dependent phosphorylation of glucose. Since the method of membrane preparation was crude and

yielded impure membranes, definitive conclusions as to the cellular location of the various components could not be drawn. Another study by Maryanski and Wittenberger (40) led to equally unsatisfactory results.

They investigated the localization of components of the glc and mannitol (mtl) PT-systems by recombining soluble and particulate extracts of induced and non-induced cells and determining the amount of sugar phosphorylation. In recombining extracts of glucose-grown cells, they were able to demonstrate the phosphorylation of glucose but not the phosphorylation of mannitol Soluble and particulate fraction from mannitolinduced cells recombined to phosphorylate both mannitol and glucose (i.e., glucose is constitutive). More importantly, they were able to demonstrate a mannitol reaction when the glucose soluble and mannitol particulate fraction were recombined. Alone neither one catalyzed phosphorylation of mannitol. This indicated that both soluble and particulate components

exist in S. mutans. However, in both cases the soluble components alone were as active in phosphorylating activities,indicating that the method






32


of membrane preparation (sonication) resulted in the formation of small membrane fragments. Interestingly, in constrast to Schachtele's work, the

pellets alonedid not show activity, supporting the conclusion that extreme fragmentation of membranes occurred during sonication. This apparent failure to clearly separate the particulate and soluble fraaction make any interpretation difficult.
Maryanski and Wittenberger (40) were able to demonstrate the presence of a heat stable component that could enhance phosphorylation of mannitol by cell-free extracts. One minor but perhaps significant point is the ability of Ca2+ to replace Mg2+ in their assay system and the inhibition that Zn2+ produced. In E. coli these ions had opposite effects (30). Mg2+ is required for the El catalyzed transfer (25); therefore, this difference may reflect a subtle diversity between El components.

Regulation of carbohydrate utilization in Gram positive organisms
The systems of regulation in the Gram negative enterics have been studied extensively by a number of laboratories. Much of this work has been concerned with Saier's model (55) of regulation. To date very little is known about regulation of transport in Gram positive cells and much less is known about the streptococci. One reason for this lack of knowledge may be the absence of genetic systems of study in these organisms.

As with studies of sugar transport, studies of regulation by these organisms have progressed little beyond the descriptive stage. Hamilton and Lo(17) surveyed various oral streptococci for the induction of lactose metabolism. A comparative study of S. salivarius and S. mutans was performed. Their data did suggest that the predominant lactose pathway in S. salvarius was via 6-galactosidase indicating the presence






33


of a non-PTS permease, whereas S. mutans contained phospho-6-galactosidase indicating a dependence on the PTS for transport. They based their conclusions on the observation that ONPG could be hydrolyzed by permeabilized cells of S. salivarius with equal efficiency in the presence or absence

of PEP. However, this result could be due to a large energy reserve within these cells. Interestingly, they observed a true diauxie with S. salivarius when these cells were grown in equimolar lactose and glucose, where in S. mutans lac PTS induction under these conditions was observed at low glucose concentrations. In a parallel experiment they demonstrated that isopropyl-8-D-thiogalactopyranoside (IPTG) and galactose could induce lactose metabolic enzymes in glucose-grown cells of both S. mutans and S. salivarius. The obvious contradiction of these

two experiments is difficult to reconcile. The latter experiment employed 28 mM glucose as a growth substrate; however, the authors do not mention the amount of glucose remaining at the time of the addition of 8 mM IPTG. The addition of IPTG alone (control) to cells of S. salivarius produced a greater amount of $-galactosidase activity than when glucose was present, indicating a repressive effect of glucose; however, the authors made no mention as to the growth substrate in these control cells. The analogous data obtained for S. mutans were not presented.

A diauxic growth pattern occurs when glucose-grown cells are transferrred to a medium containing equimolar glucose and sucrose (82). The second phase of growth correlates with induction of the suc PTS.
The mechanism (s) which allows preferential sugar utilization in Gram positive organisms has not been elucidated. The model evoked to explain this form of regulation in Gram negative cells is inadequate. One reason for the non-applicability of Saier model is the failure to






34


define a role for cAMP in these organisms. In studies on Bacillus

subtilis (8), Lactobacillus plantarum (20,68), Bacillus megaterium (91), and S. faecalis (24), cAMP addition could not relieve repression. Interestingly, in two separate studies, S. mutans was found to respond to cAMP and adenylate cyclase was detected (17,51). However, in one case the methodology used to detect the enzyme has been shown to give artifactual results (45). It is more difficult to evaluate the second study in which the reliable method of radioimmunoassay was used to detect cAMP since no subsequent reports have appeared in the literature. The second reason it is difficult to apply the Saier model to many Gram positive organisms is that in these bacteria we see one PTS sugar (glucose) regulating other PT-systems (usually for dissacharides) rather than PTS regulation over non-PTS transport.
In order to explain the hierarchy of sugar utilization via the various PT-systems, Thompson et al. (90) evoked the concept of catabolite inhibition. This mechanism involves the differential affinities of the various sugar-specific components for HPr-. P. For example in the above discussion concerning S. mutans, the sucrose-specificcomponents would have a lower affinity for the HPr. P than the glucose-specific moieties.
Catabolite inhibition could explain the diauxic effect observed in non-induced cells. For instance, in S. faecalis, the kinetics of inhibition by glucose of the lac PTS appear to be competitive (23). Alternatively, this concept may be applied to the inhibition observed with pre-induced cells. For example, glucose inhibits the uptake of galactose (PTS-mediated) in galactose-grown cells of S. lactis (90). In the former case, cataboliteinhibitionwould result in a situation analogous to





35


inducer exclusion (23), whereas a general repression would be observed in the latter case (90).

Another theory put forward involved inhibition by sugar-phosphates

(60). It is postulated that a sugar phosphate binding site of the Eli may be a means of turning off transport. For instance, an increase in intracellular glucose-6-phosphate was shown to be coincident with a decrease in fructose (frc) PTS-mediated transport in E. coli. Saier and Simoni (65) have shown that lactose uptake in S. aureus inhibits uptake of other PTS sugar. These depressed rates of uptake are dependent on the lactose-specific PTS components being functional. They suggest that galactose-6-phosphate is responsible since a lag precedes inhibition when the cells are presented with two sugar substrates (i.e., lactose phosphate must be processed to produce the putative inhibition). On the other hand, Thompson et al. (90) rules out inhibition by sugarphosphates as being the cause of glucose inhibition of galactose uptake in pre-induced cells of S. lactis since the levels of glucose-6-phosphate were essentially invariable in cells growing on galactose compared to glucose. The little-understood transphosphorylation reaction has been

interpreted by Saier and Moczydlowski (60) to support their proposal of sugar phosphate mediated-regulation; however, in transphosphorylation the binding site on EII is only for a sugar phosphate derivative homologous to the underivatized form.
A third type of proposed regulation is relevant here especially when considered with the discussion to follow;this involves the inhibition of a-MG uptake by an energized membrane. The inhibition can be
reversed by the use of ionophores and other inhibitiors of the proton motive force (60).






36


In addition to the regulation observed when cells are presented with two different growth substrates, there is a regulatory mechanism imposed by growth. A secondary transport system for glucose exists in S. mutans. This is apparently dependent on an energy source other than

PEP and evidence acquired by the use of inhibitors such as the ionophore carbonyl-m-chlorophenylhydrazone suggest it is mediated by a proton pump. The affinity of this system for glucose is 8-15 times lower than the PTS (18). Thus at lower glucose concentrations, the glc PTS would be expected to predominate. This prediction was shown experimentally using the controlled conditions of a chemostat: cells grown under a low dilution rate utilized the PTS (11,18). Surprisingly, glycolytic activity was found to be greater under conditions of glucose starvation than when excess glucose was present. At the higher dilution rate (excess glucose) the PIS was apparently repressed and the low affinity system predominated. The degree of repression was found to be proportional to the growth rate (11).

As with the glc PTS, the suc PTS of S. mutans appears to be inhibited by rapid growth and excess substrate. This has been demonstrated both in batch (82) and chemostat grown culture (10). In the latter study, the investigators calculated that the amount of transport was insufficient to

account for the total sucrose uptake observed. Rapidly growing wild-type cells in batch culture had a lower specific activity for the suc PTS than did stationary-phase cells. Mutants were able to grow in sucrose even

though they were apparently devoid of suc PTS activity. The authors in both studies interpreted these data to indicate the existence of a secondary transport system. The major problem with interpretation of






37


the results of the latter study is that the extent of leakiness of these mutants was not adequately quantitated. No activity was detected using an enzyme-linked assay which indicated that this mutation was tight; however, growth on sucrose was approximately 2-log below wild-type. PTS activity below the level of detection of the assay could account for the sucrose-supported growth of the mutants. Further studies are needed in
order to determine definitively the existence of an alternate system.

Relevant to these observations is the inhibition of a-MG uptake

(PTS-mediated) in E. coli observed during rapid growth. This inhibition may be related to the energized state of the membrane (60). Interestingly, repression of the glc (11) and suc (82) PT-systems in S. mutans is pH dependent; however, activity is increased at the lower pH. If a pH potential imposed by an ATPase is responsible for regulation, activity of the ATPase would result in activation of these PT-systems according to these results since the action of the ATPase cause a decrease in the external pH.
Regulation of sugar transport in the streptococci has been largely speculative; documentation of such regulation has been sparse. The major problem is the lack of biochemical information concerning the various PT-systems. It is not feasible to propose a regulatory protein such as the EIg111lc if it is not known whether the glucose system contains an EIII-type molecule. Until now, none of the individual PTS components has been shown to exist in S. mutans. Furthermore, mutants which totally lack a functional glc PTS have not been obtained for this species. Hamilton and St. Martin (18) isolated a glc PTS-mutant from S. mutans which retained 15% of the activity of wild-type. It is difficult to






38


attribute a regulatory function to the glc PTS using such a leaky mutant.

The first step in understanding regulation of sugar transport is a more comprehensive assignment of PTS components. This work attempts to elucidate, in part, the biochemistry of the S. mutans glc PTS and to relate this to the physiological regulations observed in this organism.













MATERIALS AND METHODS

Cultures and cell growth. Cells of Streptococcus mutans GS5 were routinely maintained in a tryptone-yeast extract(TYE) broth (17). This medium was composed of 10 g/l tryptone (Difco Laboratories, Detroit, MI),

5 g/l yeast extract (Difco Laboratories) and 3 g/l dibasic potassium phosphate. For maintenance, this medium was supplemented with 20 mM glucose (Sigma Chemical Co., St. Louis, MO).

For assay purposes, the cells were grown overnight in TYE broth plus 20 mM of a given sugar. These cells were transferred to a defined medium

(DM) broth designed for S. mutans (85). To this defined medium, 5 mM sugar was added. A 10% inoculum size was routinely used. Unless otherwise specified, cells were allowed to grow until late-log/early-stationary

phase in the DM; after which time, they were harvested by centrifugation at 12,000 x q for 10 min and washed once in 100 mM sodium phosphate (PB), pH 7.0, plus 5 mM MgCl2. The centrifuge used for routine purposes was a Sorvall RC-5B (Dupont Instruments, Newtown,CN).

Cell preparation for assays. In all the assays outlined below, decryptified cells were used. The decryptification conditions were as follows: washed cells were resuspended in PB, pH 7.0, plus 5 mM MgCl2 to one-tenth their original volume. A toluene-acetone mixture (1:3 v/v) was used to permeabilize the cells. The mixture was added in a ratio of 100

I1/ml of cell suspension and decryptification was carried out by vigorously shaking on a Vortex Mixer (Scientific Industries, Bohemia, NY) for 1-2 min intervals interspersed with cooling in ice. The total length 39






40


of shaking was 3-5 min. This methodology was based on that used by Calmes (3) for S. mutans. If a further dilution was required 100 mM PB plus 5 mM MgC12 was added after decryptification to give the desired cell concentration.

Determination of the lactose phosphotransferase system and phosphoB-galactosidase. The assay for the determination of the lac PTS utilized the lactose analogue o-nitrophenyl-3-galactoside (ONPG; Sigma Chemical Co). The standard reaction mixture contained 100 mM PB, pH 7.0, 5 mM MgCl2,

8 mM sodium phosphoenolpyruvate (PEP; Sigma Chemical Co.), 8 mM ONPG, and cells (concentrations to be detailed in legends to Tables and Figures). The total volume was 0.6 ml. This was then incubated for 30 min at 37 C after which time the reaction was stopped by the addition of 1 ml of a 5%
Na2CO3 solution (aqueous). The cells were centrifuged inaSorvall table top centrifuge and the amount of o-nitrophenol (ONP) formed was determined by reading the absorbancy of the supernatant at 420 nm in a Gilford 2600 recording spectrophotometer (Gilford Instruments, Inc., Oberlin, OH).

The amount of ONP was then determined from a standard curve and the results are expressed in terms of moles of ONP/pig cells (dry weight).

For the determination of phospho-6-galactosidase activity, the

lactose-phosphate analogue, o-nitrophenyl-8-galactose-6-phosphate (ONPG-6P; Sigma Chemical Co.) was used. Unless otherwise specified, the reaction mixture contained 100 mM PB, pH 7.0, 5 mM MgCl2, 10 mM ONPG-6-P, plus cells (amounts to be detailed in legends) in a total volume of 0.5 ml. After 30 min at 37C the reaction was terminated with 5% Na2CO3, the cells were removed by centrifugation, and the absorbancy of the supernatant was

determined at 420 nm. The results were expressed as the moles ONP formed/






41


pg of cells (dry weight). The assay for the lac PTS was based on that described by Calmes (3) and that for the phospho-6-galactosidase was described by Calmes and Brown (4).

Phosphotransferase assay: LDH/NADH-linked. For general surveys, an

LDH/NADH-linked assay was used (29). A mixture containing 80 mM PB, pH 7.0, 4 mM MgC12, 10 mM PEP, .025 mg lactate dehydrogenase (LDH; Sigma

Chemical Co.), 3 mM B-nicotinamide adenine dinucleotide, reduced form, disodium salt (NADH; Sigma Chemical Co.) plus cells (amounts given in legends to Tables and Figures) was monitored at 340 nm in a Gilford 2600 recording spectrophotometer for NADH oxidase. An initial rate was obtained by allowing the reaction to proceed for 4-5 min. The PTS reaction was

initiated by the addition of 1 mM sugar substrate. The final volume was

1.0 ml. The reaction was monitored by measuring the decrease in absorbancy at 340 nm for 4-5 min. An initial rate was obtained from this measurement and corrected for the endogeneous NADH oxidase activity. The Vmoles of NADH remaining was determined from a standard curve and from this value the amount of NAD+ formed was determined. Results are expressed as uimoles

NAD+ formed/pg cells/min.

Phosphotransferase assay: radioactive. For a more quantitative

analysis, a radiolabelled PTS substrate was used. A typical reaction mixture contained 30 mM PB, pH 7.0, 1.5 mM MgCl2, 10 mM PEP, 10 mM sodium flouride (NaF, Sigma Chemical Co.), and 0.1 mM unlabelled sugar to which

was added the specific isotope (New England Nuclear, Boston, MA). The amount of each isotope contained in each reaction mixture is given under

the legends to the Figures or in the Results section. To standardize the reaction, 90-110 pg of cells was used and the concentration was determined from a standard curve of dry weight vs. absorbancy of cells at 600 rnm






42



(to be described in a following section). For routine purposes, the time of incubation was 30 min; for kinetic studies, the time was 10 min, since the extent of the reaction was linear within this time. Unless otherwise stated, these studies were carried out at 25 C.

To assay cell-free membranes the following modifications were made: the buffer concentration was generally increased to 80 mM, PB, pH 7.0, the MgCl2 concentration was increased to 4.0 mM and the membrane concentration varied as specified in the RESULTS. Incubation was always at 37 C. In subsequent assays, the volume was scaled down to 200 pl in order to conserve material and 4 mM 2-mercaptoethanol was added.

These reactions were stopped by diluting 100-200 pl of the reaction mixture 1:9 (v/v) in 1% sugar or when the scaled down assay was used, by adding cold 1% sugar in excess of the reaction volume directly to the assay mixture. In both cases, these mixtures were rapidly cooled. From this, the phosphorylated derivative was separated from the labelled substrate by filtration under vacuum through a DE-81 anion exchange filter (Whatman, Clifton, NJ) which had been prewashed with a 1% solution of underivatized sugar. The filters were washed one time with 1% sugar and 4 x with cold H20. They were then counted in a Beckman

LS8000 scintillationcounter (Beckman Instruments, Inc., Fullerton, CA). This assay is based on the methodology developed by Simoni et al. (76). The scintillation fluor used was Aquasol (5 ml; New England Nuclear).

Transphosphorylation. In order to assay directly for the EIIglc, the glucose-glucose-6-phosphate exchange reaction was employed. The procedure was a modification of that developed by Saier (59). In this procedure, 70 mM PB, pH 6.0, 3.5 mM MgCl2, 3.5 mM 2-mercaptoethanol,






43

50 mM glucose-6-phosphate (Sigma Chemical Co.), 10 mM NaF and 50 pM D-[14C(U)]-glucose (4 pCi/pmole) were reacted with membranes. The total reaction mixture volume was 200 pl. The mixture minus the labelled glucose was prewarmed to 37 C and the reaction was begun by the addition of D-[14C(U)]-glucose. Incubation continued at this temperature for 30 min after which time the reaction was stopped by the rapid addition of cold H20 and the separation of derivatized product from reactant was accomplished by passing the mixture through a Dowex column, AGl-X8 (CI-), (0.7 x 4 cm; Biorad Laboratories, Richmond, CA). The column was washed with approximately 3 column volumes of H20 and glucose-6-phosphate was eluted in 5 ml of 1 M LiCl2 directly into a scintillation vial. To this vial, 10 ml of Aquasol was added and counts were obtained in a Beckman LS8000 liquid scintillation counter. Cpm were converted to dpm by comparison to a standard quench curve (external standard vs. percent efficiency). Results are expressed as moles glucose-6-phosphate

formed/jig cells (dry weight).

Assay for Enzyme I. To detect this enzyme, the technique of Saier et al. (64) was used. The reaction mixture was composed of 40 mM tris(hydroxymethyl)-aminomethane hydrochloride (Tris-HCl), pH 7.5, 8 M MgCl2, 10 mM NaF, 2 mM sodium pyruvate, 0.2 mM phosphoenol[l-14C]pyruvic acid, cyclohexylammonium salt (10 iCi/~mole) plus cell extract. The total reaction volume was 100 Pl. All components except the radioactive substrate were incubated at 37 C and the reaction was begun by the addition of the substrate. Incubation continued for an additional 60 min after which time 0.4 ml Sigma color reagent (Stock no. 505-2; 20 mg/100 ml of 2,4-dinitrophenylhydrazine in 1 N HCl) was added.
The tubes were mixed thoroughly and incubated for an additional 10 min






44



at 37 C. The derivatized radioactive product was separated by the addition of 1 ml ethyl acetate. After vigorous mixing, a 600 p1 sample was removed from the organic phase,placed in 5 ml Aquasol,and counted in a Beckman LS8000 liquid scintillation counter. Results are calculated per 600 pl product counted and are expressed as pmoles 14C-pyruvate formed/pg protein.

Growth curves. Cells were pre-grown in DM supplemented with 5 mM sugar. At log-phase, the cells were transferred to fresh medium supplemented with sugar(s) at a concentration of 5 mM/sugar. Growth was monitored with a Klett-Summerson photoelectric colorimeter using a red filter #66 (Klett Mfg. Co., Inc., New York, NY) and/or a Gilford 2600 spectrophotometer (600 nm).

Sugar determination in spent medium. Lactose was determined by the Boehringer-Mannheim lactose/galactose kit (Boehringer-Mannheim Biochemicals, Indianapolis, IN). The basic principle of this kit is to convert the lactose to galactose plus glucose by -galactosidase. The galactose is oxidized to galactonic acid by galactose dehydrogenase plus NAD+. The appearance of NADH is detected at 365 nm. A 100 pl aliquot of the spent medium was incubated with NAD+ plus 1.2 U of 8-galactosidase for 10 min at 25 C. After this incubation, PB, pH 6.8, and H20 was added to bring the volume to 1.34 ml. The solutions were first read at 365 nm to obtain a background and the reaction was then started by the addition of 0.4 U of galactose dehydrogenase. Incubation proceeded for 15 min. (The reaction was determined to be complete at the end of this time period.) The final volume of the reaction mixture was 1.36 ml, the NAD+ concentration was 0.45 M and the PB concentration was 0.15 M. Controls






45


lacking B-galactosidase demonstrated the absence of galactose from the medium as a contaminant or as an excreted end-product. The concentration of lactose was determined by comparison to the reduction of NAD+ by a known quantity of lactose.
Fructose was determined by the Boehringer-Mannheim glucose/fructose kit. This kit contains hexokinase plus adenosine 5'-triphosphate (ATP) to convert fructose to fructose-6-phosphate, phosphoglucose isomerase to convert fructose-6-phosphate to glucose-6-phosphate, and glucose-6phosphate dehydrogenase plus 8-nicotinamide adenine dinucleotide phosphate (NADP ) to convert glucose-6-phosphate to gluconate-6-phosphate plus NADPH (reduced). The results were read at 365 nm and the amount of fructose calculated from a standard. Controls lacking phosphoglucose isomerase demonstrated the lack of glucose in the samples tested. The procedure involved incubating 100 pl of the spent medium with 1.0 M

NADP 4.0 M ATP, 1.33 U hexokinase, and 0.67 U glucose-6-phosphate dehydrogenase. The reaction mixture was incubated for 15 min at 25 C after which time 3.27 U of phosphoglucose isomerase was added to all but the controls and incubation was continued for an additional 15 min. The results were obtained by reading the assay at 365 nm in a Gilford 2600 spectrophotometer.

The glucose determination was based on the glucose oxidase method of Raabo and Terkildsen (50). All reagents were from a diagnostic kit (Sigma Chemical Co.). Briefly, glucose is oxidized to gluconic acid plus H202. The presence of H202 is detected by its reaction with o-dianisidine which, when oxidized, becomes brown and can be detected at 450 nm. Typical samples (0.1 ml) contained up to 15 ig of glucose.






46


To this 1.0 ml of the reagent was added. This reagent contains 5 U/ml glucose oxidase, 1 Purpurogallin U/ml of peroxidase, and 4 ig/ml of o-dianisidine. After mixing, the reactants were incubated at 37 C in

the dark for 30 min.
Membrane preparation. The procedure used is a modification of the one developed by Siegel et al. (75). Typically, cells were grown in 200 ml of either DM or TYE plus 20 mM glucose. At the log-phase, the cells were harvested and washed 2 x in 0.9% NaCl, 2 x in 5 mM ethylenediaminetetraacetic acid(EDTA), and 2 x in 20 mM Tris-HCl, pH 6.8, plus 5 mM 2-mercaptoethanol and 10 mM MgCl2("lysis buffer"). The cells were frozen at some point during the washing procedure. For lysis, cells were suspended in 50 ml lysis buffer with 5 mg purified mutanolysin (MI; a gift from Kanae Yokagawa, Dainippon Chemical Co., Tokyo, Japan) and incubated 60 min at 37 C; after which time, 2.5 mg RNase and 2.5 mg DNase (Sigma Chemical Co.) were added and incubation continued with stirring for an additional 60 min at 37 C. Since it was determined that the DNase preparation contained proteolytic activity, phenylmethylsulfonyl fluoride (Sigma Chemical Co.) was included in this last step. Membranes were collected by centrifugation at 30,900 x q for 60 min and washed once in 100 mM PB, pH 7.0, containing 5 mM MgCl2 and 5 mM 2-mercaptoethanol. The supernatant from the first and second centrifugation were pooled, dialyzed for 48 h against H20 (with at least 3 changes), lyophylized, and resuspended in 25 ml of 100 mM PB, pH 7.0, containing 5 mM MgCl2 and 5 mM 2-mercaptoethanol. (This will be referred to as the cytoplasmic fraction.) The cytoplasmic fraction was stored at
-4 C. The pellet was resuspended in the same buffer and subjectedtoalow






47



speed spin at 1075 x q for 5 min in order to remove whole cells. The supernatant from this step was then centrifuged at 30,900 x q for 60 min to recover the membranes in the pellet. The purified, cell-free membranes were resuspended in 2.5 ml of the above buffer and stored at

-4 C. These were free from contamination by cell wall (75).

Cell wall-membrane complexes were prepared using the method of Bleiweis et al. (2). For this procedure, cells were grown to earlystationary phase in 200 ml DM supplemented with 20 mM glucose. They were harvested and washed twice with 100 mM PB, pH 7.0, containing 5 mM MgCl2. The washed cells were then resuspended in 10 ml of this buffer plus an approximately equal volume of glass beads and 100-200 pl of tributyl phosphate, and homogenized for 3 min in a Braun tissue homogenizer (Bronwill Scientific, Rochester, NY) cooled by CO2. The beads were removed by filtration through a scintered glass filter (coarse). During this process, the beads were washed with several volumes of buffer. The final suspension volume was 200 ml. DNase, 10 mg, and RNase, 10 mg, were added and enzymatic digestion was allowed to proceed for 2 h at 37 C under conditions of constant stirring. EDTA, 3 mM, was added and the membranes were pelleted out and washed 1 x in 100 mM PB, pH 7.0, containing 3 mM EDTA, and finally resuspended in 100 mM PB, pH 7.0, containing 5 mM MgCl2. This was then subjected to a 1075 x g centrifugation for 5 min and the membranes were collected by centrifuging the supernatant at 30,900 x g for 60 min. The "Braun-membranes" were stored at -4C in 2.5 ml 100 mM PB, pH 7.0, containing 5 mM MgCl2.

A third method of cell breakage involved homogenizing the cells

in a Bead Beater (Biospec Products, Bartlesville, OK). This apparatus






48


is analogous to a Waring Blender except that it requires glass beads. One liter of cells grown in DM plus 20 mM glucose was harvested at early-stationary phase. After harvesting and washing twice, they were resuspended in 100 mM PB, pH 7.0, containing 5 mM MgCl2 and 5 mM 2-mercaptoethanol and homogenized using a 200-300 ml volume of glass beads for 5 min. To prevent foaming, tributyl phosphate was added prior to homogenizing. After filtering through scintered glass (coarse) to remove the glass beads, the suspension was treated with DNase and RNase, 50 pg each, for 2 h at 37 C with constant stirring. The cell wallmembrane complexes were then collected and washed twice in 100 mM PB, pH 7.0, containing 3 mM EDTA; whole cells were removed by centrifugation and the cell wall-membrane complexes were pelleted and resuspended in 5 ml of 100 mM PB, pH 7.0, containing 5 mM MgCl2.

Purification of mutanolysin. The purified fraction of mutanolysin, Ml, retained proteolytic activity; therefore, it was necessary to further purify this enzyme. This was accomplished by a two-step procedure

(75). The enzyme (15-16 mg) was suspended in H20 using sonication and then centrifuged at 30,900 x I for 1 h. The supernatant was applied to a carboxymethyl Sephadex C-25 ion exchange column (30 x 0.9 cm; Pharmacia Fine Chemicals, Piscataway, NJ) and eluted with a linear gradient of 0.01 M to 0.15 M phosphate buffer, pH 7.0, (600 ml). The column had been prewashed with 0.01 M PB, 0.15 M PB, and reequilibrated with the starting buffer. Fractions of approximately 5 ml were collected by gravity at a flow rate of 12-15 ml/h. Muralytic activity eluted at 0.045 M to 0.05 M phosphate buffer. Fractions were read at 280 nm in a Gilford 2600 recording spectrophotometer. Those that had an absorbancy






49


of greater than 0.05 were pooled and concentrated by pressure dialysis. The protein recovery was approximately 50% of starting material. Removal of protease activity was monitored by the Azacoll assay (CalbiochemBehring, La Jolla CA). The pooled, concentrated material, 125 1g, was incubated with the chromogenic substrate, Azacoll, 10 mg, in 50 mM PB, pH 7.0, to a total volume of 2.5 ml. After 2 h at 37 C, the substrate was centrifuged at 12,000 x g for 10 min and the supernatant was read against a blank tube which had contained Azacoll but no enzyme. That the purification procedure had successfully removed protease activity was determined by an absence of solubilized chromogen.
Mutagenesis and mutant selection. A modification of the methanesulfonic acid ethyl ester (EMS; Sigma Chemical Co.) mutagenesis procedure developed by Shanmugam and Valentine (74) was used to mutagenize a culture of S. mutans. Cells were grown in DM plus 20 mM glucose. At early-log phase (Klett=34), 0.1 ml of EMS was added and the incubation of cells continued for 60 min at 37 C. At the end of this time period,

the cells were harvested at ambient temperature and washed once with carbon-free DM and sonicated for 30 sec. The volume was brought up to 10 ml with carbon-free medium and the treated cells were incubated at 48 C for 40 min. The cells were pelleted at ambient temperature and resuspended in a small volume of TYE before sonicating for 30 sec. The volume was brought up to 10 ml with TYE supplemented with 20 mM glucosamine and the culture was incubated until turbidimetric increases, presumed to be growth, resumed. At this time, streptozotocin was added

to select for glc PTS mutants according to the procedure of Lengeler

(33). Streptozotocin was added so that the final concentration of this






50


glucose analogue was 50 pg/ml and incubation at 37 C proceeded for 5 h. The cells were harvested, washed once with carbon-free medium and then allowed to grow overnight in TYE supplemented with 20 mM lactose. The cells were then transferred to fresh TYE supplemented with glucosamine and at early-log phase (Klett= 35) streptozotocin was added to give a final concentration of 50 ug/m]. After 5 h at 37 C, the cells were harvested, washed once with carbon-free medium, and plated on TYE supplemented with 20 mM lactose plus 20 mM 2-DG. After 48 h, the plates were replicated on TYE containing 20 mM glucosamine or 20 mM lactose. Colonies that grew on lactose but not glucosamine were patched onto lactose/2-DG containing TYE agar. These plates were finally replicated onto TYE plates containing either 20 mM glucosamine or 20 mM lactose. Colonies were chosen which grew only on lactose. These colonies were grown in TYE broth plus 20 mM mannitol and the resultant culture was stored in glycerol at -40 C.

Dry weight determination. Cells were grown in DM plus 50 mM lactose. At various times during their growth cycle, 30 ml portions were removed, harvested, and resuspended in 30 ml, 100 mM PB, pH 7.0,

containing 5 mM MgCl2. The absorbancy of this suspension was read at 600 nm in a Gilford 2600 recording spectrophotometer. The cells were collected by centrifugation and resuspended in 3.0 ml of the same
buffer. A portion, 0.95 ml,was aliquoted in triplicate to predried (72 h, 60 C) and preweighed aluminum weighing pans. Dry weight (60 C) readings were taken at 24, 48, and 72 h. The readings did not vary and therefore an average value for all three time periods was obtained; and from this average the weight of buffer alone was subtracted to obtain a mean dry weight/absorbancy unit.






51



Chemical analysis. The origin of the sugars routinely used

throughout this study was Sigma Chemical Co. for 2-deoxyglucose (2-DG), mannitol, a-methylglucopyranoside (a-MG), methyl--D-thiogalactopyranoside (TMG), galactose, and lactose and Calbiochem-Behring for fructose and mannose. In order to ascertain the degree of purity of these sugars each commercial product was analyzed by gas-liquid chromatography. The sugars were converted to their alditol acetate derivatives by the method of Griggs et al. (15). The derivatives were separated on a glass column (6 ft x 2 mm) packed with 3% SP2330 on Supelcoport 100/120 mesh (Applied Science Lab Inc., State College, PA) on a Tracor model 560 gas chromatograph (Tracor Instruments Inc., Austin, TX). The initial temperature was 180 C and after an initial hold of 5 min, the temperature was raised to 240 C at 2 C/min intervals. The flow rate of the carrier gas, N2, was 20 ml/min. Detection of the peaks was by a hydrogen flame ionization detector and an Autolab Minigrator electronic digital integrator (SpectraPhysics, Santa Okra, CA) was used to quantitate the peaks.

The amounts of glucose, a common contaminant, were as follows: 2-DG, 0.521%; mannose, 0.0626%; mannitol, 0.184%; galactose, < 0.01%, lactose, 0.0015%. The anomers of fructose and a-MG derivatives have the

same retention times as glucose, therefore, quantitation was not possible. TMG was not tested.

The method of Lowry et al. (37) was used for protein determination. To solubilize membranes, 3' sodium dodecyl sulfate (SDS) was added. Proteins added during the purification of membranes (e.g., RNase, DNase, and mutanolysin) were not subtracted from the final data determinations.






52


Radioisotopes. The following radioisotopes were purchased from New England Nuclear: D-[14C(U)]-glucose (4.0 mCi/mmole); D-[14C(U)]fructose (359 mCi/mmole); methyl (c-D-[14C(U)]-gluco)pyranoside (275 mCi/mmole); D-[l,2-3H]-2-deoxyglucose (37.3 Ci/mmole); D-[l-3H(N)]mannitol (17 Ci/mmole); methyl(8-D-[methyl-14C]thiogalacto)pyranoside (54.7 mCi/mmole); D-[l-14C]-mannose (48.6 mCi/mmole); [14C(U)]-lactose (0.97 mCi/mmole); D-[14C(U)]-2-deoxyglucose (282 mCi/mmole). Phosphoenol-[1-14C]-pyruvic acid (10.6 mCi/mmole) was purchased from Amersham (Arlington Hgts., IL).











RESULTS

General conditions for assay of phosphotransferase systems. The initial objectives of this study were concerned with learning about the regulation of lactose uptake in S. mutans GS5. Therefore, many of the basic parameters for the study of the phosphotransferase systems (PT-systems) in this strain were established using the lac-PTS as a model. For these studies, two assays were employed: (1) ONPG plus PEP and ONPG-6-P were used as substrates for the PTS and for phospho-Bgalactosidase, respectively, and (2) the generation of pyruvate from the donation of the phosphoryl group of PEP to lactose was measured by the oxidation of NADH in the presence of lactate dehydrogenase (LDH). Since there is a stoichiometric relationship between lactose and PEP and between PEP and NADH, the amount of NADH oxidized is a measure of the amount of lactose phosphorylated.

Fig. 3 demonstrates the linearity of the two PTS assays. Up to 625 pg dry weight of cells may be used to obtain a linear relationship in the utilization of ONPG plus PEP under the conditions outlined in Methods. On the other hand, the oxidation of NADH is linear up to 325 pg of cells, after whicha plateau is reached. The measurement of NADH oxidation is approximately 70-fold more sensitive than the phosphorylation and subsequent cleavage of ONPG as determined by comparison of the results obtained by the two different assays using the same dry weight measurement of cells. This may reflect the relative affinities of the lac PTS for ONPG vs. lactose. ONPG, an analogue of lactose, would be expected to have a lesser affinity than the natural substrate.

53




















Fig. 3. Lactose phosphotransferase system activity as a function of cell concentration. Cells were decryptified as described in Methods except the toluene-acetone mixture was 50i1/ml cell suspension. ONP released by the ONPG + PEP reaction ( ) and NAD generated by the LDH/NADH linkedspectrophotometric assay ( o ) were measured as described in Methods.







55

























NADH (umoles) CONVERTED TO NAD+/ min (o)

0 0 0 0 0 U ) (\J

0
0 (.D

0

.0
to





0C~
0
-OJ Lii
0

0 (0





(s(f Owi f) f i3SV 13t t dNO





56



Fig. 4 demonstrates that the pH optimum of the lac PTS is 7.0

using either of the two reaction systems described above. This optimum is dependent on the buffer system used. Table 2 shows the results of assaying for the lac PTS using the two different methods and phospho8-galactosidase in the presence of various buffers. Phosphate buffer at pH 7.0 is optimal for both PTS and phospho-B-galactosidase activities.

Lac PTS activity appears to be independent of ionic strength

between 10 and 500 mM sodium phosphate. This can be seen in Table 3. Also, phospho-8-galactosidase activity is most pronounced at ionic strengths between 50 and 1000 mM. On the other hand, NAD+ production is independent of all ionic strengths employed.

Assaying whole cells required the permeabilization of membranes

to phosphorylated compounds such as PEP using a toluene-acetone mixture. The rationale for using "decryptified" cells was to allow for controlled intracellular concentrations of reactants. Since this type of treatment may cause alterations of the membrane proteins, it was necessary to standardize the conditions of this procedure. Decryptification involves the addition of the toluene-acetone mixture followed by vigorous shaking of the cell suspensions. Table 4 outlines a variety of parameters followed during this step. Two proportions of toluene with acetone were examined: 1:3 and 1:8 (v/v). The amount of this mixture per cell volume was considered as well as the extent of mixing using a Vortex Mixer. All shaking was done at 2-min intervals with approximately 1-min cooling

in ice.
A toluene-acetone mixture of 1:3 (v/v) at a final concentration of 100 il/ml cell culture allowed for the highest lac PTS activity. The

























Fig. 4. The pH optimum for the lactose phosphotransferase system.
Cells were grown and decryptified as outlined in Table 2. A cell concentration of 120 pg (dry weight) was used to measure ONP released by the ONPG + PEP reaction ( 0 ) and NAD+ generated by the LDH/NADHlinked spectrophotometric assay ( 0 ) at indicated pH levels in 60 mM PB (ONP assay) and 70 mM PB (spectrophotometric assay).









co
L0l







NADH (nmoles) CONVERTED TO NAD+/ug CELLS (o)
0 0 0 0 0 0 0 0 0 0 0 .- o o oo t, o o o o oo a O










0
o o I ,
0


() S-1-130 b6t/(SO wu) dNO






59


Table 2. Lactose phosphotransferase system and phospho--galactosidase
enzyme activities as a function of buffer composition and pH.

nmoles ONP/Ijg cellsb
Buffer pH ONPG + PEP ONPG-6-P unoles NAD+/min/pgb cells

MESa 5.0 .09 1.38 0
6.0 .43 2.10 27.9 6.5 .71 4.44 47.9 PBa 6.5 .66 4.64 45.0
7.0 .86 5.25 75.7
a 7.5 .81 4.48 67.9 MOPS 7.0 .78 4.23 62.1 TRIS-HCla 7.5 .72 1.80 39.3
8.0 .59 1.46 45.0 9.0 .44 1.11 8.5 10.0 .21 0.18 0

aMES: 2-(N-morpholino)ethanesulfonic acid, sodium salt;
PB: sodium phosphate; MOPS: 3-(N-morpholino)propanesulfonic acid, sodium salt; Tris-HC1: tris-(hydroxymethyl)-aminomethane hydrochloride.
bCells were grown to late-log/early-stationary phase in defined

medium containing 5.0 mM lactose, harvested, washed 1 x with 100 mM PB, pH 7.0, containing 5 mM MgCl2 and then resuspended in 100 mM PB, pH 7.0,
containing 5 mM MgCl2 to a final volume 1/20th of the original culture. The cells were decryptified with a toluene-acetone mixture at a final ratio of 50 pl/ml of cells for 5 min. All reactants were made up in H20. The final concentrations of the above buffers were 70 mM for the LDH/NADHlinked assay, 60 mM for the ONPG + PEP assay, and 80 mM for the ONPG-6-P assay. Since cells were suspended in 100 mM PB, pH 7.0, the contribution of this buffer was minimized by using a 2-fold suspension of cells and
one-half the routine volume of cell-suspension. This results in a 2.5 mM concentration of PB in the LDH/NADH assay, 3.8 mM in the ONPG + PEP assay, and 1.8 mM in the ONPG-6-P assay. The concentration of cells was 140 pg, dry weight, for the LDH/NADH-linked and ONPG assays and 56 jg, dry weight, for the ONPG-6-P assay. MgCl2 concentrations were 3.5 mM for the NADH/
LDH-linked assay, 3.0 mM for the ONPG + PEP assay, and 4.0 mM for the ONPG-6-P assay. See Methods for details of each enzymatic assay procedure.






60









Table 3. Lactose phosphotransferase system and phospho--galactosidase
activities as a function of ionic strength of buffer reagent.


moles ONP/ipg cells
maa ONPG + PEP ONPG-6-P moles NAD+/min/pg cellsb



1000 .091 4.18 20.9 500 .175 3.82 20.5 100 .138 6.90 22.2 50 .166 5.72 24.0 10 .168 1.70 14.3


apB, pH 7.0.

bThe procedures used are as outlined in Table 2.






61







Table 4. Determination of optimal amounts of solvents for decryptification of cells for phosphotransferase enzyme assays.a

Toluene-acetone (v/v) 1:3 1:3 1:3 1:3 1:3 1:3 1:8

Pl/ml 100 100 100 100 50 25 100 Vortex time (min) 0 2 4 5 5 5 5

ONPG + PEPb .534 .981 .913 .816 .515 .515 .476 ONPG-6-Pc 21.7 25.7 26.9 26.9 35.0 28.1 36.2


aCells of S. mutans GS5 were cultured overnight in defined
medium containing 5.0 mM lactose, harvested, and washed as described in Methods.
bCells, 103 pg (dry weight), were incubated with 100 mM PB, pH 7.0, 5 mM MgC12, 8 mM PEP, and 8 mM ONPG (total volume: 600 pl) for 30 min at 37 C. The reaction was stopped with cold 5% NaCO3 and the cells were removed by centrifugation. The amount of ONP formed was determined by spectrophotometric measurements of the supernatants at 420 nm. Results are expressed as nmoles ONP/pg cell (dry weight).
CCells, 21 pg (dry weight), were incubated with 100 mM PB, pH 7.0,
5 mM MgCl2, and 100 mM ONPG-6-P (total volume: 500 pl) for 30 min at 37 C. The reaction was determined as above and is expressed as nmoles/g cells (dry weight).






62


permeabilization appeared to be complete after 2-min of mixing. If phospho-6-galactosidase activity is measured a somewhat different profile is obtained; optimization occurred at a 50 il/ml cell culture of a 1:3 (v/v) mixture or a 100 vl/ml cell culture of a 1:8 (v/v) mixture. Since the emphasis of the present study is on the various PT-systems, the optimum conditions were chosen on this basis.

Table 5 compares sugar transport into intact cells vs. phosphorylation of that sugar by decryptified cells. As can be seen with both fructose and glucose, transport results in a 2-fold greater amount of sugar being converted to its phosphorylated derivative compared to phosphorylation. This could be due to one of several factors. First, treatment of the membrane with toluene-acetone may result in an inactivation and/or rearrangement of the transport proteins thereby preventing maximal uptake of sugar for phosphorylation. Second, two glucose transport systems have been shown to be operative in this organism (18). Part or all of this higher transport activity may be due to a contribution of this second system. Since this latter system requires a proton motive force, it can be observed with intact cells only. Also, the presence of an ATP-dependent kinase (see Table 6) indicates an alternate fructose system; therefore, the above discussion concerning glucose transport may pertain to fructose transport. Lastly, this assay was performed with a radiolabelled sugar substrate which is converted to a labelled phosphorylated derivative. The product is separated from the reactant by an anion exchange filter which will trap the negativelycharged sugar phosphate or, alternatively if intact cells are used, the negatively charged cells. These filters may be more efficient for






63



Table 5. Comparison of sugar (glucose and fructose) transport and
phosphorylation by decryptified or untreated cells.a

Substrate
Glucose Fructose

(nmoles/ml)

Sugar transportedb 2.4 1.5 Sugar phosphorylatedc 1.2 0.7

aCells were grown in fructose according to the procedure outlined in Methods. At early-stationary phase, the cells were harvested and washed 1 x and finally resuspended to 4 x their original concentration. One-half of this suspension was decryptified with a toluene-acetone mixture as described previously and the other half was untreated.
bFor transport assays, untreated cells (115 ig) were incubated with 40 mM PB, pH 7.0, 2 mM MgCl2, 10 mM NaF, and either 100 yM D-glucose plus 125 pM D-[14C(U)]-glucose (4 vCi/pimole) or 100 V M D-fructose plus
1 ~i D-[14C(U)]-fructose (359 pCi/pmole).
CDecryptified cells (113 pg) were used to test for sugar phosphorylation. The conditions were alteredfrom the transport assay in the following manner: 10 mM PEP was added and incubation was in 50 mM PB,
pH 7.0, plus 2.5 mM MgCl2. The same amountsof radiolabelled substrates were used.

In both types of reactions, incubation was carried out at 37 C for 10 min after which time 0.2 ml was removed and diluted into 2 ml of 1% cold sugar. These were then passed through a DE-81 filter and the filters counted in a liquid scintillation counter. Raw counts were then converted to dpm and nmoles/ml were calculated from known specific activities.






64


binding the cells than the product, thus, the higher apparent transport activity may be partly due to an inherent characteristic of the assay.
Even though sugar transport appeared to lend itself to a more sensitive assay, I chose to study sugar phosphorylation using decryptified cells. The reasons for this were as follows: (1) the concentrations of the reactants could be controlled more carefully, and (2) the use of decryptified cells eliminated any secondary transport systems (e.g., via the proton motive force) that may be present.

Fig. 5 profiles the lac PTS and phospho--galactosidase activities during the growth curve described by strain GS5 when grown on this sugar. It is evident that the lac PTS reaches maximum activity at early-stationary phase. This was confirmed by the LDH/NADH assay
(data not shown). The catabolic enzyme, phospho-8-galactosidase, however, reaches maximal levels of activity at early-log phase. This maximization of phospho-a-galactosidase appears to coincide with the early detection of the lac PTS. In all subsequent experiments involving the several PT-systems, late-log or early-stationary cells were used.

Phosphorylation of sugars by the various PT-systems (e.g., glc PTS) is dependent on PEP by definition; ATP will not substitute for PEP. This is clearly seen in Table 6 in the case of glucose. However, ATP was able to donate its phosphoryl group to fructose and this reactionprovided 40% the amount of fructose-6-phosphate formed as when PEP was the donor. Mannose was also able to accept a phosphoryl group, but
only 8% of product formed as compared to assays using PEP as a phosphoryl source. This organism has been shown to contain a fructokinase
which is able to catalyze mannose phosphorylation (5). Such an enzyme



















Fig. 5. Lactose phosphotransferase system enzyme activities and growth of S. mutans GS5 as a function of time. Cells were grown in defined medium plus 5.0 mM lactose over a period of 7 h and turbidities measured using the Klett ( 0 ). At defined intervals, aliquots were removed and cells decryptified for the ONPG + PEP assay releasing ONP ( 0 ) and the ONPG-6-P assay releasing ONP ( a ). The latter assay measures phospho-6galactosidase activity.






66




ONP (nmoles) RELEASED/ug CELLS FROM ONPG-6-P (A) ONP (nmoles) RELEASED/ug CELLS x I0-' FROM ONPG+PEP(o) co oD N 0 c ( N








O



0 .




o



0




0









I I I0
O O 0 0 0 0

(') S.IINN 1L1)7






67







Table 6. Determination of phosphoenolpyruvate dependency for
phosphotransferase system-mediated phosphorylation.

Substrates for phosphorylation
Glucose Fructose Mannose

ATP 0a 0. 5a 0.06b PEP 2.9a 1.2a 0.77b

aCells grown in defined medium plus 5 mM glucose were harvested, washed, and decryptified. The permeabilized cells, 153 ug (dry weight) were incubated with 70 mM PB, pH 7.0, 3.5 mM MgC12, 10 mM ATP or PEP, 10 mM NaF and 100 0M D-glucose plus 125 pM D-[14C(U)]-glucose (4 pjCi/ inole) or 100 -M D-fructose plus 1 pM D-[14C(U)1-fructose (359 Ci/jimole). The final reaction volume was 1.0 ml.
bCells grown in defined medium plus 5 mM glucose were harvested, washed, and decryptified. The permeabilized cells, 109 lig (dry weight) were incubated with 70 mM PB, pH 7.0, 3.5 mM MgC12, 10 mM ATP or PEP, 10 mM NaF, and 100 pM D-mannose plus 10 pM D-[1-14C]-mannose (48.6 pCi/ vpnole). The final reaction volume was 1.0 ml.

In both a and b, the reaction was allowed to proceed for 10 min at 25 C. Reactions were halted when 0.1 ml was removed and diluted in 1% homologous sugar, filtered through a DE-81 filter and counted in a liquid scintillation counter. Results are expressed as nmoles sugar phosphorylated/ml.






68



is likely responsible for the small amount of phosphorylation seen here. The fructokinase is present in glucose-, mannose- and fructose-grown cells (data not shown). In all experiments performed in this study PEP was used, however, the possibility exists that some PEP was converted to ATP via pyruvate kinase. All results using fructose as a substrate are interpreted as being at least 60% dependent on the PTS. Fig. 6 demonstrates that at the concentration used in this study, 10 mM, PEP is well beyond the limiting range. Resting cells of S. lactis have been shown to contain an energy reserve in the form of 3-phosphoglycerate

(86). Presumably, S, mutans cells in early-stationary phase contain a similar reserve since transport in intact cells is observed (Table 5). If NaF is excluded, glucose phosphorylation occurs in decryptified cells without the addition of PEP (data not shown). Reproducibility of this result was not always obtained, an observation confirmed by others (Dr. G. Jacobson, personal communication). Therefore, for purposes of standardization, NaF was always included and the concentration used, 10 mM, gave maximum inhibition of glucose phosphorylation in the absence of PEP (data not shown).

Comparative study of phosphotransferase-mediated sugar phosphorylation. In order to obtain a profile of the various PT-systems in this organism, a survey was set up in which the cells were grown in various growth substrates and assayed for the presence of specific PT-systems. The results are outlined in Table 7. The glc PTS and man PTS can be detected under all the growth conditions tested leading to the conclusion that these two sugars are phosphorylated via a constitutive system. Phosphorylation of 2-deoxyglucose (2-DG) was detected in all cells

















Fig. 6. Glucose phosphotransferase system activity as a function of phosphoenolpyruvate (PEP) concentration. Cells were grown in defined medium plus 5 mM glucose and decryptified as in Methods. Cells(63 jig dry weight), were incubated with varying concentrations of PEP plus 25 IIM D-[14C(U)]-glucose (4 uCi/Fmole) and 100 ipM D-glucose in 25 mM PB, pH 7.0, containing 1.3 mM MgCl2. Product (glucose-6-phosphate) was measured by filtering cells on DE-81 filters and counts retained on filters were measured and converted to dpm and subsequently to nmoles/ml reaction mixture.





















GLUCOSE-6-PHOSPHATE (nmoles/ml) 0o C -N






.0
b






m


3o
0 0 0

O 0




0










OL






71







Table 7. Induction of phosphoenolpyruvate-dependent phosphotransferase
systems as a function of carbon source in growth media
(LDH/NADH-linked assay).

Growth carbon source
Substrate testedb Glucose Mannose Mannitol Lactose Galactose

Glucose 6.3 8.1 12.7 1.5 18.3 Mannose 4.5 4.9 4.8 1.9 15.7 Mannitol 0 0 3.0 0 0 2-Deoxyglucose 0.9 0.9 2.0 0 9.4 a-Methylglucoside 0 0 0 0 0 Galactose 0 0 0 0 0
Isopropyl -Dthiogalactopyranoside 0 0 0 1.4 3.9 Lactose 0 0 0 7.5 20.7 Glucosamine 5.5 N.T.c N.T. N.T. N.T.


aCells were grown in the several sugars as described under Methods. After harvesting and washing, they were resuspended in buffer to 1/20th of their original volumes. The cell concentration varied between 60-325 pg/ml. To insure linearity, at least two cell concentrations were used to test a given sugar for sugar phosphorylation. Where activity could not be detected the upper limits of this range (240-325 pg) were reported.
bThe LDH/NADH-linked spectrophotometric assay was used to assay for sugar phosphorylation as described in Methods. All data are expressed as imoles NADH converted to NAD+/g cells/min x 10-2.
CNot tested.






72


except those grown in lactose. This compound is a glucose analogue and thus would be expected to be transported via the glc PTS as is the case in E. coli (21). Schachtele and Mayo (71) showed that the uptake and phosphorylation of this analogue in S. mutans is through a PTS, most likely the glc PTS. The detection of PEP-dependent phosphorylation of this sugar analogue in all but lactose-grown cells argues for such a PTS being constitutive. The most likely reason for its absence in lactose-grown cells is that low levels of the glc PTS preclude the detection of 2-DG phosphorylating activity given the sensitivity of the assay used. In agreement with Schachteleand Mayo (71), mannitol-grown cells show a high level of 2-DG activity; again, most likely because glc PTS activity is high in these cells.

One means of testing for the presence of two glc PT-systems is to compare the phosphorylation of the two glucose analogues, 2-DG and a-MG

(21). It can be seen (Table 7) that a-MG is not phosphorylated, thereby leading to the tentative conclusion that there is only one glucose system operating in this organism and it would be analogous to the low affinity system in E. coli. Mannitol, as has been shown by others (40), is phosphorylated by an inducible system. Interestingly, galactose is not an apparent PTS substrate even though growth in this sugar leads to a high level of other PT-systems. This may be related to the poor growth of the cells in this carbon source. Lactose and its analogue, isopropyl-D-thiogalactopyranoside (IPTG) appeared to be phosphorylated by an inducible system. This is in agreement with a report published by Hamilton and Lo(17). Galactose as well as lactose appear to be inducers; in S. aureus, galactose-6-phosphate has been shown to be the inducer for the lac PTS (43).






73


The results obtained in Table 8 reaffirm the general pattern

discussed above. Here the cells were grown as in Table 7 but the PTS assay was conducted using radiolabelled substrate at a saturating level. Glucose, again, appears to be constitutive. The relative levels are similar to those found using the LDH/NADH assay. Glc PTS is low in lactose-grown cells and high in mannitol-grown cells. However, there

appears to bea contradiction with mannose-grown cells in that PEPdependent glucose phosphorylating activity is lower than anticipated. In agreement with the previous results, 2-DG is phosphorylated while ac-MG is not. Mannose phosphorylation does not vary greatly between mannose- and glucose-grown cells but shows a significant decrease in lactose-grown cells. Fructose, like glucose and mannose,appears to be phosphorylated by a constitutive system.

The constitutive nature of PEP-dependent phosphorylation of glucose, mannose, and fructose suggests a possible physiological relationship amongst the PT-systems for these three sugars. As discussed in the Introduction, the low affinity glc PTS in E. coli phosphorylates glucose, mannose, fructose, and glucosamine (31). In addition, this system phosphorylates 2-DG but not ac-MG. Since it appeared that an analogous system existed in this strain of S. mutans, a series of competition experiments was done to test for this possibility. In these experiments, a 14C-substrate competed for phosphorylation against an excess of unlabelled sugar. The results are presented in Table 9. Glucose and mannoseweremutually competitive. Mannose was a less efficient competitor for glucose phosphorylation than was glucose. Glucosamine was also a competitor for both mannose and glucose but at a lower degree of






74






Table 8. Induction of phosphoenolpyruvate-dependent phosphotransferase
systems as a function of carbon source in growth media
(radioactive assay).

Growth carbon source a
Substrate testedb Glucose Mannose Mannitol Lactose

(pmoles sugar-phosphate/vg cell dry weight) Glucose 28.7c 18.5 95.8 38.6 Mannose 44.0 30.5 N.T. 6.4 2-Deoxyglucose 0.9 N.T. N.T. N.T. ac-Methylglucoside 0 N.T. N.T. N.T. Methyl-8-D- d
thiogalactopyranoside N.T. N.T. N.T. 3.5 Mannitol N.T. 0 12.3 N.T. Fructose 56.1 20.0 49.3 4.8

aCells were grown as described previously (Table 5, Methods).

bCells were decryptified and assayed with various sugars. The

final concentrations of reactants were: 30 mM PB, pH 7.0, 1.5 mM MgC12, 10 mM PEP, and 10 mM NaF. The substrate concentrations included 100 pM of unlabelled sugar containing the following concentrations of the homologous radioactive substrates: 50 pM D-[14C(U)]-glucose (4 pCi/pmole),
3 pM D-[1-14C]-mannose (48.6 pCi/pmole), 0.03 pM D-[1,2-3H]-2-deoxyglucose (37.3 mCi/pmole), 0.08 pM methyl-a-D-[14C(U)]-glucoside (275 Ci/pmole), 19 pM [methyl-14C]-a-D-thiogalactopyranoside (54.7 pCi/pmole), 0.006 pM D-[l-3H(N)]-mannitol (17 mCi/ijmole) or 2 M D-[14C(U)]-fructose.(359pCi/pmole) Reaction mixtures included a range of 12-350 pg cells and specific activities were calculated from an average of those values falling within the linear portion of the curves generated (not shown).
cThe standard deviation for 5 identical samples was + 3.8.
dNot tested.
















Table 9. Competitive inhibition by unlabelled sugars of uptake of radiolabelled
sugars by S. mutans GS5.a




Radiolabelled substratesb Glucose Mannose Glucosamilne c-Methylglucoside Fructose Mannitol Methyl-6-Dgalactopyranoside
(% Inhibition)

D-[l-14C]-Mannose 96 93 67 N.T.c 39 N.T. N.T. 0-04C(U)]-Glucose 99 74 54 22 1 0 8

*89 20 0 0 0 0 0 D-[14C(U)]-2-Deoxyglucose 94 95 86 N.T. 28 N.T. 17

D-[14C(U)]-Fructose 0 25 0 0 99 0 0

aCellswere grown in defined medium plus 5.0 mM glucose and prepared as described in Methods.
bThe assay contained 30 mM PB. p1H 7.0; 1.5 mM MgC12; 10 mM PEP; 10 mM NaF; 0, 2 (shown by asterisk), or 10 mM competing unlabelled sugars; and cells, 85-100 pg (dry weight). The 14C-sugar concentrations were: glucose, 250 14; fructose, 1.5 JM; mannose, 10.5 Iid; and 2-deoxyglucose, 3.5 M. The incubation mixture was held at 25 C for 9 min and then diluted with a cold 1% solution of the homologous sugar. This was held at 4 C until filtered through a DE-81 filter. Results are expressed as percent inhibition by 2 or 10 mM unlabelled sugar of uptake of labelled sugar.
CNot tested.









Q"






76



efficiency. Glucose phosphorylation appeared to be inhibited by ac-MG but since direct phosphorylation of this analogue could not be demonstrated, it was concluded that this observed inhibition was the result of a non-specific mechanism. As expected, fructose competed with its 14C-isotope; however, tt did not inhibit glucose phosphorylation. There was a mutual competition between mannose and fructose. This may be due to some recognition of mannose by a frc-PTS or the sharing of a distinct "mannose site" on the glc PTS with fructose. If the latter explanationis correct, affinity for fructose would be extremely low since this sugar does not interfere with glucose activity. Alternatively, this mutual reaction may reflect the action of a manno-fructokinase. It has been shown that S. mutans possesses an ATP-dependent kinase capable of recognizing both fructose and mannose (5). ATP may be generated by pyruvate kinase and this may be available in PEP-supplied cells. Table 6 indicates a role for ATP in sugar phosphorylation. However, this mechanism would not account for all the inhibition observed. As would be predicted, mannitol and methyl-B-D-thiogalactopyranoside (TMG) do not

inhibit the reactions seen with glucose or fructose.

In addition, both glucose and mannose, as well as glucosamine,

inhibit 2-DG phosphorylation. Fructose is slightly inhibitory, however TMG also shows some competitive inhibition which indicates a degree of non-specificity to the reaction. If a "mannose site" exists on the glc PTS and 2-DG shows some affinity for this site, then one may postulate that this is the site where fructose is inhibiting. Since 2-DG is contaminated with glucose (see Methods), it is difficult to interpret the data shown in Table 9 using this glucose analogue as a competitive






77


inhibitor. Calculations showed that the amount of inhibition of glucose phosphorylation could be attributable to the contaminating glucose. However, PEP phosphorylation of mannose and fructose in the presence of 2-DG showed 15% and 100%, respectively, of control levels, clearly indicating competition in the case of mannose. Also, the homologous system showed 94% inhibition.

Kinetics of phosphotransferase activities and relative growth rates in various sugars. It was of interest to deduce the relative affinities of the glc/man PTS for substrates. Kinetic analyses were performed and are illustrated in Figs. 7-9. The linear transformation by a double reciprocal plot was calculated by a linear regression analysis. The linear coefficient in the case of glucose was calculated to be .999. The Km for glucose phosphorylation was calculated from the x-intercept and was found to be 64 pm and the Vmax which was calculated from the y-intercept was found to be .366 nmoles glucose-6-phosphate formed/min/ ml (Fig. 7). For mannose, the linear coefficient was calculated to be .987. The Km for mannose phosphorylation was calculated to be 90 s and the Vmax was .300 moles mannose-6-phosphate formed/min/ml (Fig. 8). Not shown is the kinetics of 2-DG phosphorylation, the Km of this reaction was calculated to be 154 pim.

Fig. 9 shows the results of a kinetic analysis using fructose as a substrate. The Km of this reaction is 42 jm and the Vmax is .176 nmoles fructose-6-phosphate formed/min/ml. The linear coefficient in this case is .998.

These relative affinities are reflected in the growth rates in these different carbon sources (Fig. 10). A transformation of these














Fig. 7. Kinetics of glucose-6-phosphate formation by the PEPdependent phosphotransferase system of S. mutans GS5. Decryptified
cells (104 pg, dry weight) obtained from cultures of strain GS5 grown in defined medium plus 5 mM glucose were suspended in a reaction mixture
(1.0 ml total volume) of 30 mM PB, pH 7.0, containing 1.5 mM MgC12, 10 mM PEP, 10 mM NaF, and various concentrations of D-glucose each containing D-[14C(U)]-glucose (0.8 uCi/pmole). After 10 min at 25 C, reactions were stopped by dilution of 0.1 ml of the mixture into 1.0 ml 1% glucose. Reactants were separated from products by a DE-81 filter. Glucose-6-phosphate was measured ( e ) from counts converted to dpm and then expressed as nmoles/min/ml. A double reciprocal plot (inset) was determined by linear regression analysis. Kinetic data are included in the text.











120
100
80 60
-2 40 E .30 20
1 I p I I I l I
-.10 0 .10 .20 .30 .40 .50 .60 .70
0 .25- /s o *
E
c .20
w



"C
. .10


m) .05
0
0


0 100 200 300 400 500 600 700 800 900 1000 GLUCOSE (tM)














Fig. 8. Kinetics of mannose-6-phosphate formation by the phosphoenolpyruvatedependent phosphotransferase system of S. mutans GS5. Decryptified cells (103 ug, dry weight), obtained from cultures grown in defined medium plus 5 mM glucose, were suspended in a reaction mixture (1.0 ml total volume) of identical composition to
that employed in Fig. 5 except for the substitution of various concentrations of D-mannose each containing D-[l-14C]-mannose (2.3 pCi/pmole). Reactions were carried out as in the previous experiment (Fig. 5) and stopped by dilution into 1% mannose. Mannose-6-phosphate was measured ( ) from counts converted to dpm and then expressed as nmoles/min/ml. A double reciprocal plot (inset) was determined by linear regression analysis. Kinetic data are discussed in the text.












140 120 100
V 80E 60
40
E
u 20 E -.10 0 .10 .20 .30 .40



.4



0


z
a .2

I .




0 1 O0 200 300 400 500 MANNOSE (uM)





co














Fig. 9. Kinetics of fructose-6-phosphate formation by the phosphoenolpyruvate-dependent phosphotransferase system of S. mutans GS5. Decryptified cells
(100 pg, dry weight) obtained from cultures grown in defined medium plus 5 mM glucose, were suspended in the reaction mixture employed in Figs. 5 and 6, except for the use of various concentrations of D-fructose, each containing D-[14C(U)]-fructose (5.4 1Ci/ pmole). Cold 1% fructose was employed to stop reactions. Fructose-6-phosphate was measured ( ) from counts converted to dpm and then expressed as nmoles/min/ml. A double reciprocal plot (inset) was done as before and kinetic data are presented in the text.









120 100

IV 80

.15 60
40E 20 E -.10 .10 .20 .30 40 .50
0 I I/s o .10
E

O




a0 LU


0 I



LL I I u
0 100 200 300 400 500 FRUCTOSE (gM)



















Fig. 10. Growth of S. mutans GS5 in glucose, fructose, and mannose. Cells were grown for 12 h in TYE plus 20 mM concentrations of either glucose, fructose, or mannose. Cultures (5% inocula) were then transferred to 50 ml defined medium containing 5 mM concentrations of the homologous sugars. At specified times, 10 ml aliquots were removed for absorbancy measurements at 600 nm.








0.8
GLUCOSE

0.7 0.6


0.5 FRUCTOSE


0 0.4 MANNOS


0.3


0.2


0.1 I



0 60 120 180 240 300 360 420 480 540 600 660 720 780 TIME (min)







0o
Ur






86


data into a semi-log plot allowed the calculation of the mean generation

times. Themean generation time in defined medium containing glucose is 75 min; whereas in mannose it is 290 min. With mannose as the growth substrate, a maximum culture density was not achieved after 12 h of growth. However, cultures grown in glucose reach their maximum levels between 5.5 h and 6.0 h under the conditions employed.

The mean generation time using fructose as a carbon source was calculated to be 121 min and maximum growth was achieved in approximately 6.5 h.

Selection of glucose phosphotransferase negative mutants. Data presented thus far indicate that S. mutans GS5 appears to possess a relatively non-specific PTS for glucose uptake. In order to further study the former system, glc PTS negative mutants were selected on the basis of their inability to grow on glucosamine. Wild-type cells noninduced for the lac PTS are unable to grow in the presence of 2-DG

since this non-metabolizableglucose analogue is transported by a constitutive system, thereby exhausting the PEP reserves. In addition, evidence will be presented below that suggests glucose represses the induction of the lac PTS. By selecting for a 2-DG resistant mutant, one may be selecting for a glc PTS-negative mutant, since 2-DG is transported by the glc PTS. However, the majority of the 2-DG resistant clones picked appeared to possess an altered glc PTS allowing them to transport and therefore grow on glucose (data not shown). In order to select true glc PTS-negative mutants, a second criterion, the inability to grow on glucosamine was used. However, glucosamine negative cells are not necessarily transport mutants. Clones which






87


grew on lactose plus 2-DG but not on glucosamine were isolated. A priori, one would expect a mutant selection based on these two criteria to provide glc PTS-negative mutant clones. In addition, glc PTS-negative cells were enriched by incubation in streptozotocin in the step prior to plating (see Methods). After mutagenesis, the cells were allowed to recover in glucosamine. As shown in Table 7, this is a PTS substrate which is carried by the glc PTS (Table 9). Streptozotocin then was added to these cells. Since it is a glucose analogue containing a nitroso group and is carried by the glc PTS (33), those cells still containing an active glc PTS will transport this toxic analogue and should be killed.

Streptozotocin-resistant clones were chosen and allowed to grow in mannitol overnight. A survey was done to determine if the lesion indeed was in the glc PTS and the extent of loss of phosphorylating functions. In order to perform a general survey, the LDH/NADH-linked assay was used. The PTS substrates studied were glucose, mannose, fructose, and mannitol.

Glucose, mannose, and fructose were used to determine what, if any, linkage exists between the PTS-mediated phosphorylations of these three sugars and mannitol phosphorylation was assayed as a positive control, since all cells were grown on this substrate. The results which are expressed as percent wild-type activity can be seen in Tables 10-12. Table 10 shows a representative listing of clones which grouped into what was labelled as Group I. These cells could not phosphorylate glucose or mannose; whereas fructose and mannitol phosphorylations were variable but did not correlate with the absence of glucose/mannose phosphorylation. Table 11 shows representatives of Group II. This group had much reduced glucose activity but with one exception mannose






88






Table 10. Glucose phosphotransferase system negative mutants:
Group I.a


Sugar substrate for transport assay b

Clone Glucose Mannose Fructose Mannitol

3A Oc 0 175 94 2A 0 0 125 35 4B 0 0 95 20 12B 0 0 54 24 18B 0 0 42 59 16B 0 NTd 50 0 23A 0 0 26 0

aMutants were selected as outlined in Methods.

bCells were grown overnight in TYE broth plus 20 mM mannitol,
harvested, washed, and decryptified. The LDH/NADH-linked spectrophotometric assay for sugar phosphorylation was run on each clone using the indicated sugars as described in Methods. In each case a 1 mM sugar concentration was employed.
CResults are expressed as percent wild-type.
dNot tested.






89







Table 11. Glucose phosphotransferase system negative mutants:
Group II.a


Sugar substrate for transport assayb

Clone Glucose Mannose Fructose Mannitol


34B 8c 0 83 45 8B 13 0 46 48 25B 16 0 104 42 26B 18 0 86 18 4A 20 0 NT d 105 10OA 21 0 71 100 26B 21 7 129 59 28B 29 0 88 31

aMutants were selected as outlined in Methods.

bSee Table 9 for assay conditions.

CResults are expressed as percent wild-type.

dNot tested.






90







Table 12. Glucose phosphotransferase system negative mutants:
Group III.a


Sugar substrate for transport assayb

Clone Glucose Mannose Fructose Mannitol


49B 30c 21 155 97 23B 33 15 117 63 11B 34 15 75 164 13A 49 17 NTd 176 46B 38 0 158 33


aMutants were selected as outlined in Methods.

bsee Table 9 for assay conditions.

CResults are expressed as percent wild-type.

dNot tested.






91



activity is totally absent. Fructose phosphorylation was variable but always detectable. Group III clones are listed in Table 12. These cells were characterized by having up to 50% wild-type activity for glucose phosphorylation and, with one exception, residual mannose activity. Fructose activity was near to or above wild-type activity in all mutants which grouped in this category. These data suggest a g1c/man PTS and a distinct frc PTS.

Studies of cell-free membranes for phosphotransferase related

activities. The study of the glc PTS using isolated membrane fractions was undertaken for a number of reasons. The first reason was to obtain a more detailed picture of this transport system at the molecular and cellular levels. Second was to compare membranes with whole cells in order to evaluate the ability to maintain the native protein structure during the fractionation procedure; specifically when using the muralytic enzyme,mutanolysin (Ml), to remove the cell wall. Lastly, and most

significantly, was to determine the site(s) of lesion(s) of the mutants described in the previous section. The latter studies are described in the following section. Since the primary focus of this study is the glc PTS, all membranes were prepared from glucose-grown cells.

Mutanolysin (Ml) treatment yielded a more active membrane preparation than when mechanical means were used to break cells. The specific activity of glucose phosphorylation using 50 pM D-[14C(U)]-glucose by Ml-prepared cell membranes was 14.7 pmoles/pg protein whereas "membranes" obtained from cells broken by glass beads was 4.6 pmoles/pg protein. The Ml-derived membranes were 3.2-fold more active than the wall-membrane complexes obtained from cells broken by glass beads in a Bead Beater.




Full Text

PAGE 1

STUDIES ON THE PHOSPHOENOLPYRUVATE-DEPENDENT PHOSPHOTRANSFERASE SYSTEMS IN Streptococcus mutans GS5 BY ELLEN S. LIBERMAN A DISSERTATION PRESENTED TO THE GRADUATE COUNCIL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 1982

PAGE 2

ACKNOWLEDGMENTS The author wishes to express her gratitude to the chairman of her committee. Dr. Arnold S. Bleiweis, for his valuable guidance and support. She would also like to thank Drs. Richard Boyce and Francis Davis, for their counsel during these studies. She owes special thanks to Dr. Keelnatham Shanmugam whose help in obtaining mutants and with various other aspects of this work was invaluable. She would also like to show her appreciation to Steven F. Hurst for his assistance with gas liquid chromtographic analysis and for his excellent graphics and photography. Lastly, two of the author's co-workers. Dr. Julie Siegel and Ms. Patti Traube-Beede, are given special thanks for their support and encouragement. ii

PAGE 3

TABLE OF CONTENTS Page ACKNOWLEDGMENTS ii LIST OF TABLES iv LIST OF FIGURES vi ABSTRACT viii INTRODUCTION AND LITERATURE REVIEW ... 1 MATERIALS AND METHODS 39 RESULTS 53 CONCLUSIONS 127 BIBLIOGRAPHY 145 BIOGRAPHICAL SKETCH 153 iii

PAGE 4

LIST OF TABLES Table Page 1 Phosphoenolpyruvate-dependent phosphotransferase systems in S^. mutans 26 2 Lactose phosphotransferase system and phospho-6galactosidase enzyme activities as a function of buffer composition and pH 59 3 Lactose phosphotransferase system and phospho-3galactosidase activities as a function of ionic strength of buffer reagent 60 4 Determination of optimal amounts of solvents for decryptifi cation of cells for phosphotransferase enzyme assays ... 61 5 Comparison of sugar (glucose and fructose) transport and phosphorylation by decryptified or untreated cells 63 6 Determination of phosphoenol pyruvate dependence for glucose phosphotransferase system-mediated phosphorylation 67 7 Induction of phosphoenolpyruvate-dependent phosphotransferase systems as a function of carbon source in growthmedia (LDH/NADHlinked assay) 71 8 Induction of phosphoenolpyruvate-dependent phosphotransferase systems as a function of carbon source in growthmedia (radioactive assay) 74 9 Competitive inhibition by unlabelled sugars of uptake of radiolabelled sugars by S^. mutans GS5 ,75 10 Glucose phosphotransferase system negative mutants: Group I 88 11 Glucose phosphotransferase system negative mutants: Group II 89 12 Glucose phosphotransferase system negative mutants: Group III 90 13 Glucose phosphorylation by cell fractions of S^. mutans GS5 93 iv

PAGE 5

LIST OF TABLES (Continued) Table Page 14 Phosphoenolpyruvate-dependent phosphorylation of glucose by mutanolysin-prepared membranes of S^. mutans GS5 .... 97 15 Phosphotransferase activities of mutanolysin-prepared membranes 98 16 Inhibitory effect of competing sugars on the phosphorylation of D-[^''^C(U)]-glucose by the phosphoenolpyruvatedependent phosphotransferase system of decryptified cells and mutanolysin-prepared membranes derived from S. mutans GS5 99 17 Glucose-glucose-6-phosphate exchange reaction (transphosphorylation) catalyzed by mutanolysin-purified membranes of S^. mutans GS5 as a function of reactant concentrations. 101 18 Pyruvate-phosphoenol pyruvate exchange reaction as a probe for Enzyme I: Distribution of activity in cell -free extracts of S^. mutans GS5 107 19 The relative glucose and fructose phosphotransferase activities in mutanolysin prepared membranes of mutant and wild-type strains of S^. mutans GS5 108 20 Transphosphorylation as a measure of Enzyme II in mutanolysin-prepared membranes of wild-type and mutant strains of S. mutans GS5 110 21 The distribution of Enzyme I as determined by the phosphoryl exchange reaction between pyruvate and phosphoenolpyruvate in cell -free extracts of wild-type and mutant strains of S^. mutans GS5 112 22 The growth and sugar uptake by cells of S^. mutans GS5 induced for lactose dissimilation in mediin supplemented with lactose and glucose 116 V

PAGE 6

LIST OF FIGURES Figure Page 1 Schematic of the phosphoenolpyruvate-dependent phosphotransferase system 4 2 Proposed scheme for the regulation of carbohydrate transport and metabolism by the phosphoenolpyruvatedependent phosphotransferase system in E. coli 17 3 Lactose phosphotransferase system activity as a function of cell concentration 56 4 The pH optimum for the lactose phosphotransferase system 58 5 Lactose phosphotransferase system enzyme activities and growth of S^. mutans GS5 as a function of time 67 6 Glucose phosphotransferase system activity as a function of phosphoenol pyruvate (PEP) concentration 70 7 Kinetics of gl ucose-6-phosphate formation by the PEPdependent phosphotransferase system of ^. mutans GS5 ... 79 8 Kinetics of mannose-6-phosphate formation by the phosphoenolpyruvate-dependent phosphotransferase system of S^. mutans GS5 81 9 Kinetics of fructose-6-phosphate formation by the phosphoenolpyruvate-dependent phosphotransferase system of S^. mutans GS5 83 10 Growth of ^. mutans GS5 in glucose, fructose, and mannose 85 11 Phosphorylation of glucose by cell-free membranes 95 12 Glucose-glucose-6-phosphate transphosphorylation by cellfree membranes 103 13 Phosphoenolpyruvate-dependent phosphotransferase component EI activity in cell-free membranes 106 14 Diauxic growth patterns by S^. mutans GS5 using glucose and lactose substrates 115 15 Growth patterns displayed by S^. mutans GS5 in fructose plus lactose 119 vi

PAGE 7

LIST OF FIGURES (Continued) Figure P^S^ 16 Growth of S. mutans GS5 (wild-type) and a glucose phosphotransferase negative mutant in lactose plus glucose 122 17 Lactose uptake by S. mutans GS5 (wildtype) and a glucose phosphotransferase negative mutant as a function of cell density ^24 VI i

PAGE 8

Abstract of Dissertation Presented to the Graduate Council of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy STUDIES ON THE PHOSPHOENOLPYRUVATE-DEPENDENT PHOSPHOTRANSFERASE SYSTEMS IN Streptococcus mutans GS5 By Ellen S. Liberman December 1982 Chairman: Dr. Arnold S. Bleiweis Major Department: Microbiology and Cell Science This dissertation is concerned with characterization of the glucose phosphoenol pyruvate (PEP)-dependent phosphotransferase system (glc PTS) in Streptococcus mutans GS5. The PTS is a mode of transport which involv the translocation of a phosphoryl group from PEP to an incoming sugar moiety. It is a complex system which requires up to four proteins, Enzyme I (EI), HPr, Enzyme III (EIII), and Enzyme II (EII), to accomplish the transport and group translocation functions. The objectives of the present study were (1) to obtain a general overview of the glc PTS, (2) to obtain a more detailed picture of the glc PTS by studying isolated membranes derived from cells of S^. mutans and (3) to study the hierarchy of sugar utilization in S^. mutans The substrates of the glc PTS in order of declining affinities were found to be glucose, mannose, and 2-deoxyglucose. The evidence for this finding comes from studies of the competitive effects exerted by the above sugars on the transport of the heterologous sugars, from kinetic studies, and from studies on glc PTS-negative mutants. vi i i

PAGE 9

Isolated membranes derived from cells grown in glucose were prepared using the muralytic enzyme, mutanolysin. These membranes were able to phosphorylate glucose and mannose when supplied with exogenous PEP. They were also able to phosphorylate glucose when the phosphoryl donor glucose-6-phosphate was used (transphosphorylation) thus demonstrating the presence of a functional EII^^^ in cell -free membranes of the wildtype strain. The presence of EI in these membrane preparations was demonstrated by the phosphoryl exchange reaction between PEP and pyruvate. Glucose in the growth-mediun prevented the induction by lactose of the lac PIS in S^. mutans GS5. Thus, glucose is a preferred sugar. Glucose did not appear to repress the induction of the lac PTS in glc PTS-negative cells even though glucose was taken up by these cells. This finding indicates the necessity for a functioning glc PTS for the regulation of lactose uptake. The mutant cells contained wild-type levels of EI but lacked a functioning EII^^^ as shown by the two phosphoryl exchange reactions. These results suggest that the EII^^^ is required for the regulation of sugar uptake in S^. mutans GS5. ix

PAGE 10

INTRODUCTION AND LITERATURE REVIEW Streptococcus mutans is the causative agent of dental caries, the most prevalent bacterial disease in humans. This species is divided into a number of serotypes (a^-5.), however serotype c is most commonly isolated from carious lesions. The growth of this organism is accompanied by the production of lactic acid which causes the demineral ization of dental enamel. The resulting cavity is the clinical manifestation of this disease (13,14). Numerous investigators have been studying the physiology of sugar metabolism of S^, mutans in order to deduce basic mechanisms of transport and dissimilation and to learn about the genetic basis of the pertinent enzymology. It is hoped that such knowledge could be applied to preventative therapy and thus aid in the elimination of this disease. At this stage only limited knowledge exists concerning the transport of sugars into S^. mutans The purpose of this dissertation is to expand our knowledge of a major transport route, the phosphotransferase system, in serotype c^ strain GS5. Mechanisms of transport Carbohydrate transport in bacteria is characterized by three general modes. The first is exemplified by the lactose system in Escherichia coli The energy required for lactose uptake is derived from proton symport, i.e. lactose is co-transported with a proton, thus dissipating the proton gradient produced during electron transport. The second type of transport utilizes adenosine 1

PAGE 11

2 5 '-triphosphate (ATP) as an energy source and is exemplified by maltose uptake in E. coli This type of transport is characterized by the requirement for a periplasmic binding protein. One way these two types of sugar accumulations are differentiated is by the loss of transport functions upon osmotic shock in the case of the latter transport system and the retention of such functions in the former system (7). The third mode of transport is characterized by glucose transport in E. coli and lactose transport in Staphylococcus aureus (21). It is termed the phosphoenol pyruvate (PEP)-dependent phosphotransferase system (PTS). This type depends on the energy inherent in the enol configuration of the phosphoryl group of PEP. Since the phosphoryl group is transferred, it falls into the general category of group translocation. Compone nts of the phosphotransferase system The two systems which have been studied in the most detail, and after which a standard scheme has been modeled, are the glucose system of E^. coli and the lactose system of _S. aureus This model has been developed by the isolation of individual components and their reconstitution in vitro A general scheme for PTS-mediated sugar transport is outlined in Fig. 1. The phosphoryl group is transferred from PEP to a small molecular-weight protein called HPr. This translocation is mediated by the enzyme termed Enzyme I (EI). EI itself forms a phosphoryl ated intermediate. From HPr, the phosphoryl group is transferred to a third protein referred to as Enzyme III (EIII). The final steps involve the concomitant transfer of the phosphoryl group from EIII to the sugar as it crosses the cytoplasmic membrane. The permease is referred to as Enzyme II (EII). Both EIII and EII are sugar specific (31,55,77).

PAGE 12

I o J= r— Q. ta to SO E dJ o C3.-(-> 0) -o s_ I cu sa. I— -o O E C (O Ol o ^ Q. to O O. N c •rLf> -P tn res -—^ §E u +-> oo to >> to to re CD s_ •r>— N t— < C t-4 UJ s (U >— N ^-l SZ O i-H (/I O M • >> SO O c , •1so -a -c C Q. O O ^jz n. to so O) -t•r4i. SO O) re O) to o Q. re to O) r— £ >, sC sre ^

PAGE 14

5 The first two proteins (HPr, EI) are general proteins in that they are involved in the transport of all "PTS sugars" by a given cell (21, 25,78). HPr has been studied in S^. aureus and £. col i In both cases it is a small molecular weight protein; the E. coli HPr is 9600 daltons (1) and the S. aureus HPr is 9000-9200 daltons (77). Its physiological activity is stable to heating at 100 C for several minutes (25,29). From studies of the rates of hydrolysis of phospho-HPr (HPr~P) under acid and alkaline conditions, it has been determined that the phosphoryl group is transferred to the N-1 of the imidizole ring of histidine (1,76). This component has been shown to be non-specific since point mutations in the gene coding for HPr produces pleiotropic effects in terms of sugar transport (55,78). Attempts to cross species lines have met with little success, although the E. col i HPr allows low levels (5% of the homologous system) of phosphorylation of thiomethyl-B-Dgalactopyranoside (TMG) by the S^. aureus PTS enzymes. However, the ^. aureus HPr cannot substitute for E. col i HPr when a-methyl gl ucoside (a-MG) is the substrate (77). Durham and Phibbs (9) were unable to demonstrate cross-complementation between Pseudomonas aeruginosa and E. col i Simoni et al. (77) reported the block occurrred between HPr~P and sugar phosphorylation when S^. aureus HPr was substituted in the E^. coli system. Cords and McKay (6) reported cross-complementation of crude extracts from S^. aureus and ^. lactis ; however, the results are difficult to evluate since a quantitative analysis was not presented. The EI protein has been isolated from S^. Aureus and E_. coli However it has been only partially purified and the failure to attain full purification of this protein has been attributed to its sensitivity to

PAGE 15

6 oxidation (25,30). In S^. aureus the phosphorylated derivative of this enzyme migrates as a single band of 80,000 daltons in sodium dodecyl sulfate-polyacryl amide gel electrophoresis (SDS-PAGE) (25). The molecular weight of this enzyme in E. coli and Salmonella typhi murium has been estimated to be 70,000 and 90,000 daltons, respectively (21). The cytoplasmic localization of EI and HPr was determined by the complete separation of cytoplasmic and membrane fractions (31,77). In S. aureus this was accomplished by ultracentrifiguation (77). The biological activity of HPr and EI has been elucidated mainly in the laboratory of Dr. S. Roseman from studies with E. coli and S. aureus Kundig and Roseman (30) demonstrated that in the presence of ^^Penolpyruvate, and Mg^"*", the rate of the reaction involving the transfer of the phosphoryl group to HPr was directly proportional to EI concentration, whereas HPr demonstrated saturability. This indicated that the catalyst was EI while HPr served as a phosphoryl carrier. The reaction is stoichiometric and dependent on the PEP concentration. Furthermore, determination of equilibrium constants using components isolated in S. aureus demonstrated that the energy inherent in the enolate bond of PEP is essentially maintained in HPr~P (76). During this transfer EI forms a transitory phosphorylated intermediate (21, 25). With high concentrations of PEP and pyruvate, an abortive complex is formed and thus the PTS reaction is inhibited. Saier et al. (64) used this observation to develop a reaction to directly assay for the presence of EI; that is, EI catalyzes a phosphoryl exchange between PEP and pyruvate.

PAGE 16

7 The sugar-specific component, EIII, shows more variation among organisms and also among PTsystems within the same organism. This is best illustrated by three systems: the lactose PTS (lac PTS) of S. aureus and the two glucose phosphotransferase systems (glc PTS) of E. coli The EIII of the lac PTS in S^. aureus was purified from the soluble fraction of an ultracentrifugation of cell extracts (22). Studies showed it to be a trimer composed of three identical sub-units. Various physical techniques such as analytical ultracentrifugation and SDSPAGE allowed a measurement of 35,700 daltons for the trimer (22). This protein is auto-catalytic; i.e., it catalyzes the transfer of the phosphoryl group from HPr to itself (22). Each sub-unit is capable of accepting a phosphoryl group on a histidyl residue. Hydrolysis under acid and alkaline conditions allowed the determination of the phosphoryl bond to be at the N-3 of the imidizole ring (22,25). The phospho-EIII (EIIKP) then is able to donate the phosphoryl group to lactose (76). Experiments with membrane-bound EIII have demonstrated that the phosphoryl ated form of EIII exhibits lipophilic behavior, whereas the non-phosphorylated does not. Sub-unit exchange was readily demonstrated with the phosphoryl ated form but not the underivatized form. Also, dissociation of the phosphoryl ated form by lipophilic agents was observed. It was suggested that physiologically EIII exists as an underivatized, non-phosphorylated trimer and a membrane-bound phosphorylated monomer. It is in the latter form that the EIII has been postulated to be able to intercalate in the membrane and to donate the phosphoryl group to the incoming sugar (25).

PAGE 17

8 As stated, in E. col i there are two different glc PTsystems. One has been characterized as a high-affinity system and is able to phosphorylate the glucose analogue, a-MG, as well as glucose. This system, similar to the lactose system of S^. aureus features a "soluble" EIII. Unlike 1 ac EIII the phosphoryl protein link is via an acyl group (21,25). Its apparent molecular weight is 20,000 (25). As will be described below, EIII^^^ may have a regulatory role (73). In contrast to the latter two systems is the low-affinity glucose system of £. coli This system has been shown to transport glucose and its anomers, mannose and fructose. In addition, it transports the glucose analogue 2-deoxygl ucose (2-DG) as well as being responsible for the accunulation of the acetyl and N-acetyl derivatives of glucosamine and mannosamine (30,55). This EIII is an integral membrane protein which is complexed with the translocating component, EII. It has been isolated by Kundig and Roseman (31) and shown by isoelectric focusing to be composed of three proteins each of which showed EIII activity, albeit each with differeing specificities. One band had EIII activity with mannose, a second was specific for fructose and the third was specific for glucose (31). The soluble components have been isolated and well characterized. Their biological activities have been established through reconstitution. Furthermore, since the phosphoryl addition involves sequential steps, the order as well as the individual proteins involved have been established through the use of a "^^P-probe and the isolation of the intermediates by electrophoresis (30) or gel filtration chromatography (76). The transport step and the concomitant phosphoryl transfer to

PAGE 18

9 the sugar is more poorly understood. Transport requires an intact membrane and conversely membrane proteins require lipid matrices to function; thus dissection of this step is a more formidable task. Various Ell's have been isolated and along with EIII~P have been shown to complete the phosphoryl transfer to the sugar. Among the Ell's that have been isolated thus far is the EII from the low-affinity glucose system in ^. coli (31) and the EII^^^ of S^. aureus (77). q1 c The L coli EIP was isolated using a butanol/urea extraction technique. It was shown to have biological activity in the presence of phosphatidyl glycerol and the requirement for this phospholipid was stringent (31). This observation was surprising since this is a minor lipid within the E. coli membrane (21). lac EII was purified by using a combination of agents including sodiim deoxycholate, Tween 40 and Triton X-100 (77). Lipid-free EII could be obtained by sucrosegradient centrifugation and biological activity was detected in the presence of the other PTS components if a lipophilic environment is created with a detergent such as Triton X-100 (25). Identification of products of phosphotransferase system activity The definitive assignment of functional phosphotransferase systems involves isolating and identifying the derivatized sugar(s) as product (s). Reconstitution of the S^. aureus lac PTS results in the formation of TMG-6-P (77), and lactose-P (78) where the phosphoryl group is esterified at the C6 position of the 3-D-galactopyranosyl moieties. The E. coli glucose systems form gl ucose-6-phosphate (30,55), a direct intermediate in glycolysis. Fructose is phosphoryl ated at CI (55).

PAGE 19

10 other phosphotransferase systems for sugar uptake Many systems for sugar transport have been investigated to date. Most of these systems conform to the model discussed in the preceding section. However, a few have been found to diverge from the accepted scheme. For illustrative purposes two will be discussed in this section. The obligate aerobe, P. aeruginosa contains an unusual PTS for fructose utilization. Only two components have been identified in this system, one membrane bound and one soluble. The soluble protein appears to be similar to EI in terms of molecular weight; however, unlike EI, this may be a peripheral membrane protein since membranes alone have residual PTS activity (9). On the other hand, this residual activity may represent cytoplasmic contamination of the membrane preparation used in the study cited. The present study of S. mutans GS5 will present a similar finding with regard to the glc PTS (see RESULTS ). Another interesting divergence from a better known system is the inducibility of these two components in the pseudomonad. In E. coli, both HPr and EI are constitutive. However, the relative levels are enhanced when glucose is present (53). The membrane components (EII, EIII) vary also with the composition of the mediun. However, the relative levels of the sugar-specific components allow for the conclusion that the E. coli low affinity glucose system is constitutive and the high affinity system is inducible (21). The sugar specific components (EII and EIII) of the S. aureus lac PTS are inducible whereas the equivalent components for glucose transport are constitutive (21). A different pattern is observed in P. aeruginosa. Both factors are inducible (9). This may be related to the presunably secondary role the PTS plays in this organism.

PAGE 20

n The occurrence of the PTS is directly related to the metabolism of the organism. In general, strict aerobes do not have a PTS. There are a number of exceptions such as P^. aeruginosa Also, heterofermentative genera do not transport sugars via a PTS. This is presumably due to the stoichiometric relationship between the fermented sugar and the generation of PEP (21,54). Again £. aeruginosa is an obvious exception to this generalization. Durham and Phibbs (9) have proposed that after being brought in by a PTS, a putative kinase is involved in phosphorylating fructose. Thus, fructose plus PEP results in fructose-l-phosphate which then accepts a second phosphoryl group from ATP to form fructose-1 ,6-di phosphate. Generally, organisms which have phosphotransferase systems are either anaerobes or facultative anaerobes. These organisms utilize the Embden-Myerhof pathway and thus generate two moles of PEP for every mole of sugar fermented (21,54). The advantage of this type of transport lies in the conservation of ATP (54). In Spirochaeta aurantia a facultative anaerobe, only mannitol is transported by a PTS. The product of this reaction is mannitol-1phosphate. Three proteins are required for this reaction. Two are soluble and on the basis of physiochemical properties are analogous to HPr and EI. The third protein is a membrane-bound EII (61). The regulation of this system differs from the model systems. In these systems HPr and EI are regulated on one operon, whereas the sugar speccific components are coord inately regulated as a separate operon. These operons map at distinct loci on the E. coli chromosome (55). In S^. aurantia the three PTS proteins are genetically regulated in a coordinate fashion. In addition, the enzyme responsible for the first

PAGE 21

12 step in mannitol dissimilation, mannitol-1-phosphate dehydrogenase, is also regulated in the operon (55). Phosphotransferase systems and the regulation of transport In addition to a transport function, the phosphotransferase systems (PT-sys terns) in the Gram negative enterics function in the control of non-PTS transport. This control is bilateral: the permeases are regulated directly through allosteric modulation and indirectly by regulating their synthesis (55). In order to demonstrate a model of regulation, the control of the glc PT-systems over the lac permease of ^. coli will be described as an example. A brief review of the lac operon will aid in explaining this model. The controlling factors in transcription of the lac operon are two-fold: a negative modulator, the repressor, and a positive modulator cyclic adenosine 5 '-monophosphate (cAT'lP). In the absence of inducer (lactose or one of its analogues) a repressor molecule inhibits the transcription of lac RNA by binding to an operator site on the DNA. This operator site is flanked by the promoter site on one side and the structural genes on the other. In the presence of an inducer, a repressor-inducer complex is formed. The binding of the inducer causes the repressor to undergo an allosteric shift and therefore causes it to lose its affinity for the sequence of nucleotides to which it binds. It is evident that in the absence of intracellular inducer the lac permease cannot be expressed and thus by controlling the transport of inducer molecules, the cell is able to control the synthesis of transport systems. The prevention of inducer entry is termed "inducer exclusion" (60).

PAGE 22

13 RNA polymerase binding and initiation of transcription is dependent on the formation of an "open complex," that is, the conformation of the DNA is altered as to produce localized melting. The effect of this is to allow the polymerase to transcribe the lac genes (45). The formation of the "open complex" is brought about by a second modulating system composed of two elements: cAMP receptor protein (CRP) plus cAMP. This system is analogous to the repressor-inducer complex except that it exerts a positive control. CRP by itself does not exhibit an affinity to the DNA. However when complexed to the low molecular weight effector molecule, cAf'IP, its affinity towards the DNA increases by virtue of an allosteric transition. The binding site of this complex is on the lac promoter in the region distal from the operator (45). Thus, it is obvious that controlling cAMP synthesis would control the level of expression of the lac permease, and therefore, lactose uptake. Synthesis of cAMP is accomplished by regulation of the allosteric enzyme, adenylate cyclase. It has long been observed that growth of E^. coli in a combination of lactose and glucose results in a biphasic growth pattern. This diauxic growth is a result of the repression of the expression of the lac genes (60). Not only is the inducer (lactose) excluded from these cells but cAf-IP levels are low. It is during the lag period preceding the second burst of growth, following glucose exhaustion, that cAMP is synthesized and sufficient quantities of inducer are accumulated to induce the lac operon. If cells are grown on a permissive growth substrate such as glycerol, where the expression of the lac genes occurs in the absence of inducer

PAGE 23

14 and glucose is added, two effects are observed: (1) there is an immediate cessation of lac gene transcription followed by (2) a resimption of transcription at a repressed level. The immediate severe repression is called transient repression and the second type of repression is termed catabolite repression (38). Both these forms of inhibition occur even though the inducer is present intracellularly. On the other hand, both phenomena reflect the cAMP levels measured. That is, glycerol is a permissive substrate because it allows cAMP to be synthesized, whereas growth in glucose causes an inhibition of adenylate cyclase; thus cAMP levels are low in cells growing on glucose (39,46). The addition of glucose to cells growing on a permissive substrate results in a severe transient decrease in intracellular cAMP followed by a resumption of synthesis of this metabolite but at a lower level than that observed in the absence of glucose (38). The lowering of cAMP levels is due to an inhibition of adenylate cyclase (46), excretion is a function of intracellular levels not of growth substrates (45). Catabolite repression is a misnomer since the effect does not require metabolism of glucose. Non-metabolizable analogues such as a-MG produces the same effect (45,47). The glucose effects (i.e., inducer exclusion, transient repression, and catabolite repression) are exerted with a wide range of metabolizable sugars. Most of these systems have in common the following: they are inducible and they are transported by no n-PTsystems, though their transport may be mediated by an ATPdependent or a proton motive force-dependent mechanism (45,55). A unifying hypothesis to explain the two underlying mechanisms of repression, inducer exclusion and adenylate cyclase inhibition, has been

PAGE 24

15 proposed by Saier (55). This is diagrammed in Fig. 2. The overall mechanism is a phosphorylation-dephosphorylation modulation of competing functions which are all dependent on a central protein. In the outline, the phosphorylation reaction is mediated by the general PTS proteins, EI and HPr. In the presence of glucose, the equilibrium of the phosphoryl donation lies in the direction of glucose phosphorylation; however in the absence of glucose, the phosphorylated HPr is free to donate the phosphoryl group to a hypothetical protein termed RPr (regulatory protein). In the phosphorylated form, this protein becomes a positive effector for adenylate cyclase, thus allowing cAMP to be synthesized thereby negating catabolite and transient repressions. In the nonphosphorylated form (i.e., when glucose is present), RPr has an affinity for the non-PTS permeases. The proposal is that these permeases are allosteric proteins and that RPr is a negative effector. Therefore, RPr binding inhibits transport of non-PTS sugars and is responsible for inducer exclusion. When phosphorylated, RPr loses its affinity for the permease and the unmodified permeases can then function. Much evidence has been accumulated to support this model. A selected amount will be summarized here. Using S. typhimuri um and E. coli it was found that mutations in the genes coding for EI or HPr had a surprising effect on the metabolism of non-PTS sugars such as melibiose and maltose. Tight mutants could not grow on these substrates, whereas the growth pattern of leaky mutants were phenotypical ly indistinguishable from wild-type. However, if a-MG is added to induced cells, the leaky mutants exhibit a profound increase in sensitivity to the repressive effects of this analogue as

PAGE 25

I 4J SO r— CL O ••> to 0) C O) C (/) o) a. Q. Min (/J (U O O +j c x: j:: ra 03 CL a. 5SI -o +-> — >, O QSJ= jC UJ ra O Q.QC7> XJ (/I 3 SO I/) ra x: O Q. l/l Ql. H+-) O r— o c I— 00 O) I— c -o o o c >+•r <0 Sro > x: s_ 4-> >,•!t-H CL+J to O) o -re c > >, OJ OJ N o sc ^ j=: LU O) Q-JD x: 00 <: > o o 4-> 4-> S10 o sa. CD I/) 4C QJ to Si. +-> O) 00 +-> (O E OJ O) C c QJ OJ Q. Q. 0) QJ T3 x: I 1— U -ro o E QJ +-> c ra o r— +J >^ o QJ Q. -o ra so O Q. I— < < u o 1 — 1 to ^ 1— 1 a. C (U o rQJ QJ ID 1 — 1 QJ VI JC in +-) O +-> QJ O • \ CL E iQi o >> >i Q. lO SX3 N ro Q. o C >, 0) E o LU SlO o +J CM I— ij ro Q. O X5 c: 00 CT> ro •rOl C7)(— •r+J +-> QJ OLl. QJ E ro S1 E QJ > E +J O o CO SSc c Q. ro 00 Q. cc ro

PAGE 26

17

PAGE 27

18 compared to wild-type. This repressive effect is evident in terms of growth and enzyme synthesis (66). The degree of this repression is related to the extent of induction. Thus, fully induced cultures are more refractory towards the repressive effects of a-MG. Also, noninduced cells cannot grow if transferred to a medium containing this glucose analogue and the non-PTS sugar, melibiose (60,63). This indicates that the primary effect being investigated in these studies is inducer exclusion (63). Furthermore, the EII specific for the PTS sugar must be present for the sugar to cause this hypersensitivity (63, 66), substantiating the involvement of PTS-mediated transport. A second mutation described by Saier and Roseman (62) mapping at a site co-transducible with the pts operon (genes for EI and HPr) but not part of the operon was found to suppress mutations within the pts operon in terms of the hypersensitivity and total repression described above. The distinguishing phenotypic characteristic resulting from these suppressor mutations was found to be depressed levels of the EI 1 1 for glucose; i.e., the soluble factor which is part of the high affinity glc PTS. These mutants were termed catabolite repression resistant and the gene responsible was termed err. It is postulated that this protein (EIII^^^) may indeed be RPr (60). A second group of mutants were isolated by Saier et al. (67) which also relieved PTS-mediated repression; i.e., repression due to an EI or HPr mutation. The characteristic that distinguished this second group from the _crr mutants, is the specificity of the relief. The err mutation was general in that both transport and metabolism of all the affected non-PTS sugars were relieved of repression, whereas in this second

PAGE 28

19 category, a number of unique mutants were isolated which exhibited a refractory response when a specific non-PTS sugar was tested. That is, there was a double mutant in which maltose transport and dissimilation was released from repression, one which melibiose transport was released, etc. The mutations responsible for this release mapped in or near the genes coding for the individual transport proteins. These results suggest that an altered permease may be responsible for this relief and thus indicate an allosteric interaction between a PTS component and the permeases (60,67). Lastly, increased levels of PEP was shown to partially overcome the inhibitory effect of a-MG-induced repression of glycerol transport in wild-type cells. This indicated phosphorylation involving PEP was utilized by these cells to regulate sugar uptake (58). All these results relate to the transport of non-PTS sugars and thus the regulation of non-PTS-mediated transport. An interaction was also observed between PTS components and adenylate cyclase. However, the effects of the various genetic lesions were not completely analogous. The following observations are relevant to the discussion on adenylate cyclase: (1) Mutations affecting the low affinity EII proteins render the adenylate cyclase of _E. coli insensitive to glucose and its analogues (i.e., 2-DG) but not a-MG, whereas mutations resulting in a disfunctional EII of the high affinity system render the enzyme insensitive to a-MGinduced but not 2-DG-induced inhibition (19). These mutations affect the regulation not the activity of the enzymes and these data indicate a functional PTS is required for adenylate cyclase regulation.

PAGE 29

20 (2) In vitro assays of wild-type adenylate cyclase fail to show glucose-mediated inhibition of this enzyme, again pointing to the need for transport activity (47). (3) EI deletion mutants show low levels of adenylate cyclase activity. Leaky EI mutants are "hypersensitive" to the inhibitory effects of glucose on adenylate cyclase activity and this "hypersensitivity" could be partially overcome in permeabi lized cells by the addition of PEP (48,57). (4) A mutation in the err gene results in depressed levels of cAMP in cells which have wild-type levels of EI and HPr (48). (5) Kinetic analysis revealed that adenylate cyclase was still functioning in vivo albeit at depressed levels in the presence of glucose. This tends to indicate positive regulation is required and taken with other data, glucose prevents the positive regulation of adenylate cyclase. These data also suggest that the err gene product (EI 11^^^) may be involved in regulation. The effect of temperature on the acivity and regulation of adenylate cyclase suggests that a stable complex, i.e., an allosteric interaction, is a prerequisite for activation (48). The isolation of the err mutation along with the observation that pre-growth on glucose enhanced the inhibition of transport suggests that a component of the inducible high-affinity glucose system is also involved in inducer exclusion. In support of this, Scholte et al. (73) have shown the identity of the err gene product to be EIII using crossimmunoel ectrophoreti c techni ques Other phosphotransferase functions There is evidence that the PTS functions in the regulation of motility (55). The responsible factor appears to be the sugar-specific EII protein (36). In addition, the EII

PAGE 30

21 protein may play a role in the fermentation of the sugar they are transporting (36). Lastly, the various Ell's are able to catalyze the transphosphorylation reaction: the transfer of a phosphoryl group from a derivatized sugar to its non-derivatized counterpart (36,59). Ttie physiological significance of this latter reaction is unknown; however, there has been speculation that it may have a regulatory function (60). Characterization of the phosptioenolpyruvate-dependent phosphotransferase systems in the strptococci The literature concerning the streptococci lacks the detail outlined above. As demonstrated in the above discussion, the model system is based on data collected from studies in E. coli and ^. aureus Studies on the streptococci do not involve the molecular dissection of the various PT-systems. Identification of PTS transport in these organisms is based on one or more of the following criteria: stimulation of transport by intracellular PEP reserves and/or PEP-dependent phosphorylation of a sugar, the demonstration that ATP cannot substitute for PEP for either of these two functions, and the identification of the transport product of a sugar and/or its non-metabol izable analogue as a phosphoryl ated derivative. The most extensive studies of the lactic streptotocci have been on the lac PTS in Streptococcus lactis McKay et al. (42) demonstrated that NaF, an inhibitor of enolase which catalyzes the glycolytic step involved in the conversion of 2-phosphoglycerate to phosphoenol pyruvate inhibits the transport of TMG, a lactose analogue. The intracellular product was identified as TMG-P in this and other studies (42,86). Thompson (86) measured PEP utilization along with TMG accumulation and found there was a stoichiometric relationship between these two parameters.

PAGE 31

22 Furthermore, the conditions used in this study did not permit the ATPdependent accumulation of amino acids; thereby demonstrating that PEP was not being converted to ATP by pyruvate kinase. In this organism, lactose metabolic enzymes are plasmid coded (32). In a strain devoid of its lac plasmid, both lac PTS and phospho-3-galactosidase activity are absent, suggesting a close linkage between these lac genes. However, regulation of these enzyme activities does not appear to be coordinated. In wild-type, the phospho-3-galactosidase is present under growth conditions which repress the expression of the lac PTS (32). This is an interesting observation in light of other studies on Gram positive bacteria suggesting that the inducer for the lac PT-systems is galactose-6-phosphate. Morse et al. (43) isolated phospho-6-galactosidase negative mutants from S^. aureus and found that these cells could not transport lactose when grown under conditions which induce the lac PT-system in wild-type. However, galactose-6-phosphate was able to induce this system in such mutants. As stated in a previous section, incoming lactose is phosphorylated at the C5 of the galactose moiety (78). Also, early reports indicate the constitutive nature of the phospho-3-galactosidase of S. aureus (41). In strains of S^. lactis galactose is a more potent inducer of the lac PTS than lactose itself (6,32). Galactose has two transport systems in these cells, one which requires ATP and a second which requires PEP. In an elegant series of experiments, Thompson demonstrated the existence of two systems (88). He accomplished this by comparing galactose dissimilation in the presence of glycolytic inhibitors such as iodoacetate or the presence of ionophores. The latter he termed the Gal P system.

PAGE 32

23 He confirmed his findings by using an ATP-generating substrate, arginine, which allowed galactose transport but not PEP-mediated lactose transport. Through competition studies, it was shown that PEP-dependent galactose transport v/as mediated by the lac PT-system. This is in agreement with LeBlanc et al. (32) who were able to demonstrate the coincident disappearance of the lac and gal PT-systems upon curing a strain of S^. lactis of one of its plasmids. In another study. Cords and McKay (6) isolated a revertant of a lac PTS" strain. However, galactose could not induce the lac-PTS in these cells and upon closer examination, it was shown that these cells only possessed the gal P system. Transport via these two different systems results in the dissimilation of galactose by two different routes. If galactose is translocated by the gal P system, it enters the cell cytosol as a free sugar. Subsequently, it is phosphorylated by galactokinase and ATP (6), and is metabolized via the Leloir pathway (88) to glucose-l-phosphate and consequently by the Embden-Myerhof pathway upon isomerization to glucose-6phosphate (90). Galactose-6-phosphate is the product of PTS transport. This compound is metabolized through the tagatose pathway (43,88). This biochemical sequence involves the conversion of galactose-6-phosphate to tagatose-6-phosphate which is phosphorylated by ATP to tagatose-1 ,6di phosphate and finally to dihydroxyacetone phosphate which can feed into the Embden-Myerhof path (87). It should be noted that in these cells, lactose is phosphorylated at the C6 of the galactose moiety. Thus it also must be metabolized via the tagatose pathway with the free glucose produced by the action of phospho-3-galactosidase being processed through the Embden-Myerhof pathway (87).

PAGE 33

24 There appears to be some controversy as to which galactose transport system is the predominant mode of transport. Based on their biochemical analysis of mutants. Cords and McKay (6) speculated that galactose-6phosphate repressed the gal P system. However, Thompson (88) found that the gal P had a 10-fold lower K for galactose than the lac PTS. In m ^ addition, he found growing cells contained low levels of tagatose phosphate but high levels of fructose phosphate which indicated the Leloir path was operating. He interpreted these results to mean that the Leloir path is the predominant route of galactose degradation in relatively low levels of sugar and thus represents the major route. In addition to these two types of galactose transport systems, a second gal PTS appears to exist (44,88). This system is specific for galactose and thus gal^ lac" cells can only grow on galactose. Other phenotypic traits of this strain are the inability to accumulate TMG and the production of galactose-l-phosphate and galactose-6-phosphate. As already discussed, these are the first intermediates in galactose metabolism when galactose is transported by the gal P and the gal PTS, respectively. Thus, these cells which are lac PTS deficient are still able to transport galactose through a PTS in addition to a proton-driven permease. It is apparent that galactose transport in this organism is complicated. A second PEP-mediated transport system in S. lactis has been defined for glucose. Thompson (86) has shown that there is a stoichiometric relationship between PEP depletion and 2-DG uptake. He also demonstrated that the selective use of inhibitors of glycolysis such as P-chloromecuribenzoate (target: glyceraldehyde-3-phosphate dehydrogenase) leads to

PAGE 34

25 inhibition of 2-DG uptake in metabolizing cells. Based on the following criteria this glc PTS was classified as being similar to the low affinity^, coli system: (1) it was refractory to the inhibitory effects of sulfhydryl reagents, (2) the following glucose analogues 14 appeared to compete with C-glucose: 2-DG, mannose, and N-acetylglucosamine, while the substrate of the high affinity system, a-MG, was without effect, and (3) it is a constitutive system (86). It appears that the predominant transport system for sugars in the cariogenic streptococcus, S. mutans is of the PEP-dependent phosphotransferase type. This is not surprising in view of the fact that this is an aerotolerant anaerobic organism (85), and, therefore, must conserve its energy reserves. The PT-systems studied to date in this organism are outlined in Table 1. The literature contains an extensive survey of the various transport systems found in the species; however, details of the individual systems are lacking. The first system to be defined was the glc PTS. Schachtele and Mayo (71) demonstrated a PEP-dependent phosphorylation reaction involving 2-DG in permeabilized cells of S. mutans ATP could not replace PEP in this reaction. Supporting evidence for a PEP dependent transport system came from the observation that NaF inhibited the uptake of 2-DG. Based on the failure of 6-DG to inhibit phosphorylation of 2-DG, they concluded that this analogue is phosphorylated at C6. These investigators extrapolated their findings to the natural substrate, glucose. Even though it is an obvious assumption that 2-DG is transported and phosphorylated by the same proteins involved in glucose uptake, competitive inhibition studies to prove this point have never been done.

PAGE 35

26 Table 1. Phosphoenolpyruvate-dependent phosphotransferase systems in S^. mutans Glucose (Schachtele and Mayo, 1973) Mannitol (Maryanski and Wittenberger 1975) Sorbitol (Maryanski and Wittenberger, 1975) Lactose (Calmes, 1978) Sucrose (St. Martin and Wittenberger, 1979)

PAGE 36

27 Two other monosaccharides studied in this organism are the hexitols: sorbitol and mannitol. Both of these sugars are substrates of a PEPdependent phosphorylation reaction. These PT-systems are inducible and therefore are evident in cells only when grown on the respective sugars. The enzymes, mannitol-l-phosphate dehydrogenase and sorbitol-6-phosphate dehydrogenase are coinduced indicating a regulon codes for these PTsystems (40). The most thoroughly investigated PTS in this organism is the sucrose (suc)PTS. The reason for this is the medical implication of sucrose metabolism in this species. This organism is able to synthesize a dextran capsule through the action of a glucosyltransferase and it is this capsule that allows adhesion to tooth surfaces (13,14). Slee and Tanzer (80,81) described a unique PTS for sucrose in this species. Using a variety of strains, they demonstrated a PEP-dependent phosphorylation of sucrose to sucrose-6-phosphate which appeared inducible; i.e., it was not present in glucose adapted cells. St. Martin and Wittenberger (83,84) verified the inducibility of the sue PTS by employing a greater variety of growth substrates. Upon translocation of sucrose, the dissacharide is phosphorylated at the C6 position of the glucose moiety (83,84). Hydrolysis of sucrose6-phosphate requires a unique invertase. A sucrose-6-phosphate hydrolase was described in S. mutans (5,84). This enzyme had a high affinity for its substrate and thus could only be detected directly by using sucrose6-phosphate. It is synthesized constituti vely, unlike the sue PTS (84). It is interesting to compare this system to the lactose system in S. aureus. Inthis system galactose-6-phosphate is the inducer (43) and this compound

PAGE 37

28 is the product of a constitutive hydrolase (41). It may be that sucrose6-phosphate hydrolase activity results in the production of the sue PTS inducer. Indeed, sucrose-6-phosphate hydrolase negative cells which have an evident sue PTS are isolated as PTS constitutive (84). However, using the S^. aureus analogy, it seems unlikely that glucose-5-phosphate is the inducer since glucose-grown cells have repressed levels of sue PTS activity (81,83,84). The other product resulting from the action of this hydrolase is fructose. Growth of cells on fructose results in lower levels of the sue PTS than growth on glucose (83). Further experiments must be performed to clarify these contradictory data. Thompson and Chassy (89) have recently described a sue PTS in S^. lactis It is interesting that in this organism they found the sucrose-6phosphate hydrolase to be inducible. They also found a fructokinase which was induced in sucrosegrown cells. The identification of this latter enzyme in cells metabolizing sucrose along with the finding that the cytosol of these cells contains high levels of free fructose, lead these investigators to conclude that the fate of the fructose resulting from the hydrolysis of sucrose-6-phosphate was phosphorylation by an ATP-dependent kinase. Recently a sue PTS has been reported in _E. coli (35,72). This is an interesting finding since it had been previously thought that in this organism the PTS was used exclusively for the transport of monosaccharides (55). The ability to utilize sucrose is a plasmid-borne function, the origin of which appears to be Klebsiella pneumoniae (35). This plasmid bears two genes coding for sucrose metabolic enzymes, EII^"^ and sucrose6-phosphate hydrolase (72). There are some interesting features of this

PAGE 38

29 system. First, there is a coordinate induction of these two proteins and genetic studies suggest that the inducer is fructose or its phosphorylated derivative. This is different from S^. mutans where fructose appears to be a repressive sugar for the sue PTS (84). In addition the genes for these two proteins do not appear to be transcribed as a regulon in S^. mutans Additional evidence that a regulon exists in E. coli is the isolation of a mutant constitutive for both proteins. Another unusual feature is the apparent requirement for the EI 11^^^. This was demonstrated by transducing the sue PTS into EI 1 1 negative strains of 1coli and was confirmed using in vitro reconstitution of the individual protein (35). Using the criteria of PEP dependency, NaF inhibition, and product isolation, Cal-nes (3) concluded that lactose uptake in S^. mutans is mediated by a group translocation mechanism. He used a variety of assays to study this system including the hydrolysis by permeabi lized cells of the lactose analogue, o-nitrophenyl-6-galactopyranoside (ONPG), in the presence of PEP and the transport of TMG into intact cells followed by the extraction and identification of its phosphorylated derivative. A phospho-ggalactosidase has also been identified (16,17) and characterized (4) in this species. Conflicting data as to the regulation of this enzyme have been reported. Results published by Calmes and Brown (4) suggest that this is an inducible enzyme. Furthermore, the preferred inducer is lactose. Galactose is also an inducer but the amount of the enzyme synthesized in the presence of this monosaccharide is less than that obtained upon lactose induction.

PAGE 39

30 These results parallel those obtained by Calmes (3) when investigating the lac PTS; namely, both lactose and galactose are inducers with the dissacharide more active in this regard. These findings are in direct disagreement with the induction pattern of the lac PTS postulated to occur in S^. aureus The results of Hamilton and Lo (17), however, suggest a similarity between the mechanisms of induction of the lac PTS in S. mutans with that in S^. aureus That is, in ^. mutans the levels of phospho-B-galactosidase vary to some extent with the growth condition; however, this enzyme is always present. Also the levels are highest when cells are grown in galactose even though with some strains tested the relative levels were close. The reasons for the discrepancies between the authors may be due to the different strains used in the respective studies. Finally, in a later report published by Hamilton and Lebtag (16), much more dramatic increase in the levels of phospho-6galactosidase are shown in cells grown in lactose and galactose when compared with glucose. For both the lac PTS and phospho-6-galactosidase, galactose is a better inducer than lactose. They interpret their data to mean a co-regulation of the lac PTS and phospho-6galactosidase genes. Enzymes of the tagatose pathway have been demonstrated in S^. mutans (16). The levels of these enzymes increase, though not as dramatically in all strains, when lactose is the carbon source. Furthermore, this increase is evident only when lactose, not galactose, is included in the growth medium. Growth in galactose induces enzymes of the Leloir pathway (16). Thus in this respect, lactose metabolism follows the model defined in S. lactis.

PAGE 40

31 Most of the work on transport in the streptococci has been physiological in nature, The biochemistry of PT-systems in these organisms has not been extensively investigated. One exception is the study of the glc PTS of S^. faecal is Interestingly, the PTS appears to be similar to that of the low affinity of E^. coli in that the EI 1 1 has been characterized to be membrane bound and complexed to EI I (27). Two attempts at dissecting the PTS in S^. mutans are noteworthy. Schachtele (70) isolated membranes from glucose-grown cells of S^. mutans. These membranes alone were able to carry out a PEP-dependent phosphorylation of glucose. Since the method of membrane preparation was crude and yielded impure membranes, definitive conclusions as to the cellular location of the various components could not be drawn. Another study by Maryanski and Wittenberger (40) led to equally unsatisfactory results. They investigated the localization of components of the glc and mannitol (mtl) PT-systems by recombining soluble and particulate extracts of induced and non-induced cells and determining the amount of sugar phosphorylation. In recombining extracts of glucose-grown cells, they were able to demonstrate the phosphorylation of glucose but not the phosphorylation of mannitol Soluble and particulate fraction from mannitolinduced cells recombined to phosphoryl ate both mannitol and glucose (i.e., glucose is constitutive). More importantly, they were able to demonstrate a mannitol reaction when the glucose soluble and mannitol particulate fraction were recombined. Alone neither one catalyzed phosphorylation of mannitol. This indicated that both soluble and particulate components exist in S^. mutans However, in both cases the soluble components alone were as active in phosphoryl ating activities, indicating that the method

PAGE 41

32 of membrane preparation (soni cation) resulted in the formation of small membrane fragnents. Interestingly, in constrast to Schachtele's work, the pellets alonedidnot show activity, supporting the conclusion that extreme fragmentation of membranes occurred during soni cation. This apparent failure to clearly separate the particulate and soluble fraaction make any interpretation difficult. Maryanski and Wittenberger (40) were able to demonstrate the presence of a heat stable component that could enhance phosphorylation of mannitol by cell -free extracts. One minor but perhaps significant point is the 2+ 2+ ability of Ca to replace Mg in their assay system and the inhibition 2+ that In produced. In E. coli these ions had opposite effects (30). 2+ Mg is required for the EI catalyzed transfer (25); therefore, this difference may reflect a subtle diversity between EI components. Regulation of carbohydrate utilization in Gram positive organisms The systems of regulation in the Gram negative enterics have been studied extensively by a number of laboratories. Much of this work has been concerned with Saier's model (55) of regulation. To date very little is known about regulation of transport in Gram positive cells and much less is known about the streptococci. One reason for this lack of knowledge may be the absence of genetic systems of study in these organisms. As with studies of sugar transport, studies of regulation by these organisms have progressed little beyond the descriptive stage. Hamilton and Lo(17) surveyed various oral streptococci for the induction of lactose metabolism. A comparative study of S. salivarius and S. mutans was performed. Their data did suggest that the predominant lactose pathway in S^. salvarius was via 3-galactosidase indicating the presence

PAGE 42

33 of a non-PTS permease, whereas S. mutans contained phospho-3-galactosidase indicating a dependence on the PTS for transport. They based their conclusions on the observation that ONPG could be hydrolyzed by pemieabilized cells of S^. salivarius with equal efficiency in the presence or absence of PEP. However, this result could be due to a large energy reserve within these cells. Interestingly, they observed a true diauxie with S^. salivarius when these cells were grown in equimolar lactose and glucose, where in S^. mutans lac PTS induction under these conditions was observed at low glucose concentrations. In a parallel experiment they demonstrated that isopropyl-B-D-thiogalactopyranoside (IPTG) and galactose could induce lactose metabolic enzymes in glucose-grown cells of both S^. mutans and S. salivarius The obvious contradiction of these two experiments is difficult to reconcile. The latter experiment employed 28 mM glucose as a growth substrate; however, the authors do not mention the amount of glucose remaining at the time of the addition of 8 mM IPTG. The addition of IPTG alone (control) to cells of S^. salivarius produced a greater amount of 3-galactosidase activity than when glucose was present, indicating a repressive effect of glucose; however, the authors made no mention as to the growth substrate in these control cells. The analogous data obtained for S^. mutans were not presented. A diauxie growth pattern occurs when glucose-grown cells are transferrred to a medium containing equimolar glucose and sucrose (82). The second phase of growth correlates with induction of the sue PTS. The mechanism (s) which allows preferential sugar utilization in Gram positive organisms has not been elucidated. The model evoked to explain this form of regulation in Gram negative cells is inadequate. One reason for the nonapplicability of Saier model is the failure to

PAGE 43

34 define a role for cAMP in these organisms. In studies on Bacillus subtil is (8), Lactobacillus plantarum (20,68), Bacillus megateriun (91), and S. faecal is (24), cAMP addition could not relieve repression. Interestingly, in two separate studies, _S. nutans was found to respond to cAMP and adenylate cyclase was detected (17,51). However, in one case the methodology used to detect the enzyme has been shown to give artifactual results (45). It is more difficult to evaluate the second study in which the reliable method of radioimmunoassay was used to detect cAMP since no subsequent reports have appeared in the literature. The second reason it is difficult to apply the Saier model to many Gram positive organisms is that in these bacteria we see one PTS sugar (glucose) regulating other PT-systems (usually for dissacharides) rather than PTS regulation over non-PTS transport. In order to explain the hierarchy of sugar utilization via the various PT-systems, Thompson et al. (90) evoked the concept of catabolite inhibition. This mechanism involves the differential affinities of the various sugar-specific components for HPr. p. For example in the above discussion concerning S. mutans, the sucrose-specific components would have a lower affinity for the HPr~ P than the glucose-specific moieties. Catabolite inhibition could explain the diauxic effect observed in non-induced cells. For instance, in S. faecalis the kinetics of inhibition by glucose of the lac PTS appear to be competitive (23). Alternatively, this concept may be applied to the inhibition observed with pre-induced cells. For example, glucose inhibits the uptake of galactose (PTS-mediated) in galactose-grown cells of S. lactis (90). In the former case, catabolite inhibition would result in a situation analogous to

PAGE 44

35 inducer exclusion (23), whereas a general repression would be observed in the latter case (90). Another theory put forward involved inhibition by sugar-phosphates (60). It is postulated that a sugar phosphate binding site of the EII may be a means of turning off transport. For instance, an increase in intracellular glucose-6-phosphate was shown to be coincident with a decrease in fructose (frc) PTS-mediated transport in L col i Saier and Simoni (65) have shown that lactose uptake in S^. aureus inhibits uptake of other PTS sugar. These depressed rates of uptake are dependent on the lactose-specific PTS components being functional. They suggest that galactose-6-phosphate is responsible since a lag precedes inhibition when the cells are presented with two sugar substrates (i.e., lactose phosphate must be processed to produce the putative inhibition). On the other hand, Thompson et al. (90) rules out inhibition by sugarphosphates as being the cause of glucose inhibition of galactose uptake in pre-induced cells of S. lactis since the levels of glucose-6-phosphate were essentially invariable in cells growing on galactose compared to glucose. The little-understood transphosphorylation reaction has been interpreted by Saier and Moczydlowski (60) to support their proposal of sugar phosphate mediated-regulation; however, in transphosphorylation the binding site on EII is only for a sugar phosphate derivative homologous to the underivatized form. A third type of proposed regulation is relevant here especially when considered with the discussion to follow; this involves the inhibition of a-MG uptake by an energized membrane. The inhibition can be reversed by the use of ionophores and other inhibitiors of the proton motive force (60).

PAGE 45

36 In addition to the regulation observed when cells are presented with two different growth substrates, there is a regulatory mechanism imposed by growth. A secondary transport system for glucose exists in S^. mutans • This is apparently dependent on an energy source other than PEP and evidence acquired by the use of inhibitors such as the ionophore carbonyl-m-chlorophenylhydrazone suggest it is mediated by a proton pump. The affinity of this system for glucose is 8-15 times lower than the PTS (IS). Thus at lower glucose concentrations, the glc PTS would be expected to predominate. This prediction was shown experimentally using the controlled conditions of a chemostat: cells grown under a low dilution rate utilized the PTS (11,18). Surprisingly, glycolytic activity was found to be greater under conditions of glucose starvation than when excess glucose was present. At the higher dilution rate (excess glucose) the PTS was apparently repressed and the low affinity system predominated. The degree of repression was found to be proportional to the growth rate (11). As with the glc PTS, the sue PTS of S^. mutans appears to be inhibited by rapid growth and excess substrate. This has been demonstrated both in batch (82) and chemostat grown culture (10). In the latter study, the investigators calculated that the amount of transport was insufficient to account for the total sucrose uptake observed. Rapidly growing wild-type cells in batch culture had a lower specific activity for the sue PTS than did stationary-phase cells. Mutants were able to grow in sucrose even though they were apparently devoid of sue PTS activity. The authors in both studies interpreted these data to indicate the existence of a secondary transport system. The major problem with interpretation of

PAGE 46

37 the results of the latter study is that the extent of leakiness of these mutants was not adequately quantitated. No activity was detected using an enzymelinked assay which indicated that this mutation was tight; however, growth on sucrose was approximately 2-log below wild-type. PTS activity below the level of detection of the assay could account for the sucrose-supported growth of the mutants. Further studies are needed in order to determine definitively the existence of an alternate system. Relevant to these observations is the inhibition of a-MG uptake (PTS-mediated) in L coli observed during rapid growth. This inhibition may be related to the energized state of the membrane (50). Interestingly, repression of the glc (11) and sue (82) PT-systems in S. mutans is pH dependent; however, activity is increased at the lower pH. If a pH potential imposed by an ATPase is responsible for regulation, activity of the ATPase would result in activation of these PT-systems according to these results since the action of the ATPase cause a decrease in the external pH. Regulation of sugar transport in the streptococci has been largely speculative; documentation of such regulation has been sparse. The major problem is the lack of biochemical information concerning the various PT-systems. It is not feasible to propose a regulatory protein such as the EI 1 1 if it is not known whether the glucose system contains an Elll-type molecule. Until now, none of the individual PTS components has been shown to exist in S. mutans. Furthermore, mutants which totally lack a functional glc PTS have not been obtained for this species. Hamilton and St. Martin (18) isolated a glc PTS-mutant f rom S. mutans which retained 15% of the activity of wild-type. It is difficult to

PAGE 47

38 attribute a regulatory function to the glc PTS using such a leaky mutant. The first step in understanding regulation of sugar transport is a more comprehensive assignment of PTS components. This work attempts to elucidate, in part, the biochemistry of the S. mutans glc PTS and to relate this to the physiological regulations observed in this organism.

PAGE 48

MATERIALS AND METHODS Cultures and cell growth Cells of Streptococcus mutans GS5 were routinely maintained in a tryptone-yeast extract(TYE) broth (17). This medium was composed of 10 g/1 tryptone (Difco Laboratories, Detroit, MI), 5 g/1 yeast extract (Difco Laboratories) and 3 g/1 dibasic potassium phosphate. For maintenance, this mediisn was supplemented with 20 mM glucose (Sigma Chemical Co., St. Louis, MO). For assay purposes, the cells were grown overnight in TYE broth plus 20 mM of a given sugar. These cells were transferred to a defined medium (DM) broth designed for S. mutans (85). To this defined medium, 5 sugar was added. A 10% inoculum size was routinely used. Unless otherwise specified, cells were allowed to grow until late-log/early-stationary phase in the DM; after which time, they were harvested by centrifugation at 12,000 X £ for 10 min and washed once in 100 mM sodium phosphate (PB), pH 7.0, plus 5 mM MgCl2. The centrifuge used for routine purposes was a Sorvall RC-5B (Dupont Instruments, Newtown, CN). Cell preparation for assays In all the assays outlined below, decryptified cells were used. The decryptifi cation conditions were as follows: washed cells were resuspended in PB, pH 7.0, plus 5 mM MgCl^ to onetenth their original volume. A toluene-acetone mixture (1:3 v/v) was used to permeabilize the cells. The mixture was added in a ratio of 100 yl/ml of cell suspension and decryptifi cation was carried out by vigorously shaking on a Vortex Mixer (Scientific Industries, Bohemia, NY) for 1-2 min intervals interspersed with cooling in ice. The total length 39

PAGE 49

40 of shaking was 3-5 min. This methodology was based on that used by Calmes (3) for S^. mutans If a further dilution was required 100 mi^ PB plus 5 mM MgCl2 was added after decryptifi cation to give the desired cell concentration. Determination of the lactose phosphotransferase system and phosphog-galactosidase The assay for the determination of the lac PTS utilized the lactose analogue o-nitrophenyl-B-galactoside (ONPG; Sigma Chemical Co). The standard reaction mixture contained 100 mM PB, pH 7.0, 5 mM MgCl2, 8 mM sodium phosphoenolpyruvate (PEP; Sigma Chemical Co.), 8 mM ONPG, and cells (concentrations to be detailed in legends to Tables and Figures). The total volume was 0.6 ml. This was then incubated for 30 min at 37 C after which time the reaction was stopped by the addition of 1 ml of a 5% Na^COg solution (aqueous). The cells were centrifuged inaSorvall table top centrifuge and the amount of o-nitrophenol (ONP) formed was determined by reading the absorbancy of the supernatant at 420 nm in a Gilford 2600 recording spectrophotometer (Gilford Instruments, Inc., Oberlin, OH). The amount of ONP was then determined from a standard curve and the results are expressed in terms of ymoles of ONP/ug cells (dry weight). For the determination of phospho-6-galactosidase activity, the lactose-phosphate analogue, o-nitrophenyl-B-galactose-6-phosphate (ONPG-6P; Sigma Chemical Co.) was used. Unless otherwise specified, the reaction mixture contained 100 mM PB, pH 7.0, 5 mM MgCl2, 10 mM 0NPG-6-P, plus cells (amounts to be detailed in legends) in a total volume of 0.5 ml. After 30 min at 37C the reaction was terminated with 5% Na2C02, the cells were removed by centrifugation, and the absorbancy of the supernatant was determined at 420 nm. The results were expressed as the ymoles ONP formed/

PAGE 50

41 Vig of cells (dry weight). The assay for the lac PTS was based on that described by Calmes (3) and that for the phospho-6-galactosidase was described by Calmes and Brown (4). Phosphotransferase assay: LDH/NADH-li nked For general surveys, an LDH/NADH-1 inked assay was used (29). A mixture containing 80 mM PB, pH 7.0, 4 TcH MgCl2, 10 mM PEP, .025 mg lactate dehydrogenase (LDH; Sigma Chemical Co.), 3 mM 3-nicotinamide adenine dinucleotide, reduced form, disodium salt (NADH; Sigma Chemical Co.) plus cells (amounts given in legends to Tables and Figures) was monitored at 340 nm in a Gilford 2600 recording spectrophotometer for NADH oxidase. An initial rate was obtained by allowing the reaction to proceed for 4-5 min. The PTS reaction was initiated by the addition of 1 mM sugar substrate. The final volume was 1.0 ml. The reaction was monitored by measuring the decrease in absorbancy at 340 nm for 4-5 min. An initial rate was obtained from this measurement and corrected for the endogeneous NADH oxidase activity. The ymoles of NADH remaining was determined from a standard curve and from this value the amount of NAD^ formed was determined. Results are expressed as ymoles NAD"*" formed /yg eel Is /min. Phosphotransferase assay: radioactive For a more quantitative analysis, a radiolabelled PTS substrate was used. A typical reaction mixture contained 30 mM PB, pH 7.0, 1.5 mM MgCl^, 10 mM PEP, 10 mM sodium flouride (NaF, Sigma Chemical Co.), and 0.1 mM unlabelled sugar to which was added the specific isotope (New England Nuclear, Boston, MA). The amount of each isotope contained in each reaction mixture is given under the legends to the Figures or in the Results section. To standardize the reaction, 90-110 yg of cells was used and the concentration was determined from a standard curve of dry weight vs. absorbancy of cells at 600 nm

PAGE 51

42 (to be described in a following section). For routine purposes, the time of incubation was 30 min; for kinetic studies, the time was 10 min, since the extent of the reaction was linear within this time. Unless otherwise stated, these studies were carried out at 25 C. To assay cell-free membranes the following modifications were made: the buffer concentration was generally increased to 80 mM, PB, pH 7.0, the MgCl2 concentration was increased to 4.0 mf^ and the membrane concentration varied as specified in the RESULTS Incubation was always at 37 C. In subsequent assays, the volume was scaled down to 200 pi in order to conserve material and 4 mM 2-mercaptoethanol was added. These reactions were stopped by diluting 100-200 yl of the reaction mixture 1:9 (v/v) in 1% sugar or when the scaled down assay was used, by adding cold 1% sugar in excess of the reaction volume directly to the assay mixture. In both cases, these mixtures were rapidly cooled. From this, the phosphoryl ated derivative was separated from the labelled substrate by filtration under vacuum through a DE-81 anion exchange filter (Whatman, Clifton, NJ) which had been prewashed with a 1% solution of underivatized sugar. The filters were washed one time with 1% sugar and 4 x with cold H^O. They were then counted in a Beckman LS8000 scintillation counter (Beckman Instruments, Inc., Fullerton, CA). This assay is based on the methodology developed by Simoni et al. (76). The scintillation fluor used was Aquasol (5 ml; New England Nuclear). Transphosphorylation In order to assay directly for the EII^^^, the gl ucose-gl ucose-6-phosphate exchange reaction was employed. The procedure was a modification of that developed by Saier (59). In this procedure, 70 mM PB, pH 6.0, 3.5 mM MgCl^, 3.5 mM 2-mercaptoethanol,

PAGE 52

43 50 mM glucose-6-phosphate (Sigma Chemical Co.), 10 mM NaF and 50 yM D-[^'^C(U)]-gl ucose (4 yCi/;jnole) were reacted with membranes. The total reaction mixture volume was 200 yl. The mixture minus the labelled glucose was prewarmed to 37 C and the reaction was begun by the addition of D-[^^C(U)]-glucose. Incubation continued at this temperature for 30 min after which time the reaction was stopped by the rapid addition of cold H2O and the separation of derivatized product from reactant was accomplished by passing the mixture through a Dowex column, AG1-X8 (CI'), (0.7 x 4 cm; Biorad Laboratories, Richmond, CA). The colimn was washed with approximately 3 column volumes of H^O and glucose-6-phosphate was eluted in 5 ml of 1 M LiCl2 directly into a scintillation vial. To this vial, 10 ml of Aquasol was added and counts were obtained in a Beckman LS8000 liquid scintillation counter. Cpm were converted to dpm by comparison to a standard quench curve (external standard vs. percent efficiency). Results are expressed as ijnoles glucose-6-phosphate formed/yg cells (dry weight). Assay for Enzyme I To detect this enzyme, the technique of Saier at al. (64) was used. The reaction mixture was composed of 40 mM tris(hydroxymethyl)-aminomethane hydrochloride (Tris-HCl), pH 7.5, 8 mM MgCl^, 10 mM NaF, 2 mM sodium pyruvate, 0.2 mM phosphoenol [l-^^C]pyruvic acid, cyclohexyl ammonium salt (10 yCi/ymole) plus cell extract. The total reaction volume was 100 yl. All components except the radioactive substrate were incubated at 37 C and the reaction was begun by the addition of the substrate. Incubation continued for an additional 60 min after which time 0.4 ml Sigma color reagent (Stock no. 505-2; 20 mg/100 ml of 2, 4-dinitrophenyl hydrazine in 1 N HCl ) was added. The tubes were mixed thoroughly and incubated for an additional 10 min

PAGE 53

44 at 37 C. The derivatized radioactive product was separated by the addition of 1 ml ethyl acetate. After vigorous mixing, a 600 yl sample was removed from the organic phase, placed in 5 ml Aquasol,and counted in a Beckman LS8000 liquid scintillation counter. Results are calculated per 600 yl product counted and are expressed as pmoles ^^C-pyuvate formed/yg protein. Growth curve s. Cells were pregrown in DM supplemented with 5 mM sugar. At log-phase, the cells were transferred to fresh medium supplemented with sugar(s) at a concentration of 5 mM/sugar. Growth was monitored with a Klett-Summerson photoelectric colorimeter using a red filter #66 (Klett Mfg. Co., Inc., New York, NY) and/or a Gilford 2600 spectrophotometer (600 nm). Sugar determination in spent medium Lactose was determined by the Boehringer-Mannheim lactose/gal actose kit (Boehringer-Mannheim Biochemicals, Indianapolis, IN). The basic principle of this kit is to convert the lactose to galactose plus glucose by 3-galactosidase. The galactose is oxidized to galactonic acid by galactose dehydrogenase plus NAD"*". The appearance of NADH is detected at 365 nm. A 100 yl aliquot of the spent medium was incubated with NAD plus 1.2 U of 8-galactosidase for 10 min at 25 C. After this incubation, PB, pH 6.8, and H2O was added to bring the volume to 1.34 ml. The solutions were first read at 365 nm to obtain a background and the reaction was then started by the addition of 0.4 U of galactose dehydrogenase. Incubation proceeded for 15 min. (The reaction was determined to be complete at the end of this time period.) The final volume of the reaction mixture was 1.36 ml, the NAD"^ concentration was 0.45 M and the PB concentration was 0.15 M. Controls

PAGE 54

45 lacking 3-galactosidase demonstrated the absence of galactose from the medium as a contaminant or as an excreted end-product. The concentration of lactose was determined by comparison to the reduction of NAD^ by a known quantity of lactose. Fructose was determined by the Boehringer-Mannheim glucose/fructose kit. This kit contains hexokinase plus adenosine 5'-triphosphate (ATP) to convert fructose to fructose-6-phosphate phosphoglucose isomerase to convert fructose-6-phosphate to glucose-6-phosphate, and glucose-6phosphate dehydrogenase plus 3-nicotinamide adenine dinucleotide phosphate (NADP''") to convert gl ucose-6-phosphate to gluconate-6-phosphate plus NADPH (reduced). The results were read at 365 nm and the amount of fructose calculated from a standard. Controls lacking phosphoglucose isomerase demonstrated the lack of glucose in the samples tested. The procedure involved incubating 100 yl of the spent medium with 1.0 M NADP"*", 4.0 M ATP, 1.33 U hexokinase, and 0.67 U gl ucose-6-phosphate dehydrogenase. The reaction mixture was incubated for 15 mi n at 25 C after which time 3.27 U of phosphoglucose isomerase was added to all but the controls and incubation was continued for an additional 15 min. The results were obtained by reading the assay at 365 nm in a Gilford 2600 spectrophotometer. The glucose determination was based on the glucose oxidase method of Raabo and Terkildsen (50). All reagents were from a diagnostic kit (Sigma Chemical Co.). Briefly, glucose is oxidized to gluconic acid plus H2O2. The presence of H2O2 is detected by its reaction with o-dianisidine which, when oxidized, becomes brown and can be detected at 450 nm. Typical samples (0.1 ml) contained up to 15 yg of glucose.

PAGE 55

46 To this 1.0 ml of the reagent was added. This reagent contains 5 U/ml glucose oxidase, 1 Purpurogallin U/ml of peroxidase, and 4 yg/ml of o-dianisidine. After mixing, the reactants were incubated at 37 C in the dark for 30 min. Membrane preparation The procedure used is a modification of the one developed by Siegel et al. (75). Typically, cells were grown in 200 ml of either DM or TYE plus 20 mM glucose. At the log-phase, the cells were harvested and washed 2 x in 0.9% NaCl 2 x in 5 mM ethylenediaminetetraacetic acid(EDTA), and 2 x in 20 mM Tris-HCl, pH 6.8, plus 5 mM 2-mercaptoethanol and 10 mM MgCl2("lysis buffer"). The cells were frozen at some point during the washing procedure. For lysis, cells were suspended in 50 ml lysis buffer with 5 mg purified mutanolysin (Ml; a gift from Kanae Yokagawa, Dainippon Chemical Co., Tokyo, Japan) and incubated 60 min at 37 C; after which time, 2.5 mg RNase and 2.5 mg DNase (Sigma Chemical Co.) were added and incubation continued with stirring for an additional 60 min at 37 C. Since it was detennined that the DNase preparation contained proteolytic activity, phenylmethylsulfonyl fluoride (Sigma Chemical Co.) was included in this last step. Membranes were collected by centri fugation at 30,900 x ^ for 60 min and washed once in 100 mM PB, pH 7.0, containing 5 mM MgCl^ and 5 mM 2-mercaptoethanol. The supernatant from the first and second centri fugation were pooled, dialyzed for 48 h against H^O (with at least 3 changes), lyophylized, and resuspended in 25 ml of 100 mM PB, pH 7.0, containing 5 mM MgCl^ and 5 mM 2-mercaptoethanol. (This will be referred to as the cytoplasmic fraction.) The cytoplasmic fraction was stored at -4 C. The pellet was resuspended in the same buffer and subjected to a low

PAGE 56

47 speed spin at 1075 x ^ for 5 min in order to remove whole cells. The supernatant from this step was then centrifuged at 30,900 x ^ for 60 min to recover the membranes in the pellet. The purified, cell -free membranes were resuspended in 2.5 ml of the above buffer and stored at -4 C, These were free from contamination by cell wall (75). Cell wal 1 -membrane complexes were prepared using the method of Bleiweis et al. (2). For this procedure, cells were grown to earlystationary phase in 200 ml DM supplemented with 20 mM glucose. They were harvested and washed twice with 100 mM PB, pH 7.0, containing 5 mM MgCl2The washed cells were then resuspended in 10 ml of this buffer plus an approximately equal volume of glass beads and 100-200 yl of tri butyl phosphate, and homogenized for 3 min in a Braun tissue homogenizer (Bronwill Scientific, Rochester, NY) cooled by CO^. The beads were removed by filtration through a scintered glass filter (coarse). During this process, the beads were washed with several volumes of buffer. The final suspension volume was 200 ml. DNase, 10 mg, and RNase, 10 mg, were added and enzymatic digestion was allowed to proceed for 2 h at 37 C under conditions of constant stirring. EDTA, 3 mM, was added and the membranes were pelleted out and washed 1 x in 100 mM PB, pH 7.0, containing 3 mM EDTA, and finally resuspended in 100 mM PB, pH 7.0, containing 5 mM MgCl^. This vas then subjected to a 1075 x ^ centrifugation for 5 min and the membranes were collected by centrifuging the supernatant at 30,900 x ^ for 60 min. The "Braun-membranes" were stored at -4C in 2.5 ml 100 mM PB, pH 7.0, containing 5 mM MgCl^. A third method of cell breakage involved homogenizing the cells in a Bead Beater (Biospec Products, Bartlesvi 1 le OK). This apparatus

PAGE 57

48 is analogous to a Waring Blender except that it requires glass beads. One liter of cells grown in DM plus 20 mM glucose was harvested at early-stationary phase. After harvesting and washing twice, they were resuspended in 100 mM PB, pH 7.0, containing 5 mM MgCl2 and 5 mM 2-mercaptoethanol and homogenized using a 200-300 ml volume of glass beads for 5 min. To prevent foaming, tri butyl phosphate was added prior to homogenizing. After filtering through scintered glass (coarse) to remove the glass beads, the suspension was treated with DNase and RNase, 50 yg each, for 2 h at 37 C with constant stirring. The cell wallmembrane complexes were then collected and washed twice in 100 mM PB, pH 7.0, containing 3 mM EDTA; whole cells were removed by centrifuga-. tion and the cell wall-membrane complexes were pelleted and resuspended in 5 ml of 100 mM PB, pH 7.0, containing 5 mM MgCl2. Purification of mutanolysin The purified fraction of mutanolysin. Ml, retained proteolytic activity; therefore, it was necessary to further purify this enzyme. This was accomplished by a two-step procedure (75). The enzyme (15-16 mg) was suspended in H2O using sonication and then centrifuged at 30,900 x £ for 1 h. The supernatant was applied to a carboxymethyl Sephadex C-25 ion exchange colunn (30 x 0.9 cm; Pharmacia Fine Chemicals, Piscataway, NJ) and eluted with a linear gradient of 0.01 M to 0.15 M phosphate buffer, pH 7.0, (600 ml). The column had been prewashed with 0.01 M PB, 0. 15 M PB, and reequilibrated with the starting buffer. Fractions of approximately 5 ml were collected by gravity at a flow rate of 12-15 ml/h. Muralytic activity eluted at 0.045 M to 0.05 M phosphate buffer. Fractions were read at 280 nm in a Gilford 2600 recording spectrophotometer. Those that had an absorbancy

PAGE 58

49 of greater than 0.05 were pooled and concentrated by pressure dialysis. The protein recovery was approximately 50% of starting material. Removal of protease activity was monitored by the Azacoll assay (CalbiochemBehring, La Jolla CA). The pooled, concentrated material, 125 ng, was incubated with the chromogenic substrate, Azacoll, 10 mg, in 50 mM PB, pH 7.0, to a total volume of 2.5 ml. After 2 h at 37 C, the substrate was centrifuged at 12,000 x c[ for 10 min and the supernatant was read against a blank tube which had contained Azacoll but no enzyme. That the purification procedure had successfully removed protease activity was determined by an absence of solubilized chromogen. Mutagenesis and mutant selection A modification of the methanesulfonic acid ethyl ester (B^S; Sigma Chemical Co.) mutagenesis procedure developed by Shanmugam and Valentine (74) was used to mutagenize a culture of S^. mutans Cells were grown in DM plus 20 mM glucose. At early-log phase (Klett=34), 0.1 ml of EMS was added and the incubation of cells continued for 60 min at 37 C. At the end of this time period, the cells were harvested at ambient temperature and washed once with carbon-free DM and sonicated for 30 sec. The volume was brought up to 10 ml with carbon-free medium and the treated cells were incubated at 48 C for 40 min. The cells were pelleted at ambient temperature and resuspended in a small volume of TYE before sonicating for 30 sec. The volume was brought up to 10 ml with TYE supplemented with 20 mM glucosamine and the culture was incubated until. turbidimetric increases, presumed to be growth, resumed. At this time, streptozotocin was added to select for glc PTS" mutants according to the procedure of Lengeler (33). Streptozotocin was added so that the final concentration of this

PAGE 59

50 glucose analogue was 50 yg/ml and incubation at 37 C proceeded for 5 h. The cells were harvested, washed once with carbonfree medium and then allowed to grow overnight in TYE supplemented with 20 mM lactose. The cells were then transferred to fresh TYE supplemented with glucosamine and at early-log phase (Klett = 35) streptozotocin was added to give a final concentration of 50 yg/ml. After 5 h at 37 C, the cells were harvested, washed once with carbonfree medium, and plated on TYE supplemented with 20 mM lactose plus 20 mM 2-DG. After 48 h, the plates were replicated on TYE containing 20 mM glucosamine or 20 mM lactose. Colonies that grew on lactose but not glucosamine were patched onto lactose/2-DG containing TYE agar. These plates were finally replicated onto TYE plates containing either 20 mM glucosamine or 20 mM lactose. Colonies were chosen which grew only on lactose. These colonies were grown in TYE broth plus 20 mM mannitol and the resultant culture was stored in glycerol at -40 C. Dry weight determination Cells were grown in DM plus 50 mM lactose. At various times during their growth cycle, 30 ml portions were removed, harvested, and resuspended in 30 ml, 100 mM PB, pH 7.0, containing 5 mM MgCl^. The absorbancy of this suspension was read at 600 nm in a Gilford 2600 recording spectrophotometer. The cells were collected by centrifugation and resuspended in 3.0 ml of the same buffer, A portion, 0.95 ml, was aliquoted in triplicate to predried (72 h, 60 C) and preweighed aluminum weighing pans. Dry weight (60 C) readings were taken at 24, 48, and 72 h. The readings did not vary and therefore an average value for all three time periods was obtained; and from this average the weight of buffer alone was subtracted to obtain a mean dry weight /absorbancy unit.

PAGE 60

51 Chemical analysis The origin of the sugars routinely used throughout this study was Sigma Chemical Co. for 2-deoxyglucose (2-DG) mannitol, a-methylgl ucopyranoside (a-MG) methyl -3-D-thiogal actopyranoside (TMG), galactose, and lactose and Calbiochem-Behring for fructose and mannose. In order to ascertain the degree of purity of these sugars each commercial product was analyzed by gas-liquid chromatography. The sugars were converted to their alditol acetate derivatives by the method of Griggs et al (15). The derivatives were separated on a glass column (6 ft X 2 mm) packed with 3% SP2330 on Supelcoport 100/120 mesh (Applied Science Lab Inc., State College, PA) on a Tracor model 560 gas chromatograPh (Tracor Instruments Inc., Austin, TX). The initial temperature was 180 C and after an initial hold of 5 min, the temperature was raised to 240 C at 2 C/min intervals. The flow rate of the carrier gas, H^, was 20 ml/min. Detection of the peaks was by a hydrogen flame ionization detector and an Autolab Minigrator electronic digital integrator (SpectraPhysics, Santa Okra, CA) was used to quantitate the peaks. The amounts of glucose, a common contaminant, were as follows: 2-DG, 0.521%; mannose, 0.0626%; mannitol, 0.184%; galactose, < 0.01%, lactose, 0.0015%. The anomers of fructose and a-MG derivatives have the same retention times as glucose, therefore, quantitation was not possible. TMG was not tested. The method of Lowry et al. (37) was used for protein determination. To solubilize membranes, 3"; sodium dodecyl sulfate (SDS) was added. Proteins added during the purification of membranes (e.g., RNase, DNase, and mutanolysin) were not subtracted from the final data determinations.

PAGE 61

52 Radioisotopes The following radioisotopes were purchased from New England Nuclear: D-[^^C{U)]-glucose (4.0 mCi/mmole); D-[^^C(U)]fructose (359 mCi /mmole) ; methyl (a-D-[^^C(U)]-gluco)pyranoside (275 mCi/mmole); D-[l ,2-^H]-2-deoxyglucose (37.3 Ci/mmole); D-[1--^H(N)]mannitol (17 Ci/mmole); methyl (6-D-[methyl-^^C]thiogalacto)pyranoside (54.7 mCi/mmole); D-[l-^^C]-mannose (48.6 mCi /mmole); [^^C(U)]-lactose (0.97 mCi/mmole); D-[^^C(U)]-2-deoxyglucose (282 mCi/mmole). Phosphoenol-[l-^'*C]-pyruvic acid (10.6 mCi/mmole) was purchased from Amersham (Arlington Hgts. IL).

PAGE 62

RESULTS General conditions for assay of phosphotransferase systems The initial objectives of this study were concerned with learning about the regulation of lactose uptake in S. mutans GS5. Therefore, many of the basic parameters for the study of the phosphotransferase systems (PT-systems) in this strain were established using the lac-PTS as a model. For these studies, two assays were employed: (1) ONPG plus PEP and 0NPG-6-P were used as substrates for the PTS and for phospho-Bgalactosidase, respectively, and (2) the generation of pyruvate from the donation of the phosphoryl group of PEP to lactose was measured by the oxidation of NADH in the presence of lactate dehydrogenase (LDH). Since there is a stoichiometric relationship between lactose and PEP and between PEP and NADH, the amount of NADH oxidized is a measure of the amount of lactose phosphorylated. Fig. 3 demonstrates the linearity of the two PTS assays. Up to 625 yg dry weight of cells may be used to obtain a linear relationship in the utilization of ONPG plus PEP under the conditions outlined in Methods On the other hand, the oxidation of NADH is linear up to 325 pg of cells, after which a plateau is reached. The measurement of NADH oxidation is approximately 70-fold more sensitive than the phosphorylation and subsequent cleavage of ONPG as determined by comparison of the results obtained by the two different assays using the same dry weight measurement of cells. This may reflect the relative affinities of the lac PTS for ONPG vs. lactose. ONPG, an analogue of lactose, would be expected to have a lesser affinity than the natural substrate. 53

PAGE 63

4-> Q. cu >> 1 O -a X cu E O) "O O cu E to •rto to •ro 4-> -O fO o U o cu x: c 3:: +-> cu Q cu cu S<: s: 2: ro QE •r10 o Q _i -o CU >, JQ c •r™ o •4J si > so •r— o to >i to +J to c cu O cu cu •o lO -o Q.-0 to cu to E to =! 03 0) (0 to (0 4-> s-o cu CU >, > < q; S s_ rc3 o cu O ph cu • to to 1. o +J >> Q. cu X E rC c_> o to O) "i •rto to +J 03 o O -> 03 O u o o CU •r— n3 +-) Si_l +-> cu M o a. (U 1(0 LU E +-) 1 o CO c o M O cu cu CU cu SI sz CJ 4-> +-> to

PAGE 64

55

PAGE 65

56 Fig. 4 demonstrates that the pH optimum of the lac PTS is 7.0 using either of the two reaction systems described above. This optimum is dependent on the buffer system used. Table 2 shows the results of assaying for the lac PTS using the two different methods and phospho3-galactosidase in the presence of various buffers. Phosphate buffer at pH 7.0 is optimal for both PTS and phospho-g-gal actosidase activities. Lac PTS activity appears to be independent of ionic strength between 10 and 500 mM sodiim phosphate. This can be seen in Table 3. Also, phospho-6-gal actosidase activity is most pronounced at ionic strengths between 50 and 1000 mM. On the other hand, NAD^ production is independent of all ionic strengths employed. Assaying whole cells required the permeabilization of membranes to phosphorylated compounds such as PEP using a toluene-acetone mixture. The rationale for using "decryptified" cells was to allow for controlled intracellular concentrations of reactants. Since this type of treatment may cause alterations of the membrane proteins, it was necessary to standardize the conditions of this procedure. Decryptifi cation involves the addition of the toluene-acetone mixture followed by vigorous shaking of the cell suspensions. Table 4 outlines a variety of parameters followed during this step. Two proportions of toluene with acetone were examined: 1:3 and 1:8 (v/v). The amount of this mixture per cell volime was considered as well as the extent of mixing using a Vortex Mixer. All shaking was done at 2-min intervals with approximately 1-min cooling in ice. A toluene-acetone mixture of 1:3 (v/v) at a final concentration of 100 yl/ml cell culture allowed for the highest lac PTS activity. The

PAGE 66

Fig. 4. The pH optimum for the lactose phosphotransferase system. Cells were grown and decryptified as outlined in Table 2. A cell concentration of 120 yg (dry weight) was used to measure ONP released by the ONPG + PEP reaction { • ) and NAD+ generated by the LDH/NADH1 inked spectrophotometric assay ( o ) at indicated pH levels in 60 mM PB (ONP assay) and 70 mM PB (spectrophotometric assay).

PAGE 68

59 Table 2. Lactose phosphotransferase system and phospho-3-galactosidase enzyme activities as a function of buffer composition and pH. Buffer pH nmoles ONP/yg cells'^ ONPG + PEP 0NPG-6-P + b umoles NAD /min/yg cells 1 I to 3. U .09 1.38 U 6.0 .43 2.10 27.9 6.5 .71 4.44 47.9 6.5 .66 4.64 45.0 7.0 .86 5.25 75.7 7.5 .81 4.48 67.9 MOPS^ 7.0 .78 4.23 62.1 TRIS-HCr 7.5 .72 1.80 39.3 8.0 .59 1.46 45.0 9.0 .44 1.11 8.5 10.0 .21 0.18 0 MES: 2(N-morpholino)ethanesulfonic acid, sodium salt; PB: sodium phosphate; MOPS: 3-(N-morpholino)propanesulfonic acid, sodium salt; Tris-HCl: tris(hydroxymethyl )-aminomethane hydrochloride. Cells were grown to late-log/early-stationary phase in defined medium containing 5.0 mM lactose, harvested, washed 1 x with 100 mM PB, pH 7.0, containing 5 mM MgCl2 and then resuspended in 100 mM PB, pH 7.0, containing 5 mM MgCl^ to a final volume 1/20^^ of the original culture. The cells were decryptified with a toluene-acetone mixture at a final ratio of 50 yl/ml of cells for 5 min. All reactants were made up in H2O. The final concentrations of the above buffers were 70 mM for the LDH/NADHlinked assay, 60 mM for the ONPG + PEP assay, and 80 mM for the 0NPG-6-P assay. Since cells were suspended in 100 mM PB, pH 7.0, the contribution of this buffer was minimized by using a 2-fold suspension of cells and one-half the routine volume of cell-suspension. This results in a 2.5 mM concentration of PB in the LDH/NADH assay, 3.8 mM in the ONPG + PEP assay, and 1.8 mM in the 0NPG-6-P assay. The concentration of cells was 140 pg, dry weight, for the LDH/NADHlinked and ONPG assays and 56 yg, dry weight, for the 0NPG-6-P assay. MgCl2 concentrations were 3.5 mM for the NADH/ LDH-1 inked assay, 3.0 mM for the ONPG + PEP assay, and 4.0 mM for the 0NPG-6-P assay. See Methods for details of each enzymatic assay procedure.

PAGE 69

60 Table 3. Lactose phosphotransferase system and phospho-3-galactosidase activities as a function of ionic strength of buffer reagent. nmoles ONP/yq cells^ ONPG + PEP 0NPG-6-P ymoles NAD''"/min/yg cells^ 1000 .091 4.18 20.9 500 .175 3.82 20.5 100 .138 6.90 22.2 50 .166 5.72 24.0 10 .168 1.70 14.3 ^PB, pH 7.0. The procedures used are as outlined in Table 2.

PAGE 70

61 Table 4. Determination of optimal amounts of solvents for decryptification of cells for phosphotransferase enzyme assays.^ Toluene-acetone (v/v) 1:3 1:3 1:3 1:3 1:3 1:3 1:8 yl/ml 100 100 100 100 50 25 100 Vortex time (min) 0 2 4 5 5 5 5 ONPG + PEP^ .534 .981 .913 .816 .515 .515 .476 ONPG-6-P*^ 21.7 25.7 26.9 26.9 35.0 28.1 36.2 Cells of S^. mutans GS5 were cultured overnight in defined mediun containing 5.0 mM lactose, harvested, and washed as described in Methods ''cells, 103 yg (dry weight), were incubated with 100 mM PB, pH 7.0, 5 mM MgCl2, 8 mM PEP, and 8 mM ONPG (total volime: 600 yl) for 30 min at 37 C. The reaction was stopped with cold 5% NaCO^ and the cells were removed by centrifugation. The amount of ONP formed was determined by spectrophotometric measurements of the supernatants at 420 nm. Results are expressed as nmoles ONP/yg cell (dry weight). Cells, 21 yg (dry weight), were incubated with 100 mM PB, pH 7.0, 5 mM MgCl^, and 100 mM 0NPG-6-P (total volume: 500 yl) for 30 min at 37 C. The reaction was determined as above and is expressed as nmoles/yg cells (dry weight).

PAGE 71

•62 permeabilization appeared to be complete after 2-min of mixing. If phospho-3-ga1actosidase activity is measured a somewhat different profile is obtained; optimization occurred at a 50 yl/ml cell culture of a 1:3 (v/v) mixture or a 100 yl/ml cell culture of a 1:8 (v/v) mixture. Since the emphasis of the present study is on the various PT-systems, the optimum conditions were chosen on this basis. Table 5 compares sugar transport into intact cells vs. phosphorylation of that sugar by decryptified cells. As can be seen with both fructose and glucose, transport results in a 2-fold greater amount of sugar being converted to its phosphorylated derivative compared to phosphorylation. This could be due to one of several factors. First, treatment of the membrane with toluene-acetone may result in an inactivation and/or rearrangement of the transport proteins thereby preventing maximal uptake of sugar for phosphorylation. Second, two glucose transport systems have been shown to be operative in this organism (18). Part or all of this higher transport activity may be due to a contribution of this second system. Since this latter system requires a proton motive force, it can be observed with intact cells only. Also, the presence of an ATP-dependent kinase (see Table 6) indicates an alternate fructose system; therefore, the above discussion concerning glucose transport may pertain to fructose transport. Lastly, this assay was performed with a radiolabelled sugar substrate which is converted to a labelled phosphorylated derivative. The product is separated from the reactant by an anion exchange filter which will trap the negativelycharged sugar phosphate or, alternatively if intact cells are used, the negatively charged cells. These filters may be more efficient for

PAGE 72

63 Table 5. Comparison of sugar (glucose and fructose) transport and phosphorylation by decryptified or untreated cells.^ Substrate Glucose Fructose (nmoles/ml ) Sugar transported'^ 2.4 1.5 Sugar phosphorylated 1.2 0.7 Cells were grown in fructose according to the procedure outlined in Methods At early-stationary phase, the cells were harvested and washed 1 x and finally resuspended to 4 x their original concentration. One-half of this suspension was decryptified with a toluene-acetone mixture as described previously and the other half was untreated. For transport assays, untreated cells (115 ug) were incubated with 40 mM PB, pH 7.0, 2 mM MgCl^, 10 mM NaF, and either 100 yM D-glucose plus 125 v!A D-[^^C(U)]-glucose (4 yCi/ymole) or 100 \M D-fructose plus 1 yM D-[^\(U)]-fructose (359 yCi/ymole). ^Decryptified cells (113 yg) were used to test for sugar phosphorylation. The conditions were altered from the transport assay in the following manner: 10 mM PEP was added and incubation was in 50 mM PB, pH 7.0, plus 2.5 mM MgCl^The same amounts of radiolabelled substrates were used. In both types of reactions, incubation was carried out at 37 C for 10 min after which time 0.2 ml was removed and diluted into 2 ml of 1% cold sugar. These were then passed through a DE-81 filter and the filters counted in a liquid scintillation counter. Raw counts were then converted to dpm and nmoles/ml were calculated from known specific activities.

PAGE 73

64 binding the cells than the product, thus, the higher apparent transport activity may be partly due to an inherent characteristic of the assay. Even though sugar transport appeared to lend itself to a more sensi tive assay, I chose to study sugar phosphorylation using decryptified cells. The reasons for this were as follows: (1) the concentrations of the reactants could be controlled more carefully, and (2) the use of decryptified cells eliminated any secondary transport systems (e.g., via the proton motive force) that may be present. Fig. 5 profiles the lac PTS and phospho-6-gal actosidase activities during the growth curve described by strain GS5 when grown on this sugar. It is evident that the lac PTS reaches maximum activity at earlystationary phase. This was confirmed by the LDH/NADH assay (data not shown). The catabolic enzyme, phospho3gal actosidase, however, reaches maximal levels of activity at early-log phase. This maximization of phospho-B-galactosidase appears to coincide with the early detection of the lac PTS. In all subsequent experiments involving the several PT-systems, late-log or early-stationary cells were used. Phosphorylation of sugars by the various PT-systems (e.g., glc PTS) is dependent on PEP by definition; ATP will not substitute for PEP. This is clearly seen in Table 6 in the case of glucose. However, ATP was able to donate its phosphoryl group to fructose and this reaction pro vided 40% the amount of fructose-6-phosphate formed as when PEP was the donoi". Mannose was also able to accept a phosphoryl group, but only 8% of product formed as compared to assays using PEP as a phosphoryl source. This organism has been shown to contain a fructokinase which is able to catalyze mannose phosphorylation (5). Such an enzyme

PAGE 74

m c 0) 00| o (/I (O > o. sz: O) o 4-> •Ic "O l/l OJ ra C Q} •II— Q. 4(U I CO. I o Q. > sC +J (0 i/> g LU D(/) •r-a + (/) +J >•.li +-> OJ m x: 1 — C 0) 0) +J Q) -o x: O) 4-> to C > 2 to o sO) O) to iJO) O) s to C to to r— S_ r— 4J O) o o x: Q. to • O O) -£= E Q.-I-t- 0) to 4O O +J O C to O —I -r+-> O • c ID 3 o y 0)4c •I-a to 0)' 3 •!^t' O) 4-> SQ.^ = ?^ lO Sc to u : 0) o c E -o to to 0) r— •1— r— +-) O) •1O o •I-o JO C sto +-> T3 O) o c to 4O o> to "O to o +-> to Io 53 O) oCL-rLf) o to to o> c to to 0) O) >> >> to 4- to •!to > lO •!(-> a. o I to I (U O w) Q. to Z "O O -rto OJ o -C 4-) +-> o to "O I— C to rO O)

PAGE 75

66 ONP (nmoles) RELEASED/ug CELLS FROM 0NPG-6-P (a) ONP(nmoles) RELEASED/jug CELLS x |0"' FROM ONPG+PEP(o) g g: CVj O 00 CD ^ CM I I 1 1 1 1 1 1 1 1 (•) SlINPl 113n>1

PAGE 76

67 Table 6. Determination of phosphoenolpyruvate dependency for phosphotransferase system-mediated phosphorylation. Substrates for phosphorylation Glucose Fructose Mannose ATP 0^ 0.5^ 0.06^ PEP 2.9^ 1.2^ 0.7?'' Cells grown in defined medium plus 5 mM glucose were harvested, washed, and decryptified. The permeabilized cells, 153 yg (dry weight) were incubated with 70 mf^ PB, pH 7.0, 3.5 mM MgCl2, 10 mM ATP or PEP, 10 mM NaF and 100 yf'l D-glucose plus 125 yM D-[^\(U)]-glucose (4 yCi/ Unole) or 100 yM D-fructose plus 1 yM D-[^^C(U)]-fructose (359 yCi/ymole). The final reaction volime was 1.0 ml. '^Cells grown in defined medium plus 5 mM glucose were harvested, washed, and decryptified. The permeabilized cells, 109 yg (dry weight) were incubated with 70 mM PB, pH 7.0, 3.5 mM MgCl2, 10 mM ATP or PEP, 10 mM NaF, and 100 yM D-mannose plus 10 \M D-Ll-l^cj-mannose (48.6 yCi/ ymole). The final reaction volume was 1.0 ml. In both a and b, the reaction was allowed to proceed for 10 min at 25 C. Reactions were halted when 0.1 ml was removed and diluted in 1% homologous sugar, filtered through a DE-81 filter and counted in a liquid scintillation counter. Results are expressed as nmoles sugar phosphorylated/ml.

PAGE 77

68 is likely responsible for the small amount of phosphorylation seen here. The fructokinase is present in glucose-, mannoseand fructose-grown cells (data not shown). In all experiments performed in this study PEP was used, however, the possibility exists that some PEP was converted to ATP via pyruvate kinase. All results using fructose as a substrate are interpreted as being at least 60% dependent on the PTS. Fig. 6 demonstrates that at the concentration used in this study, 10 mM, PEP is well beyond the limiting range. Resting cells of S. lactis have been shown to contain an energy reserve in the form of 3-phosphoglycerate (86). Presumably, S. mutans cells in early-stationary phase contain a similar reserve since transport in intact cells is observed (Table 5). If NaF is excluded, glucose phosphorylation occurs in decryptified cells without the addition of PEP (data not shown). Reproducibility of this result was not always obtained, an observation confirmed by others (Dr. G. Jacobson, personal communication). Therefore, for purposes of standardization, NaF was always included and the concentration used, 10 mM, gave maximum inhibition of glucose phosphorylation in the absence of PEP (data not shown). Comparative study of phosphotransferase-mediated s.ug:3.r phosphorylation. In order to obtain a profile of the various PT-systems in this organism, a survey was set up in which the cells were grown in various growth substrates and assayed for the presence of specific PT-systems. The results are outlined in Table 7. The glc PTS and man PTS can be detected under all the growth conditions tested leading to the conclusion that these two sugars are phosphorylated via a constitutive system. Phosphorylation of 2-deoxyglucose (2-DG) was detected in all cells

PAGE 78

o T3 • (U c LO o a> CM re •1— 3. o. a 0) C +-> O) tn > • c CO +-> (U CO lO •p OJ o o 03 4-> o •r— x: •rm to "O 4-> 4-> o x: 03 o CO E 0) c c CD O O) 1 Q. >> +-> o Q s_ r~ CO c CO +J >> ro o 31 CO C 4-> o 3 4c CL (0 c SZ (/) (U >> > (0 ra E o -> n3 c: o o CO So 0) +-> (U 3 Sro 4-> u a 1 o cu O s o o 4-> E ^ o E s1 — Q. Q. Qc a 3. Q. f — TD (/) LU ro OJ \ fO a. +j O SZ 0) (0 o -(-> Cl CO 3. CM co 0) O 3 1 -o (D 4-> U ^ O LlJ 1/1 CD Q 4-> O > r— s: O cu C i. O) CO E o > >i s: SO c Q. E (U U CO o 3 CO 1 — o O LD 1 — EI rt C7) r— O) T3 4J CO T3 1 1 c p— ro 1 o 0) O £ o u '<*^ B

PAGE 79

I I. ^ q u3 cj CO csi cvi — — (Hu/S8|ouju) 31VHdS0Hd-9-3S03nn9

PAGE 80

71 Table 7. Induction of phosphoenolpyruvate-dependent phosphotransferase systems as a function of carbon source in growth media (LDH/NADHlinked assay). Q 1 1 hiC 4" V* a ^ a 4" e 4/-I ^ oUDstrate testea Gl ucose Growth Mannose carbon source^ Mannitol Lactose Galactos Glucose 6.3 8.1 12.7 1.5 18.3 Mannose 4.5 4.9 4.8 1.9 15.7 Mannitol 0 0 3.0 0 0 2-Deoxyglucose 0.9 0.9 2.0 0 9.4 a-Methylglucoside 0 0 0 0 0 Galactose 0 0 0 0 0 Isopropyl-6-Dthiogalactopyranoside 0 0 0 1.4 3.9 Lactose 0 0 0 7.5 20.7 Glucosamine 5.5 N.T.^ N.T. N.T. N.T. Cells were grown in the several sugars as described under Methods After harvesting and washing, they were resuspended in buffer to 1/20*^ of their original volumes. The cell concentration varied between 60-325 yg/ml. To insure linearity, at least two cell concentrations were used to test a given sugar for sugar phosphorylation. Where activity could not be detected the upper limits of this range (240-325 ug) were reported. '^The LDH/NADHlinked spectrophotometric assay was used to assay for sugar phosphorylation as described in Methods All data are expressed as ymoles NADH converted to NADVyg cells/min x 10"^. %t tested.

PAGE 81

72 except those grown in lactose. This compound is a glucose analogue and thus would be expected to be transported via the glc PTS as is the case in ^. coli (21). Schachtdle and Mayo (71) showed that the uptake and phosphorylation of this analogue in S^. mutans is through a PTS, most likely the glc PTS. The detection of PEP-dependent phosphorylation of this sugar analogue in all but lactose-grown cells argues for such a PTS being constitutive. The most likely reason for its absence in lactose-grown cells is that low levels of the glc PTS preclude the detection of 2-DG phosphorylating activity given the sensitivity of the assay used. In agreement with Schachtele and Mayo (71), manni to 1grown cells show a high level of 2-DG activity; again, most likely because glc PTS activity is high in these cells. One means of testing for the presence of two glc PT-systems is to compare the phosphorylation of the two glucose analogues, 2-DG and a-MG (21). It can be seen (Table 7) that a-MG is not phosphorylated thereby leading to the tentative conclusion that there is only one glucose system operating in this organism and it would be analogous to the low affinity system in _E. coli Mannitol, as has been shown by others (40), is phosphorylated by an inducible system. Interestingly, galactose is not an apparent PTS substrate even though growth in this sugar leads to a high level of other PT-systems. This may be related to the poor growth of the cells in this carbon source. Lactose and its analogue, isopropyl3-D-thiogalactopyranoside (IPT6) appeared to be phosphorylated by an inducible system. This is in agreement with a report published by Hamilton and Lo(17). Galactose as well as lactose appear to be inducers; in S. aureus galactose-6-phosphate has been shown to be the inducer for the lac PTS (43).

PAGE 82

73 The results obtained in Table 8 reaffinn the general pattern discussed above. Here the cells were grown as in Table 7 but the PTS assay was conducted using radiolabel led substrate at a saturating level. Glucose, again, appears to be constitutive. The relative levels are similar to those found using the LDH/NADH assay. Glc PTS is low in lactose-grown cells and high in mannitol -grown cells. However, there appears to be a contradiction with mannose-grown cells in that PEPdependent glucose phosphorylating activity is lower than anticipated. In agreement with the previous results, 2-DG is phosphorylated while a-MG is not. f'lannose phosphorylation does not vary greatly between mannoseand glucose-grown cells but shows a significant decrease in lactose-grown cells. Fructose, like glucose and mannose, appears to be phosphorylated by a constitutive system. The constitutive nature of PEP-dependent phosphorylation of glucose, mannose, and fructose suggests a possible physiological relationship amongst the PT-systems for these three sugars. As discussed in the Introduction the low affinity glc PTS in E^. coli phosphorylates glucose, mannose, fructose, and glucosamine (31). In addition, this system phosphorylates 2-D6 but not a-MG. Since it appeared that an analogous system existed in this strain of S^. mutans, a series of competition experiments was done to test for this possibility. In these experiments, 14 a C-substrate competed for phosphorylation against an excess of unlabel led sugar. The results are presented in Table 9. Glucose and mannosewere mutual ly competitive. Mannose was a less efficient competitor for glucose phosphorylation than was glucose. Glucosamine was also a competitor for both mannose and glucose but at a lower degree of

PAGE 83

74 Table 8. Induction of phosphoenolpyruvate-dependent phosphotransferase systems as a function of carbon source in growth media (radioactive assay). Growth carbon source ^ Substrate tested'' Glucose Mannose Mannitol Lactose (pmoles sugar-phosphate/yg cell dry weight) Gl ucose 28.7^ 18.5 95.8 38.6 Mannose 44.0 30.5 N.T. 6.4 2-Deoxygl ucose 0.9 N.T. N.T. N.T. a-Methylglucoside 0 N.T. N.T. N.T. Methyl 3-Dthiogalactopyranoside N.T.^ N.T. N.T. 3.5 Mannitol N.T. 0 12.3 N.T. Fructose 56.1 20,0 49.3 4.8 Cells were grown as described previously (Table 5, Methods ). ''cells were decryptified and assayed with various sugars. The final concentrations of reactants were: 30 mM PB, pH 7.0, 1.5 mM MgCl2, 10 mM PEP, and 10 mM NaF. The substrate concentrations included 100 yM of unlabel led sugar containing the following concentrations of the homologous radioactive substrates: 50 yM D-[^^C(U)]-gl ucose (4 yCi/ymole), 3 yM D-[l-^^C]-mannose (48.6 yCi/ymole), 0.03 yM D-[l ,2-^H]-2-deoxyglucose (37.3 mCi/ymole), 0.08 yM methyl-a-D-[^\(U)]-gl ucoside (275 yCi/jjnole), 19 yM [methyl^C]-B-D-thiogal actopyranoside (54.7 yCi/ymole), 0.006 yM D-[lH(N)]-mannitol (17 mCi/ymole) or 2 yM D-[^^C(U)]-fructose (359 yCi /ymole) Reaction mixtures included a range of 12-350 yg cells and specific activities were calculated from an average of those values falling within the linear portion of the curves generated (not shown). The standard deviation for 5 identical samples was 3.8. ^Not tested.

PAGE 84

75 z: I a i CJ O ir> i~ u o t/t •*-* o di rO I/I i_ <_> fO (U L. 5" I/I 3 4-> c 4-1 C o X o § E Of -C o c VI CVi at 4-* fi u c o" o 3 I/I u u c u. 1.
PAGE 85

— tn — 76 efficiency. Glucose phosphorylation appeared to be inhibited by a-MG but since direct phosphorylation of this analogue could not be demonstrated, it was concluded that this observed inhibition was the result of a non-specific mechanism. As expected, fructose competed with its 14 C-isotope; however, tt did not inhibit glucose phosphorylation. There was a mutual competition between mannose and fructose. This may be due to some recognition of mannose by a frc-PTS or the sharing of a distinct "mannose site" on the glc PTS with fructose. If the latter explanation is correct, affinity for fructose would be extremely low since this sugar does not interfere with glucose activity. Alternatively, this mutual reaction may reflect the action of a manno-fructokinase. It has been shown that S^. mutans possesses an ATP-dependent kinase capable of recognizing both fructose and mannose (5). ATP may be generated by pyruvate kinase and this may be available in PEP-supplied cells. Table 6 indicates a role for ATP in sugar phosphorylation. However, this mechanism would not account for all the inhibition observed. As would be predicted, mannitol and methyl-3-D-thiogalactopyranoside (TMG) do not inhibit the reactions seen with glucose or fructose. In addition, both glucose and mannose, as well as glucosamine, inhibit 2-DG phosphorylation. Fructose is slightly inhibitory, however TMG also shows some competitive inhibition which indicates a degree of non-specificity to the reaction. If a "mannose site" exists on the glc PTS and 2-DG shows some affinity for this site, then one may postulate that this is the site where fructose is inhibiting. Since 2-DG is contaminated with glucose (see Methods ), it is difficult to interpret the data shown in Table 9 using this glucose analogue as a competitive

PAGE 86

77 inhibitor. Calculations showed that the amount of inhibition of glucose phosphorylation could be attributable to the contaminating glucose. However, PEP phosphorylation of mannose and fructose in the presence of 2-DG showed 15% and 100%, respectively, of control levels, clearly indicating competition in the case of mannoso. Also, the homologous system showed 94% inhibition. Kinetics of phosphotransferase activities and relative growth rates in various sugars It was of interest to deduce the relative affinities of the glc/man PTS for substrates. Kinetic analyses were performed and are illustrated in Figs. 7-9. The linear transformation by a double reciprocal plot was calculated by a linear regression analysis. The linear coefficient in the case of glucose was calculated to be .999. The for glucose phosphorylation was calculated from the x-intercept and was found to be 64 ym and the V^^^ which was calculated from the y-intercept was found to be .366 nmoles glucose-6-phosphate formed/min/ ml (Fig. 7). For mannose, the linear coefficient was calculated to be .987. The K^^ for mannose phosphorylation was calculated to be 90 pm and the V^^^ was .300 nmoles mannose-6-phosphate formed/min/ml (Fig. 8). Not shown is the kinetics of 2-DG phosphorylation, the K^^^ of this reaction was calculated to be 154 ym. Fig. 9 shows the results of a kinetic analysis using fructose as a substrate. The K^ of this reaction is 42 ym and the V is .176 nmoles fructose-6phosphate formed/min/ml. The linear coefficient in this case is .998. These relative affinities are reflected in the growth rates in these different carbon sources (Fig. 10). A transformation of these

PAGE 87

•a sc 3 C ta to -l-> 0 s_ ta •> X 0 CU E x> "O Q Q. (U p QJ i SZ n 1 0 ^ — •0 LlJ •r— cn 0 +-> 3 n 14_ 0 (0 0 40 CU un 0 CJJ (U Lf) 4-> l-> 0 oo •rCM C 1 c io (U >— Ln V3 OJ 0 ta Q i* +J (0 ra -M X Q. > Q +-> f— w cn [ J / — \ E x> P— ta in "4''^ rci -a 00 o CU to u ei -n 0 s. sz +-) 0 0 (/) If. sz i. c~ >— c i~ T3 0 "rCU
CL 0 SCU 4^ 3 0 tJ Q. S(O E '3 — CO Q sScu o 0 •4-> CU 0 (4 (/) n 1 C SXI to Q 0 s_ CU 0 0 M. ^ 3 •rf~ Ms(U 0 0 to Q_ O 5 rc c "O • 1 Q. 0 0 CU TJ < ta s OJ 10 c!3 (C c •!-> 0 OQ to 30 s_ ta (/I (/) *p0 n 3 n3 -0 >1 3 0 00 4J Q. (/) s: 0 > "O 0) ta to (0 E n (U i. 0) E CU ^s+-> to >^ 0 (O 2 sQ. E fO 1 Orc CU c ro +J >> 3 1 1 0 +-> 4-) CU c o sE +J 0 ta to c x: T3 3 0 s: ZD to fO sz (O Q. •r> E cu 0. CO a C_3 CU oc {/) •a O cn 0 ^ S0 CU 3. E 1 — cu sz 1/1 Q. 4-> 1 1 CL to -M <|T3 0 1 aj 1 (U •CJ X +J 0 CU 4J QQ to to to sCU CU C UJ c 0 cx c !-> a; a. 01 0 0 CU X U. -o ifl E C 3 to CU CU Q. 1— •r— (13 3 CU +-> cu C r— 0 fO CU CU o 0 •r+-> S5 +-> 0

PAGE 88

t 79 (|UJ/u!UJ/sa|OUJU) 31VHdSOHd-9-3SO0ni9

PAGE 89

-o 1 (U (U • (i> s_ •rQJ C7) (U o Jto 3. +-> iO O (O E >> CO c tj E E o o to ta 0) CO c QJ E JE s • o +J o S+-> QJ o QJ E o I/) 3 to +J T3 c 1 — O E E O) ( Q. sto o to So QJ E -M c 4-> QJ O s: o o E E +J Q. E o QJ •r— •r— Q. 0) • (/) o O +-> o o o O) ID 1— c o E X •1 — n3 o
1 — to 3 T3 Q) JZ Q. Q. 3 1 — OJ — x: -(J >) QJ O • •r— +J +-) -p SE o o — T3 SQJ >> (J 3 •r— QJ OJ E jQ (U • r— — >, > E Q •a 4> O E *r— C 3 c 3 o 3O E Q. O il • r— r— -l-> 3 to o I/) MO 4-> CO to O 1 — tc 0) > 3 • (_) ra o rO •a M TD u cu -l-> 1 — •r— *— E E o QJ l-> fd 4J (O o iS(0 E •f— -!-> to QJ SQ. (0 J= o to ^ — V 4Q. • E 3 O ID o CO I/) 00 s to ^ — ^ OJ +J O o 1 — tz • ifO sz 45E QJ ra -a Q. o D1 E •rOJ 1 o Ll_ o ID to • 1 — 1 JD 1 0) 0) r— S(_) — 3 OJ +J S^ — o 4-> O QJ to > -IJ QJ 1 QJ QJ ^ lO 1 — i. M 1 — E S_ < c 3 3 Q. 1 1 3 (0 (U O +J QJ 1 z. to e 1/5 X O Q 0) rtS (O E X Q. QJ r— to 4so E 0) O) X E •r~" O O) c QJ to 4c LD to c >1 i/J (/) o E (/) •r— r — O c X> •r— 3 ra •r— (C (U -p cn lO O c 4J s. £1 o •r+-> QJ tn re 0) 'r(0 Ll. E > +-> QJ c o fO OJ O QJ IT3 E •r--> C o S_ s: O O Q. Q. a. E •f— l/l o x: to E to O •o o QJ o to x: c QJ (0 JE 1/1 QJ 00 Q. >> QJ Q. SO 1 C7> +-) o OJ E to o QJ C CD 0) Q. to QJ S•f— O) •r— T3 E O QJ to U. X3 QJ c QJ c to to to s_ 2 0) E (T3 O QJ re QJ D. 4-> ro E SQ) Q>5 to to E 4J E Q. E 0) i3 3 na X o a to o O 21 OJ

PAGE 91

to QJ o to o to 10 o 3 3. re XI 1 3 CU 0) ^ t — -p (U CD CU E c o 4- LO +J CU s: re c l/l 1 "O sCU 01 o CU Q. E So • p— LTl <+to to Q. c (f. o o 0) to +-> +-> x: CU 0) o Q. o o. Q. f — OJ 3 o re CL Q. o Sto E 4/1 c X 4c re o (J QJ 1 CU +-> 0) 1 — 1 t/1 to re Q. Q .i— o re x> "O to t-> CD o (O o x: 4-> E to je: 0) to r— o to Cl. re Q. -(-) 0) Q. re -a CO I/) s_ E CU o CU o >, 3 q; M o E JZ to 4-> 4-> O a. (U CU "O XJ Ol 3 Sto CU "O vo to O 3 o >, CU c/l 1 lO 4- -> o +-) re 4-> 10 tfMd) 4-> o c o (U (0 T3 o Q 1/1 u to s_ iCU tre c f+J C to o o O +J jC re re to CU c o Q. 0) C to 3 lO So o o o to o o +-> CJ r— o sz u Q. CL x: re 3 E •Ij 4-> +-> sSo 4-> +-) HSre c c CD c c o (U CU &S o 5 o oj u L. c s ID c £3. OJ 0) o •r— Q. >> XJ o • o O) S"o CU o C3 CD 3 •pCU U. re to sS> re CU 3 3 O CU > to o so s_ O re X3 >> r — O) 4CU Q. 5 O E <:

PAGE 93

i/l -o o o +O O) (O o -o E 3 c o to 0) (/J o 4-> O o (J 1O O) Lf) 4-> O 4-> T3 sO) to c (O O 1/1 c o •-> s+J 1— c O) O) o c c •rO o ID O E (/) c IC +-> E C\J to to 3 3 O I— o Q. C 00 UJ >^ "4I— un o ^ h O 3 SCM +-> O 1— I— 3 So • o O 4r— • s i/i cn o O •1s_ c 3 O E o 0) u (U Q. to 4-> • < E sz • o 3 ta to to to 4-> 3 C o cu O) E o cu O 3 E t/) o
, +J o c o ^ sto O E to o ^ •IfO (O sso +-> 4c o E ai itn S0) cn s c •Ito E 4-> •rO fO 3 4-> OC -1O I— U ra

PAGE 94

85

PAGE 95

86 data into a semi-log plot allowed the calculation of the mean generation times. The mean generation time in defined mediun containing glucose is 75 min; whereas in mannose it is 290 min. With mannose as the growth substrate, a maximum culture density was not achieved after 12 h of growth. However, cultures grown in glucose reach their maximum levels between 5.5 h and 6.0 h under the conditions employed. The mean generation time using fructose as a carbon source was calculated to be 121 min and maximum growth was achieved in approximately 6.5 h. Selection of glucose phosphotransferase negative mutants Data presented thus far indicate that i. mutans GS5 appears to possess a relatively non-specific PTS for glucose uptake. In order to further study the former system, glc PTS negative mutants were selected on the basis of their inability to grow on glucosamine. Wild-type cells noninduced for the lac PTS are unable to grow in the presence of 2-DG since this non-metabolizable gl ucose analogue is transported by a constitutive system, thereby exhausting the PEP reserves. In addition, evidence will be presented below that suggests glucose represses the induction of the lac PTS. By selecting for a 2-DG resistant mutant, one may be selecting for a glc PTS-negative mutant, since 2-DG is transported by the glc PTS. However, the majority of the 2-DG resistant clones picked appeared to possess an altered glc PTS allowing them to transport and therefore grow on glucose (data not shown). In order to select true glc PTS-negative mutants, a second criterion, the inability to grow on glucosamine was used. However, glucosamine negative cells are not necessarily transport mutants. Clones which

PAGE 96

87 grew on lactose plus 2-DG but not on glucosamine were isolated. A priori one would expect a mutant selection based on these two criteria to provide glc PTS-negative mutant clones. In addition, glc PTS-negative cells were enriched by incubation in streptozotocin in the step prior to plating (see Methods ). After mutagenesis, the cells were allowed to recover in glucosamine. As shown in Table 7, this is a PTS substrate which is carried by the glc PTS (Table 9). Streptozotocin then was added to these cells. Since it is a glucose analogue containing a nitroso group and is carried by the glc PTS (33), those cells still containing an active glc PTS will transport this toxic analogue and should be killed. Streptozotocinresistant clones were chosen and allowed to grow in mannitol overnight. A survey was done to determine if the lesion indeed was in the glc PTS and the extent of loss of phosphorylating functions. In order to perform a general survey, the LDH/NADH-1 inked assay was used. The PTS substrates studied were glucose, mannose, fructose, and mannitol. Glucose, mannose, and fructose were used to determine what, if any, linkage exists between the PTS-mediated phosphorylations of these three sugars and mannitol phosphorylation was assayed as a positive control, since all cells were grown on this substrate. The results which are expressed as percent wild-type activity can be seen in Tables 10-12. Table 10 shows a representative listing of clones which grouped into what was labelled as Group I. These cells could not phosphorylate glucose or mannose; whereas fructose and mannitol phosphorylations were variable but did not correlate with the absence of glucose/mannose phosphorylation. Table 11 shows representatives of Group II. This group had much reduced glucose activity but with one exception mannose

PAGE 97

88 Table 10. Glucose phosphotransferase system negative mutants: Group I.^ Clone Sugar substrate for transport assay ^ Mannitol V3 1 UUUo C r ructose 3A 0^ 0 175 94 2A 0 0 125 35 4B 0 0 95 20 12B 0 0 54 24 188 0 0 42 59 168 0 NT^ 50 0 23A 0 0 26 0 ^Mutants were selected as outlined in Methods. Cells were grown overnight in TYE broth plus 20 mM mannitol, harvested, washed, and decryptified. The LDH/NADHlinked spectrophotometric assay for sugar phosphorylation was run on each clone using the indicated sugars as described in Methods In each case a 1 mM sugar concentration was employed. Results are expressed as percent wild-type. Not tested.

PAGE 98

89 Table 11. Glucose phosphotransferase system negative mutants: Group 11.^ Sugar substrate for transport assay^ Clone Glucose Mannose Fructose Mannitol 34B 0 83 45 8B 13 0 46 48 25B 16 0 104 42 26B 18 0 86 18 4A 20 0 NT^ 105 lOA 21 0 71 100 26B 21 7 129 59 28B 29 0 88 31 ^Mutants were selected as outlined in Methods. '^See Table 9 for assay conditions. '^Results are expressed as percent wildtype. Not tested.

PAGE 99

90 Table 12. Glucose phosphotransferase system negative mutants: Group II I. ^ Clone Sugar substrate for transport assay^ Mannitol Gl ucose Mannose Fructose 49B 30^ 21 155 97 23B 33 15 117 63 IIB 34 15 75 164 13A 49 17 NT^ 176 468 38 0 158 33 Mutants were selected 'See Table 9 for assay Results are expressed Not tested. as outlined in Methods conditions. as percent wild-type.

PAGE 100

91 activity is totally absent. Fructose phosphorylation was variable but always detectable. Group III clones are listed in Table 12. These cells were characterized by having up to 50% wild-type activity for glucose phosphorylation and, with one exception, residual mannose activity. Fructose activity was near to or above wildtype activity in all mutants which grouped in this category. These data suggest a glc/man PTS and a distinct frc PTS. Studies of cell -free membranes for phosphotransferase related activities The study of the glc PTS using isolated membrane fractions was undertaken for a number of reasons. The first reason was to obtain a more detailed picture of this transport system at the molecular and cellular levels. Second was to compare membranes with whole cells in order to evaluate the ability to maintain the native protein structure during the fractionation procedure; specifically when using the muralytic enzyme, mutanolysin (Ml), to remove the cell wall. Lastly, and most significantly, was to determine the site(s) of lesion(s) of the mutants described in the previous section. The latter studies are described in the following section. Since the primary focus of this study is the glc PTS, all membranes were prepared from glucose-grown cells. Mutanolysin (Ml) treatment yielded a more active membrane preparation than when mechanical means were used to break cells. The specific activity of glucose phosphorylation using 50 yf-l D-[^'^C(U)]-gl ucose by Ml-prepared cell membranes was 14.7 pmoles/yg protein whereas "membranes" obtained from cells broken by glass beads was 4.6 pmoles/ug protein. The Ml-derived membranes were 3.2-fold more active than the wall-membrane complexes obtained from cells broken by glass beads in a Bead Beater.

PAGE 101

92 Since a different glucose concentration (25 yM) was used to assay wallmembrane complexes obtained from cells broken in the Braun Tissue Homogenizer, it is more difficult to obtain an exact comparison. However, the results showed a specific activity of 2.3 pmoles/yg protein which again is clearly less than that obtained with Ml-prepared membranes. For this reason, all studies involved Ml-prepared membranes. Since PT-systems have two sets of reactions, one that involves soluble proteins and a second that involves the membrane proteins, it was of interest to determine if membranes prepared with Ml would show a requirement for cytoplasmic constituents to carry out the phosphorylation of glucose. The results are outlined in Table 13. It is apparent that the membranes alone are sufficient to carry out a PEP-dependent phosphorylation of glucose. The cytoplasmic material appears to be slightly inhibitory perhaps due to phosphatases present in the soluble fraction. These data should not be interpreted to mean that unlike the majority of PT-systems studied to date only membrane proteins are involved in glucose phosphorylation. As will be discussed below, cytoplasmic constitutients are quite likely trapped within membrane vesicles or perhaps these "cytoplasmic" constituents interact with the membrane as transitory peripheral proteins. Fig. n demonstrates the proportionality of the PEP-dependent phosphorylation of glucose with added membranes. This proportionality exists up to 160 yg of membrane protein. There appears to be some saturability indicating that at least one of the PTS proteins is present in a rate-limiting amount.

PAGE 102

93 Table 13. Glucose phosphorylation by cell fractions of S^. mutans 6S5.^ Fraction'' CPM Cytoplasm 0 Membrane 2025 Cytoplasm + Membrane 1160 Cells, previously grown in defined medium plus 5.0 mM glucose, were fractionated to yield membranes and cytoplasm as described in Methods The mutanolysin (Ml) procedure was employed for this purpose. '^Reaction mixtures (final volume: 200 yl) consisted of 80 mM PB, pH 7.0, 4 mM MgCl2, 4 mM 2-mercaptoethanol 10 mM PEP, 10 mM NaF, 25 yg membrane protein, 114 yg cytoplasmic protein, and 12.5 yM D-[^4c(LI)]glucose. The reactants were incubated for 30 min at 37 C after which time they were diluted with excess sugars and the count of radioactivity incorporated into phosphorylated products was determined as described in Methods.

PAGE 103

I/) (U c ta E 10 XJ O) o so ^sI a. o o E I— > o c (T3 >> X) I c 0) o c o (J O) -M O SQ. ex. o 4-> C 0) o I c o o (/) s-p X c o •r— -t-> o CU (/) -a c o c: +-> o Q. O O c o -p CO >> so J= Q. (/) O x: Q. 0) 1 o CO ^ to (/) to a. Q 0 0) to (O c to E (0 Sn3 >< PB O) fO in s SE 5: 0) E r— E or tn n (U Dl. D1 Q) •rS_ to o o sz o •r— P fO s•p ra O +J I— E -p 0 SfO S0 <4MT3 CU -0 T3 P CU CU T) 4-^ ^ jQ rtS 3 St/) • 0 (O 1 — Q. CU CU E to +-> 0) CU 0 sto S3 (U ra cu 0 3: 2 0 i. CU Irt Q. 1 — +-> Sra cu to SI +-> CU 0 Or~ 0 C/5 •1 — 0 CM 0 tE J= c Q. cu 0 ra to 0 to I— c CU (0 1 — 0 Sp cn •rQ. 0 p X •a (O CU ^ — CU — p CU c c CU
+-> ra XI (0 sQ 4-> CU c c t/5 > c 3 •P 0 0 0 CO CT) > (U i. -0 P c: 0 CU rO <+ra S. 0 X) C_5 CU •r— E X) X3 CU ra IB E CO 1 — S-

PAGE 104

95

PAGE 105

96 Table 14 demonstrates the PEP requirement to catalyze glucose phosphorylation by cell-free membranes and also, the inability of ATP to substitute for this compound in the PTS reaction. The omission of NaF from the reaction yields a slightly higher concentration of product than when it is included. The difference between these two results yields a result close to that obtained when both PEP and NaF are excluded. However since PEP is in saturating amounts this difference may not be significant. The detection of activity in the absence of both PEP and NaF indicates a residual energy reserve. The inclusion of NaF in the absence of PEP (last line, Table 14) supports this conclusion. These data suggest cytoplasmic contamination of the Ml -prepared membranes. As with whole decryptified cells grown in glucose, membranes prepared from glucose-grown cells are able to carry out the PEP-dependent phosphorylation of glucose, mannose, and fructose (Table 15). These specific activities simply illustrate that these membranes reflect the constitutivity of certain phosphotransferase systems found in the whole decryptified cells. No attempt was made to quantify the loss of activity during membrane preparation due to the complexity of the systems under investigation. The results of competition experiments using an excess of unlabelled sugar vs. D[^'^C(U)] -glucose are described in Table 16. The pattern is similar to that obtained with decryptified cells. Glucose competes with itself. Mannose and 2-DG appear to compete with glucose phosphorylation by membranes more efficiently than in whole cells; however, this may be a reflection of the lower concentration of labelled glucose used in studies with membranes (250 yM vs. 125 yM). In both cells and membranes.

PAGE 106

97 Table 14. Phosphoenolpyruvate-dependent phosphorylation of glucose by mutanolysin-prepared membranes of S^. mutans GS5.9 Conditions pmoles gl ucose-6-phosphate formed/yg protein'^ +PEP, +NaF 7.3 +PEP, -NaF 8.6 -PEP, -NaF 1.1 +ATP, +NaF 0 Cells were grown in defined medium plus 5.0 mM glucose and membranes were prepared using mutanolysin as described in Methods '^Membranes, 40 yg, were assayed for phosphorylating activity in the presence of 80 mM PB, pH 7.0,4.0mM MgCl2,4.0mM 2-mercaptoethanol 10 mM NaF, 10 mM PEP or 10 mM ATP, and 175 yM D-[^\(U)]-glucose (4.0 yCi/ymole) in a total volume of 200 yl. After 30 min at 37 C, the mixture was diluted in 1% cold glucose. Labelled glucose-6-phosphate was measured after filtration of products through a DE-81 filter and expressed as pmoles/yg membrane protein.

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98 Table 15. Phosphotransferase activities of mutanolysin-prepared membranes.^ Sugar substrate Sugar-phosphate produced pmoles/ug cell dry weight Glucose 12.2 Mannose 12.2 Fructose 12.5 ^Membranes were prepared from glucose-grown cells by the mutanolysin procedure (see Methods ). ^Membranes were incubated in 85 mM PB, PH 7.0, 4 niM MgCl^, 4 mM 2-mercaptoethanol 10 mM PEP, and sugar. The sugar concentrations were as follows: 175 viA D-[^^C(U)]-gl ucose (4 yCi /nmole) 100 yM D-mannose plus 10 yM D-D'*C]-mannose (48.6 yCi/ymole), or 100 yM D-fructose plus 1 yM D-[^'''C(U)]-fructose (359 yCi/ymole). In order to obtain a measurement within the linear range two membrane concentrations were used (24 and 48 yg). The reaction mixture (total volime: 200 yl ) was incubated at 37 C for 30 min after which time excess cold sugar was added to stop the reaction. Product was collected onto a DE-81 filter for counting in a liquid scintillation counter. Data are expressed as pmoles sugar-phosphate/yg protein.

PAGE 108

99 Table 16. Inhibitory effect of competing sugars on the phosphorylation of D-[^'^C(U)]-gl ucose by the phosphoenolpyruvate-dependent phosphotransferase system of decryptified cells and mutanolysinmembranes^ derived from S. mutans GS5. % Inhibition Competing sugar Membranes'^ Cells'^ None 0 0 Glucose 100 99 Mannose 93 74 2-Deoxyglucose 86 54 Fructose 7 1 Mannitol 0 0 Membranes were prepared by mutanolysin-induced lysis of glucosegrown cells (see Methods ). '^To assay membranes, the reaction mixture (final volume: 200 yl) contained 50 mM PB, pH 7.0, 2.5 mM MgCl^, 2.5 mf^ 2-mercaptoethanol 10 mM PEP, 10 mM NaF, 0 or 10 mM competing sugar, 125 yM D-[^^C(U)]glucose, and 50 yg membranes. Incubation was carried out at 37 C for 30 min. The amount of radioactivity adhering to DE-81 filters was determined as described in Methods Results were calculated from the percent decrease in cpm adhering in samples with unlabelled sugar as compared to cpm adhering in samples without unlabelled sugar (percent inhibition). Data presented in Table 9.

PAGE 109

100 fructose does not compete with glucose phosphorylation. It is evident, therefore, that membranes prepared by mutanolysin treatment mimic the cells from which they are derived; that is, it does not appear that the binding site(s) of the glc PTS has been altered. The transphosphorylation reaction was first described by Saier et al. (59) as a means of assaying for EII. As described, the EII^^^ catalyzes the phosphoryl exchange between glucose-6-phosphate and glucose. The ability of membranes to carry out this reaction was studied in order to determine the integrity of the EII^^'' in cell -free membranes. The reaction is known to be very sensitive to substrate inhibition and requires a ratio of phosphoryl donor to acceptor of 1000:1 to provide an optimum reaction. Table 17 outlines the optimization of the reaction with respect to substrate concentrations. The ratio of donor to acceptor was always 1000:1. The maximum product yield was obtained with 50 mM glucose-6phosphate and 50 yM ^^C-glucose. At 75 mM donor and 75 \M acceptor, inhibition was evident. The reaction was proportional to the amount of membrane protein added up to at least 195 yg (Fig. 12). This reaction, therefore, verified the localization and function of EII^^^ in membranes. As described previously, EI is able to catalyze the phosphoryl exchange from PEP to yield pyruvate (54). If ^^C-PEP is used, the amount of exchange can be determined by the amount of ^^C-pyruvate formed. Pyruvate is separated from PEP by reacting it with phenyl hydrazine to form its osazone derivative which is water insoluble and readily separated from PEP by partitioning in ethyl acetate. The observation that membranes alone could catalyze a PTS reaction indicated that EI and HPr are either trapped within membrane vesicles

PAGE 110

101 Table 17. Gl ucose-gl ucose-6-phosphate exchange reaction (transphosphorylation) by mutanolysin-purified membranes^ of S^. mutans GS5 as a function of reactant concentrations. ^^C-glucose (yM) 12.5 25 50 75 100 Gl ucose-6-phosphate (mM) 12.5 25 50 75 100 14 C-Gl ucose-6-phosphate (nmoles formed )b 0.18 0.64 1.32 0.90 0.74 Membranes were prepared from glucose-grown cells using mutanolysin as described in Methods. Membranes, 158 yg, were assayed in the presence of 50 mM PB, pH 6.0, 2.5 mM MgCl,, 2.5 mM 2-mercaptoethanol 10 mM NaF, and various concen14 trations of D-[ C(U)]-gl ucose (4 yCi /ymole) and gl ucose-6-phosphate in a total volume of 200 yl. The reaction was held at 37 C for 30 min and then diluted with cold H^O. This was held on ice until the products were separated from reactant by anion exchange chromatography. Glucose-6phosphate was eluted with 1 M Li CI 2 and the eluate was counted in a liquid scintillation counter. The concentration of product formed was calculated from the measured cpm as described in Methods.

PAGE 111

to 1 (U a r— +J o 1 — re sz to OJ o to 4-) cu o > •r> • tn 'oi f — Q. cu c ( — c 1 O 1 to CO o O ^ — ^ o CO 1 — 1 > to — ^ re •rto 1 +-> T3 +-> =D 1 — cu >> "O (0 +-> O ro CM ro t/l a> I 3 j:= r— O +-> o Q. 1/1 >> B +-> >> o ^ O CJ re to O) iC71.— +-> 3 s_ o o> 5: o SI 1 1 oi ^ c sz 1 o CL •r— Cl s: Q to +J X 1/1 to cu m E c "O E cu o to O o CU o x: ^ x: un 'l~ i~ CU o. >> Q. 4- r— • iCL) 1/1 CO o (O CU o re c: to C Sj:^ (O •o O (0 *. +-> ro cu sz S(1) o io c 1 CD o +J S3 +-> CU cu c •r(0 I — CD -(-> o re (U Q. CD cu to • +-> 0) +-> o E o CO ts: n3 £3. Q. CJ •p€ x: Q. E JZ to E CU OJ CU 1 o_ 1 o to E >i (U E cu o O to (U E to o 1 — • o o o CD 3 (O to M o •rCJ c 1 — S00 -t-> 0) •a 3 •f— CD cu c •u 1 — +-> re CU CD SE CD •rcu re (-> i1 1 J (O E \j — ^ (U T3 CU M o re to (/I CU +-> C o d) cu cu re o C CO o tn JSZ -)-> Scu o •rso -(-> re E 3 o 44J JCl E CU to to r— •o 3 o XI CJD o c cu o c u sc o Sc ro c tre m c E 3 (O 0) cu -M o OJ c -a X +-> cu CU cu ( — ro c •r— cu r— +J 1 — s2 o E o o cu re jD o +-> +-> 2 scu CD E Sa. re •r— cu cn T3 o (0 CL U_ E (U CJ p— cu re to >i •-> sC_) "3to I— 0) fO o cu p. (U CO fO E o o s(U to cu 1 o re E o O rc3 CM CM 2 re CM

PAGE 112

o {|Tf 002/S9|0UJU) 31VHdS0Hd-9-3S00niD

PAGE 113

104 or are associated with the membrane. As is evident (Fig. 13) membranes are indeed able to catalyze this exchange reaction indicating that they contain EI. Furthermore, this reaction is quite easily achieved by these preparations, being detected with as little as 2 yg of membrane protein. Since EI is generally considered to be a soluble enzyme, it was of interest to determine if it normally is located in the cytoplasmic fraction of S. mutans Table 18 outlines the distribution of EI during the fractionation procedure. As can be seen, 65% of the activity is associated with the soluble fractions; however, 35% is left in the membrane. Further attempts to wash the EI from the membrane with 100 mM phosphate buffer were unsuccessful. In addition, protein could not be detected in these additional washes. The small amount of activity in the second wash most likely represents EI present in the interstitial spaces of the membrane preparations. If closed membrane vesicles impermeable to cytoplasmic proteins are being formed, then trapped EI would not be expected to be eluted from such vesicles. Membranes prepa red from glucose phosphotransferase negative mutants vs. membranes prep ared from wild-type cells For this study cells of mutants and the wild-type strain were grown to mid-log phase in TYE plus mannitol. Membranes were prepared using Ml according to the description in the Methods section. Two PEP-dependent PTS reactions involving glucose and fructose were examined. The results of a typical experiment are outlined in Table 19. Of the three mutants studied, only one showed the ability to phosphorylate glucose using PEP as a phosphoryl donor. This was 8B, a mutant of the group II category. Interestingly, this showed 10% of wild-type levels which closely correlated with the activity shown by whole cells (13%, Table 11).

PAGE 114

in T3 cu 4f 1—4 LlJ te an ^ an o SQ) 4-> Q) (U o C (U •-J OsJ E • cu (U • 0) £Z j > _Q c ^c o E *^ C ftj o 1 •rE to _Q Q• fO r \ *fO Q a (O D +-* i_ O c t. e\ 1 Q_ lO o OJ o LO > > Q) (_J 4-> (U 3 a; cr to to Sto Q LO 4_} CO o /rt *o o >> J_ QJ to r >— o < >Q. cu CO cu 1 ^ Q_ (j +j CO > E CO o QJ t/) 4_) £Z o 3 to QJ s: c: CM c r \ E to So r— fK lO r >QJ CO O o O U. T— 1 Opi 10 CM 03 cu d) Q) UJ (/) Z CO 4— Q) 4_> CU o> ^ I/) o O C7> 4_) O n 1 ;3 s: l/> QJ QJ fO Cl fO OJ c1 — r 2^ E o •rj_ 1 1 +^ o *" fO 4_) QJ (/) *'~ 3Q_ x~ cu C "O n 1 • QJ cu Q t/l "O f-^ w j_ to (/) f~ E / 1 "O QJ 1 n 1 ni w c:3 >• fO O QJ — i O.J— "O f~ CD n 1 _c CO OJ [ y s: ^ ^ ^ +-> ;j ~o O c: QJ CU QJ 1 t r~ s: -o 1— (11 ri to H— E QJ (U CJ i_ to "O Q-) CO O 1 1 1 > VI (/^ Q < — d) c o t/5 t/) c >•rLf) •r— CU "n w CO ''^ -O > _o QJ QJ O. f— >3 rCI +-* •o r" Z3 "O si fO ( — (J O cz cz o O c (J (3 CD QJ c OJ q; (U -M C Q. o o 1 — 1 a CO to QJ QJ 0) +-> +-> o {/) sOJ +-> o o to O 2 c ^O E S1 +J Q) c QQ. rO X3 (/I to 1 J +-) o to f— > co ro Q) +-> to cu r— VI > +J •r— u o C_) o C 3 ro Sx: 00 u c 3Z o So >> SQJ +-> r— c (0 CU >^ QJ (0 J3 s !-> 1/5 o £3. o CO I/) E O S10 3 CO (+CO to GJ o a> >> cu E il £2. ST3 OJ (O E •r— 1— to +-> CU CU sto U. fO CU o c c Cl ss: O) o x: po •f— +J E Q. a 'oj N O) +-> to to 4-> E c (0 LO •rre U 0) o o s: o (0 CO tOJ *a E !-> E _J O •r— o C31 1— s_

PAGE 115

009/s8|oujd) 31VAndAd

PAGE 116

107 Table 18. Pyruvate-phosphoenolpyruvate exchange reaction as a probe for Enzyme I: Distribution of activity in cell -free extracts^ of S. mutans GS5. Cell fraction nmoles CPyr/ml cell extract^ Total activity % total b c Cytoplasm + wash 1 0.06 1.500 48 Wash 2^ 0.18 0.54 17 Membranes^ 0.44 1.10 35 Cell -free extracts were prepared by mutanolysin-induced lysis according to the procedure described in Methods. Cytoplasm refers to the supernatant obtained from the first centrifugation of lysed cells at 30,900 x ^ for 60 min. ^Wash 1 and 2 refer to wash supernatants obtained at 30,900 x ^ for 60 min from the first and second centrifugation of washed pellets after cell lysis. These washes were conducted using 100 mM PB, pH 7.0, containing 5 mM MgCl^ plus 5 mM 2-mercaptoethanol (see Methods). Membranes refers to the final pellet obtained after the above centrifugation and removal of whole cells. The reaction mixture contained a final volume of 100 yl and consisted of 40 mM tris-HCl, pH 7.5, 8 mM MgCl^, 10 mM NaF, 2 mM pyruvate, 0.2 mM phosphoenol[l-^^C]-pyruvic acid (10.6 yCi/ymole), and cell extracts. The extracts varied in the following manner: cytoplasm plus wash 1: 2.15 and 4.29 pg protein; wash 2: 92 yg protein; and membrane: 2-20 yg protein. Where more than one amount was used, nmoles/ml of extract/600 yl counted is expressed as the mean. The reactants minus PEP were preincubated at 37 C and the reaction was initiated by the addition of this compound. After 60 min at this temperature, the reaction was terminated by the addition of phenylhydrazine and '^C-pyruvate was detected as its osazone derivative (see Methods ).

PAGE 117

108 Table 19. The relative glucose and fructose phosphotransferase activities in mutanolysin-prepared membranes of mutant and wildtype strains of S^. mutans GS5.^ Glc PTS^ Frc PTS^ Strain pmoles glucose-6phosphate/yg membrane protein % Wildtype pmoles fructose-6phosphate/vig membrane protein % Wildtype Wildtype 12.2 100 12.50 100 3A 0 0 0.56 4.5 4B 0 0 0.57 4.6 83 1.2 10 0.47 3.9 ^Membranes were prepared from cells grown in TYE supplemented with 20 mM mannitol. The procedure as described in Methods was followed. Menbranes, 90-180 ug protein, were incubated with 85 PB, pH 7.0, 4 mM MgCl,, 4 mM 2-mercaptoethanol 10 mM PEP, 10 mM NaF, and 125 yM 14 D-[ C(U)]-glucose (4 yCi/vimole) or 100 yM D-fructose containing 1 \iA D-[^^C(U)]-fructose (359 yCi/ymole). The reactants minus the membranes were incubated at 37 C and the reaction was initiated by the addition of the membranes. After 30 min at 37 C, and the reactants were diluted with cold ^% sugar (homologous) and filtered through a DE-81 filter. The filter was counted in a liquid scintillation counter and the pmoles of derivatized product formed was determined from the number of counts remaining on the filter after extensive washing. To insure linearity, at least two membrane amounts were used and results are expressed as a mean specific activity.

PAGE 118

109 Frc PTS activity deviated from that seen in whole cells (Table 19). With whole cells, this activity ranged from 175% (3A) to 46% (8B) wildtype activity (Tables 10 and 11). However, membranes prepared from these cells showed significantly decreased activity when compared to decryptified wildtype cells. Possible explanations for this result are that the frc PTS is labile and/or there are stereochemical rearrangements of this system during membrane preparation. It was of i nterest to biochemically define the site of lesion in these mutants. Since in the bacteria studied to date the PEP-dependent PT-systems involve up to four distinct proteins, a series of experiments was undertaken to more closely determine the affected protein (s). It should be reiterated that very little detail of the PT-systems in the lactic acid bacteria is known. As detailed in the Introduction the model systems are based on studies of ^. coli and S^. aureus In these organisms sugar specificity lies in the EI I and EI 1 1 components (21). It was assumed that in this organism a similar situation occurred. Therefore, the transphosphorylation reaction was used to study the putative EI I in these mutant strains vs. the wildtype. The data outlined in Table 20 demonstrate that this activity was not evident in either of the mutant strains examined, including the leaky strain 8B. A second set of membrane preparations yielded corroborating results. As stated previously, this assay is specific for the transport protein EII (59). It was therefore concluded that the membrane-bound glucose carrier protein (s) had been irreversibly altered. In order to understand the physiology of these mutants in more detail, EI was assayed in both the cytoplasm and the membrane fractions.

PAGE 119

no Table 20. Transphosphorylatipn as a measure of Enzyme II in mutanolysinprepared membranes'^ of wild-type and mutant strains of S. mutans GS5. Strain 14 pmoles C-gl ucose-6-phosphate % Wild-type formed /yg membrane protein Wildtype 3.3^ 100 4B 0 0 8B 0 0 Transphosphorylation was carried out in an incubation mixture of 70 mM PB, pH 7.0, 3.5 mM MgCl2, 3.5 mM 2-mercaptoethanol 10 mM NaF, 50 mM gl ucose-6-phosphate 50 yM D-[^^C(U)]-gl ucose (4 yCi/Mmole) and membranes. To insure linearity two amounts of membrane protein were used, the lower ranged from 180-240 ug/200 yl (total reaction volume) and the upper ranged from 455-475 yg/200 yl. In the case of 4B, the amount of membranes obtained was limited; therefore, only the upper amount was used. The reactants were incubated for 30 min at 37 C and then passed through an ion-exchange column (see Methods ). After extensive washing with H2O the glucose-6-phosphate was eluted using 5 ml of a 1 M Li CI 2 solution. The eluate was counted in a liquid scintillation counter and the pmoles of product was calculated from the known specific activity. ^Membranes were prepared by mutanolysin-induced lysis (see Methods ) of cells grown to log phase in TYE with 20 mM mannitol. ^In a separate experiment five identical samples were assayed, each containing 126 yg membrane protein. The mean amount of 1 4 C-glucose-6-phosphate formed/yg protein was 4.3 0.9,

PAGE 120

The results using two separate membrane preparations are given in Table 21. There is considerable variation between experiments, however it is apparent that the EI levels approach wild-type when both experiments are considered. The one striking difference between wildtype and mutant strains is the distribution of this enzyme. In wild-type, the membrane fraction contains 50-75% of the activity. This differs somewhat from membranes prepared from glucosegrown cells in that membranes prepared from the latter cells contained 35% of the EI activity. However, both these results differ significantly from those obtained when extracts from mutant strains are examined. It is evident that only a small fraction, 3-13% of the total EI activity is detected in the membrane fraction; the majority of it separates with the cytoplasmic constituents. The reduced amount of frc PTS activity in membranes when compared to whole cells may be a reflection of a limiting amount of EI or of a defect in El-binding to the mutant membranes. An attempt was made to reconstitute the membranes with cytoplasm; however, this was unsuccessful. One major problem was the decay of EI in the soluble fraction. The lability of cytoplasmic EI was not studied systematically; however, it has been observed throughout these studies. For instance in one study the level of activity of the PEP-pyruvate exchange reaction decreased by 51% and 86% in 2 and 7 days, respectively. The EI from other organisms has been shown to be labile in purified form but stable to freeze-thawing when associated with membranes (77). The same assay (PEP-pyruvate exchange reaction) using membranes of S^. m utans showed onlya27% decrease in 5 days with repeated freeze-thawing. A second reason for the failure of the reconstitution experiments may be due to the impermeability of membrane vesicles to large molecular weight compounds.

PAGE 121

112 Table 21. The distribution of Enzyme I as determined by the phosphoryl exchange reaction between pyruvate and phosphoenol pyruvate in cell -free extracts^ of wild-type and mutant strains of S. mutans GS5. 14 pmoles Cpyruvate/yg protein/600 Experiment 1 % Total EI activity m1 found in cell fractions pmoles '"'Cpyruvate/yg protei n/600 Experiment 2 14, % Total EI activity yl found in cell fractions Wi Idtype Membrane'^ 48.6 52 276.0 77 Cytoplasm^ 44.0 48 82.3 23 Total 92.6 358.3 3A Membrane 17.0 13 12.6 6 Cytoplasm 117.0 87 212.9 94 Total 134.0 225.5 4B Membrane 21.3 5 9.1 3 Cytoplasm 379.0 95 283.2 97 Total 400.3 292.3 8B Membrane 21.5 10.0 23.1 13 Cytoplasm 200.0 90.0 153.0 87 Total 221.5 176.1 Cell-free extracts were prepared by mutanolysin-induced lysis of cells grown to log phase in TYE supplemented with 20 mM mannitol. The procedure followed is as outlined in Methods ^Cell membranes were incubated in 25 mM tris-HCl, pH 7.5, 12 mM MgCl2. 2 mM 2-mercaptoethanol 2 mM pyruvate, and 0.2 mM phosphenol [l-^^C]-pyruvic acid (10.6 yCi/ymole). Since membranes were suspended in PB, pH 7.0, this buffer was present at a concentration of 40 mM. A linear range of 5-45 yg of protein was assayed for each strain listed. The total assay volume was 100 yl. ^Soluble extracts were in tris-HCl, pH 6.8. Therefore, the following conditions were employed: 12 mM tris-HCl, pH 6.8, 10 mM MgCl^, 3 mM 2-mercaptoethanol, 10 mM NaF, 2 mM pyruvate, and 0.2 mM phosphoenoi[lC]-pyruvic acid. Amounts of extracts (2-12 yg protein) within a linear range of activity were used. The total assay volume was 100 yl. In b and c, the reaction mixture was incubated for 60 min at 37C and ^C-pyruvate was detected as its osazone derivative (see Methods ).

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113 Regulation of the lactose phosphotransferase system by monosac charides If non-induced cells are grown in a mediim containing glucose and lactose, a biphasic growth pattern is observed. This can be seen in Fig. 14. The first phase of growth coincides with the uptake of glucose whereas the second phase of growth coincides with the uptake of lactose. These two phases of growth are separated by a 60-min lag period. Glucose is completely exhausted from the mediun before lactose is taken up by the cells, and lactose uptake is preceded by a typical diauxic lag period. The lag period usually is indicative of the induction of enzymes required for uptake of the second sugar. This appears to be the case in these cells. The induction of the lac PTS, as measured using the chromogenic analogue ONPG, is concomitant with the second phase of growth and, furthermore, is not evident until the lag phase is completed. Its specific activity then increases coincident with the second phase of growth. Phospho-6-galactosidase, on the other hand, appears during the first phase of growth. This enzyme can be detected, albeit at low levels, in glucosegrown cells (data not shown). Its synthesis appears to precede the synthesis of the lac PTS (Fig. 5). The possibility exists that one of its products, galactose-6-phosphate, is an inducer of the lac PTS as has been found in S^. aureus (43). It appears that the presence of glucose is able to prevent induction of the lac PTS in S^. mutans. However, it does not appear to regulate lactose metabolism once the lactose dissimilating enzymes are present. This can be seen from the data listed in Table 22 where lactose-grown cells were grown in a glucose-lactose mixture. Even though glucose uptake seems to precede lactose uptake, once log phase growth commences

PAGE 123

in O) 4-> 10 S4-> 00 J3 1/5 0) CO o o .— +J •.< s. O) 4- •rO -a o CO a> (/) o o zs o> o> c •r— (/J IT) CO CD CO C (O --> 3 lO C/1 (O o OJ 4J CO OJ > s. (0 0) s o • o c 4-> ra me o • r— • CO rO 3 4J CM O) 1 — lO r — 1 O cn ^ C_J Ce oa 1 > 3 H+-> s: o •r+-> Sc ds 1— +-> c o o en x: U +J Q. >> CO rO 3 J-o (0 (O C > c c Q. CL (0 o > o c rO jQ tO CO J3 (O •> ^ CU fsO 4O O 4co 4J r— D> CO >> (O CO CO I CD E o S4T3 CU E 3 -o (U e CO XJ O 4J s: -a (U •rso CO -o o SCO CO s o pe a T3 2 t> c CO cu CU (O • c > cu •o cu CU •r— O o J= 4E c cu <+I 1— CU cu o c •r• (U -o s•r— 4-> TD e D. CO 0) CU SO "i s^ cu 3+J CU CO +J o o O CX3 5 (0 cu cu LlJ (0 S o -D D. Q. CO in +-> •o o + (O XJ c O c +J c c l-> ifl 3 (0 (O CD ra (U cr cu D. CO o X3 •a a o Ss0) C o n3 C71 E 4CO c 3 4ai CO o CU 4cu 3 c (0 c Q. E •r— s_ •rX CO 1 scu l/) CU O) 3 o 3 o +-> ( — 3 CO CO ^ CO c E (U to (U cu CO (U x: o So c +J u CO lO c o 3 o O) (U cu c T3 S1— c E SE o O) O) (O •r 2 Sa Si. 3 cu CU -o o CU T3 CO l/l o s CU 4l/l E CU (0 c CO •r(O •r1 (U OJ 3 a LlCM +cu E CL +J (U Q. o CO cu CO >> 1 U o 3 CO fO C7) o CU cu o CO E CO O CM Q. x: CO CU CO _l CO h< s_ o fO

PAGE 124

SlINn 1131)H

PAGE 125

116 Table 22. The growth and sugar uptake by cells of S. mutans GS5 induced for lactose dissimilation in medium supplemented with lactose and glucose.^ t 1 1 il U Lcb NieiL units Disappearance of suqar from medium Lactose^ Glucose^ 0 7 0 0 180 15 0 1.0 240 ?? 0 1.4 300 38 1.2 1.9 345 52 0.5 1.5 370 66 1.8 2.0 390 83 2.5 2.5 430 115 3.9 3.6 450 137 4.8 1.7 540 175 6.5 3.3 575 185 6.6 4.1 595 186 ND Cells were grown in 10 ml defined medium supplemented with 5.0 mM lactose (to log phase). This culture was transferred to 100 ml defined medium containing 5 mM lactose plus 5 mM glucose. At specified times, 5 ml aliquots were removed to measure Klett units. Cells were removed from this aliquot by centrifugation and the lactose and glucose concentration in the spent medium was detennined according to the methodology described under Methods I he concentration determined at each point was subtracted from the starting concentration and this difference, in mM, is the expressed Not determined.

PAGE 126

117 both are utilized in parallel. The slightly higher than expected concentration of lactose found in the medium most likely reflects carry-over from the inocul um. One interesting point is the apparent expulsion of glucose from cell at 450 min. This was a reproducible observation. Reizer and Panos (52) have shown that Streptococcus pyogenes exhibits a phenomenon they termed "inducer expulsion." When these cells are allowed to transport TMG (form ing phosphorylated TMG) and are presented with a metabolizable sugar such as glucose, free TMG is expelled. However, in the case presented in Table 22 it is the metabolizable compound being excreted. A simpler explanation may be that free glucose, formed from the cleavage of lactose phosphate to galactose-6-phosphate plus glucose, is being expelled in favor of the gl ucose-6phosphate formed by the glc PTS. Fig. 15 shows the results of growing non-induced cells in fructose and lactose. Growth is not apparently biphasic. However, a semi-log plot of the data reveals two growth rates; the first is equal to a generation time of 82 min and is commensurate with fructose uptake (0-310 min; Fig. 15) while the second rate is equal to a generation time of 225 min and commences with lactose uptake (310 min; Fig. 15). Lactose uptake does not begin until the exhaustion of fructose from the medium. However, as is demonstrated in Fig. 15, a diauxic induction period is not evident. Indeed the induction of the lac PTS proceeds parallel with growth (inset Fig. 15). A semi-log plot of the data confirms this conclusion. The frc PTS does not vary since it has been optimized by prior growth in fructose.

PAGE 127

•T3 (U U i(U c +-> o ra o i+j +J 1 — et us re p— Q. 2 in so T3 >> rO (/) 1/1 to -o c to I c u c o o 0) 4-> Q. to o o to > o u_ OJ to (0 Q. o. -U SD. 1 CO o 0) o St 1 I— C_3 Q) -M Qj 3QJ 1 — 1 (/) CU +J O £^ 2: 0 CO QJ QJ :3 Q. O E 0 -0 0 O o CM •M 0 t/) ^z 0 (tJ QJ to 0 c >— (/) CO 1 % Ifto r— to ii o GJ cu lO 0 £Z to li 1 0 SI J cC to o "O O. CM — a; fO •p3 0 -Q CU LO O It c >— (/) 01 CO r~" to O .u E 1 *o CU 0 C p1 .cz > — i r~ • *^ 1 — t^ o CO CO -M 0 E •r~ "O QJ d LO to CU LU • 0 cu CM (/7 0 "O CO u 0 cu >CO +-> fO 0) >> 1 — O LO r >— to Oc -M o < — c "3QJ • (l a. 0) S — C31 E E. r— 0) 3 \U o o 0 Q 0 > •rC SCQ 4-> s_ to 0 r— ra XI Q. O -M o 3 Q. sz CM 0 cu Q. E fO o t/) to 4-) F U t/1 a. E 3 QJ cn rO 2: (U Ol QJ to to S. 13 CU E ST3 0 QJ c: 1 — •o ME QJ S_ 0 o OJ E OJ •r< 0 CU i. 3 & c 3 sto to QJ 4-> a to 0) -M hLn 1— +-> X OJ c o s C ro r— to •r™ +-> to S(U O) 0) •rQJ E to c -o o >> o cu 0) to •0 E 0 E 0) o 3 X) +J c t3 0 0 Q. sto Q. c ao c •4-> 0 to +J QJ to c 'f o (0 cu CM to (J +-> QJ x: MQ. 3 QJ to to s_ M o (U to > cu to SO) to QJ to CL s. T3 o o -M 3 M3. 0 0 t. to X o a> to E c CU to 1 +-> QJ QJ s0) CU (0 o 1 — 1 0 0 to QJ ej E E S•a CM 3 to c >1 t/) c ZD 1 SQJ 4J 0 o +-> CU -P n3 cx> 4S. •r— OJ r— CD • -C r — Q. 4-> 4-> > Ln O) o •r— CU •r1 r to 1 — U QJ CU 14CO Ms 0 to X SQJ >> •r— p— 0) 0 QJ "prO (J to >> 4-> c QJ r™ u. JZ +-> (U Q) 0 a> So 3 U un to •o QJ o 4-> c sc sx: to c i. T3 cC rd 4-a +-> Q) E to to

PAGE 128

119 SUGAR UPTAKE (mM) P o o o o ^2 c\i — SlINfl 1131>1

PAGE 129

120 In order to compare the regulation of the lac PTS of the mutant strains with the wild-type strain, cells were pre-grown in TYE plus 20 mM mannitol for 16 h. As can be seen in Table 7, cells grown under these conditions do not exhibit a lac PTS. A 5% inoculum of such cultures was used to inoculate a Klett tube containing TYE supplemented with 5 mM lactose and 5 mM glucose. The tubes were incubated at 37 C in a candle jar and at various times, the absorbancies were measured in a Klett-Summerson photometer. The results of a growth curve using the leaky strain 8B and wild-type cells are shown in Fig. 16. With wild-type cells, as expected, two phases of growth are separated by a lag phase. Transformation of these data via a semi-log plot (not shown) revealed two growth rates. The first phase of growth showed a mean generation time of 64 min whereas the second phase corresponded to a mean generation time of 120 min. The mutant strain exhibited a uni phasic growth pattern and this was confirmed by plotting the data as a semi-log graph (not shown). Here, the mean generation time was calculated to be 71.6 min. Interestingly, the cell density was equal for both strains suggesting glucose was fully taken up by the mutant strain. As previously noted, strain 8B is leaky for the glc PTS. In addition, S^. mutans has been shown to contain an alternative glucose transport system which plays a significant physiological role under high glucose conditions. PTS negative mutants of S^. nutans have been shown to transport glucose (18). Fig. 17 demonstrates the differential onset of ^^C-lactose uptake by wildtype cells as compared to the mutant strain 3A in the presence of glucose. This mutant strain appeared not to have a functional glc PTS (Tables 10 and 19). (Interestingly this strain did show delayed

PAGE 130

Fig. 17. Lactose uptake by S^. mutans GS5 (v/ild-type) and a glucose phosphotransferase negative mutant as a function of cell density. Cells (wildtype or glc PTS" mutant strain 3A) were grown for 16 h in TYE plus 20 mM mannitol. After harvesting, washing, and concentrating two-fold, a 10% inoculun was used to initiate growth in 10 ml TYE supplemented with 5 mM glu12 cose plus 4.9 mM lactose (4.7 mM [ C]-lactose plus [^^C]-lactose: 0.97 yCi/ymole). Cultures were incubated at 37 C in a candle jar. At specified absorbancies as read on the Klett, aliquots (0.1 ml) were removed to measure counts per minute (cpm) taken up by cells (wildtype A 3A • ).

PAGE 131

sliNn ii3n>i

PAGE 132

1/1 <0 ro sJ4-> to +-> tz c (O la 4-> 3 CLI (/I OO O I— JZ o. Q. O dJ .— t/j cn O u s3 O D1 d) Q. +-> O I c -o (O I— a CO Z3 sOl (U CU S/> O 00 c (U -t-> to c o o u ire O) r— Q. Q. O) E I > T3 Iin r— lo 3 3 2 O) — o Q. !-> >> to T+J 1— c I 1— c I— o E tn to o t/5 O CM 3 to 1 1— 3 a>i— Q. t/> 3 UJ .— >CL[— to 10 c re 3 to •!4O O -l-> ^ O x: re to 4> r— I— Ic^ 5-Io t!3 44J C C re 2 to +-> o r3 S_ E C75 CT) CU CU •r> S_ u. -,CU -i-> 3 re CU CQ C 00 o i/i s: XI E to LD T3 CZ re •rS_ 3 re -3 CU CU +-> I— C TD CU C E re CU cj r— • £3. re 4-> a. +-> 3 c re \— CO D) c c •r!-> •!re to x: 3 4-3 T3 S CU -o O 4-> CU s_ re scnja 3 3 10 so re o c
PAGE 133

KLETT UNITS

PAGE 134

125 but sparse growth on TYE agar plates supplemented with glucosamine.) The experiment in Fig. 17 was done by transferring cells grown in TYE plus 20 mM mannitol to Klett tubes containing TYE plus 5.0 mM glucose and 4.9 mM lactose of which 0.21 mM was the radioisotope. At specified times, an absorbancy reading was obtained and 100 yl were removed for filtration through a 0.2 y membrane filter (Gelman Sciences, Inc., Ann Arbor, MI.). These filters were washed extensively with H2O and transferred to 5 ml Aquasol for counting. Duplicate counts were obtained. (A second experiment using centrifugation yielded similar results to those shown here. ) Lactose uptake, as measured by counts incorporated in cells (Fig. 17), correlated with growth of the mutant strain (data not shown); that is, even in the presence of glucose, the mutant's primary growth substrate is lactose. On the other hand, even though lactose internalization is evident at an early stage of growth in wild-type cells, it is significantly depressed and this repression is not relieved until the second phase of growth as would be predicted by data shown in Fig. 14. Thus, there are two phases of lactose uptake. The first phase of uptake is at a low rate, 5.2 cpm/Klett unit, and may be explained as the internalization of inducer. The second rate, 19.4 cpm/Klett unit, most likely represents the utilization of lactose as a growth substrate. These rates can be compared to that observed for 3A which was 40 cpm/Klett unit. Rates were obtained by linear regression analysis and these data gave correlation coefficients of .985 for 3A and .952 and .872 for the first and second phases observed for wild-type.

PAGE 135

126 As in other growth studies (Fig. 16), two growth rates were obtained with wild-type culture (mean generation times 95 and 165 min, respectively). However, a single growth rate with a mean generation time of 109 min was observed for strain 3A. The maximum growth obtained for this strain was less than with wild-type. As measured in Klett units, the maximum absorbancy was 140 vs. 170 for 3A and wildtype GS5, respectively. The cell densities achieved by the respective strains reflect the amount of glucose left in the spent media. In a separate experiment where glucose was measured, wildtype 6S5 exhausted the glucose (5.0 mM) from the medium whereas 2.2 mM remained after the cessation of growth of 3A. Again, uptake of glucose by mutants may be explained by the observation of Hamilton and St. Martin (18) that the alternate glucose transport system is a low affinity system.

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CONCLUSIONS Previous studies on the cariogenic organism, S^. mutans have been concerned mainly with the ecology of this organism, with emphasis on the initiation of pathogenesis. Not emphasized in the literature is the relationship between the physiology of this microbe and pathogenesis. In the oral cavity, S^. mutans adheres to the tooth surface where it establishes a focus of infection through various mechanisms. These mechanisms of adherence have been studied extensively (13,14) and will not be discussed here. Cariogenicity is actually caused by metabolic events occurring subsequent to the initial attachment of the bacteria to dental enamel. It is the production of lactic acid from the fermentation of a select group of sugars that is responsible for the degradataion of the dental structure (13,14). Knowledge concerning the physiology of this streptococcus, however, is minimal. It is known that this organism grows on a limited number of sugar substrates and that these substrates are transported by PEP-dependent phosphotransferase systems (Table 1). This mode of transport can be viewed as the first step in fermentation since the sugar becomes modified to the first intermediate in its dissimilatory pathway during passage through the membrane. This dissertation describes the glc PTS in detail since this sugar very likely represents the major carbon source of S. mutans. Since basic knowledge concerning transport of this monosaccharide 127

PAGE 137

128 is limited, the focus of this project was general in nature. This study deals with three interrelated areas and in each case attempts to obtain a basis for further study. The objectives of these areas were thus: (1) to obtain a general description of the glc PTS in whole cells (e.g., its substrates, under what growth conditions it is present, etc.); (2) to obtain a more detailed description of the glc PTS by using cellfree extracts (e.g., the existence of soluble vs. membrane factors); and (3) to definitively ascribe a regulatory function of the glc PTS over the transport of other PTS substrates. General studies on the glucose phosphotransferase system The methodology used for these studies allowed the study of phosphorylation rather than actual transport. The reason for studying phosphorylation was the convenience of this approach and the elimination of competing alternate systems (i.e., those driven by the proton motive force or by ATP) when decryptified cells were used. Much evidence has been accumulated to strongly suggest that group translocation (i.e., phosphoryl transfer) and transport are tightly coupled processes in intact cells. These include the inability of EIand HPr-negative mutants to accumulate sugars even though the permease (EII) is present. Sugars are not transported even by means of a facilitated diffusion mechanism in such mutants (21,49,78). More direct evidence comes from the work of Kundig and Roseman (30) showing that the isolated PTS components require a membrane factor to ultimately transfer the phosphoryl group to a sugar. Parallel work in the laboratory of Hengstenberg (26) demonstrated the tight coupling of phosphorylation and transport in S. aureus Thus, by defining PEP-mediated phosphorylation, one is also describing the underlying mechanism for transport.

PAGE 138

129 Using the two glc PT-systems of E. coli as models, initial studies were concerned with defining the nature of the glucose system (s) in _S. mutans Early in the study, it was observed (Tables 7 and 8) that S^. mutans GS5 was capable to transporting 2-DG but not a-MG. In addition, glucose, mannose, and 2-DG transport systems appeared to be constitutive. This indicated that a system somewhat analogous to the low affinity system of g. coli exists in this organism. But in E. coli this system transports glucose, mannose, 2-DG, and fructose (21,30). Upon further investigation, S^. mutans appeared to diverge from the above model. Results of competition experiments (Table 9) showed that glucose and fructose do not share receptor sites on common PTS proteins. However, the results of these experiments supported the conclusion that a glc/man PTS was present in this organism and that this system also utilizes 2-DG as a substrate. A puzzling observation was the competition between fructose and mannose. This observation may be the result of one or a combination of several factors: (1) the existence of an ATPdependent manno-fructokinase, (2) the affinity of the frc PTS for mannose, or (3) the affinity of a "mannose site" on the glc/man PTS for fructose. Kinetic studies indicate that the affinity of the glc/man PTS is: glc>man> 2-DG. The relative growth rates of these cells in the various carbon sources reflect these relative affinities. Thus, the value for glucose and mannose are 64 yM and 90 yM and the mean generation times are 75 and 290 min, respectively. An observation relevant to this discussion is the extent of inhibition each sugar exhibited for the heterologous phosphorylating reaction (Table 9). That is.

PAGE 139

130 glucose is a more effective competitor of mannose phosphorylation (96% inhibition) than mannose is of glucose phosphorylation (74% inhibition). Since growth in mannose is so poor, it would seem that factors in addition to the relative affinity of the PTS for mannose must be responsible. Interestingly the V for these two sugars are similar (.366 nmoles/ ^ max min/ml for glucose vs. .300 nmoles/min/ml for mannose). This further indicates a factor beyond transport is responsible for the limited growth of S^. mutans in mannose. Results obtained using mutants selected specifically for glc PTS negativity confirm and extend the conclusions presented thus far: (1) in this organism there are distinct PT-systems for the transport of glucose and mannose vs. fructose, and (2) glucose is a more efficient substrate for the glc/man PTS than is mannose. Tables 10-12 demonstrate these points. Apparently tight, as well as moderately leaky, glc PTS negative mutants were unable to phosphorylate mannose (Tables 10 and 11). Mutants with an increased capacity to phosphorylate glucose (30%; Table 12) showed moderate mannose phosphorylation. This pattern again suggested that the glc/man PTS phosphorylated (transported) glucose more efficiently than mannose and that a minimum amount of glc PTS activity (30%) must be present before mannose phosphorylation is evident. Fructose phosphorylation again appears to be an independent reaction in the selected mutants described here. One of the more relevant observations in terms of the overall physiology of this organism is the determination of a separate frc PTS. This organism is characterized by its possession of gl ucosyl transferase on its cell surface. This enzyme synthesizes capsular dextran from

PAGE 140

131 sucrose providing the cells with the capability of adhering to enamel surfaces. In order to obtain the glucose necessary for dextran polymerization, sucrose is first split by this enzyme with the formation of free fructose (13,14). Fructose is then taken up by the frc PTS and metabolized to lactic acid. Thus, fructose plays an important role in the physiology and pathogenesis of this organism. Collectively, these data indicate that the glc PTS resembles the low affinity glc PTS of E. coli This is in agreement with results obtained in other laboratories working on S^. lactis (86) and S^. faecal is (27). Unfortunately, the work in these laboratories and the studies presented here are inconclusive. To demonstrate the molecular nature of the glc PTS in S^. mutans it is necessary to isolate and describe the individual PTS proteins as was done for £. coli (30,31; see Introduction ) and S^. aureus (26,76,77; see Introduction ). The next section describes an attempt to set a foundation to begin the molecular dissection of the glc PTS in this species. Studies on the glucose phosphotransferase system using cell-free extracts Establishing the various parameters for glc PTS activity in whole cells allowed this work to progress to the level of the cell-free system. The method of choice for preparing such a system was that described by Siegel et al. (75) using the muralytic enzyme, mutanolysin (Ml). Before proceeding with any further discussion concerning the use of cell -free extracts, it would be helpful to examine the advantages and limitations involved in the use of this methodology. The advantages of using Ml-prepared membranes, i.e., membranes prepared from osmotically-lysed cells vs. membranes prepared from

PAGE 141

132 mechanically-broken cells were: (1) the higher specific activity in terms of the PTS, and (2) the apparent reflection of these membranes of the physiology of whole cells, at least as measured by the various PT-systems. This latter conclusion was based on the comparison of the PTS-induction patterns (Table 15) and the results of competition experiments (Table 16) using membranes with data obtained using permeabi 1 i zed cells. Membranes prepared from glucose-grown cells were able to phosphorylate glucose, mannose, and fructose; thereby reflecting the constitutive nature of these PT-systems found in the cells from which they were derived. The pattern of competition using radiolabelled sugars, as elucidated in whole cells, was reproduced using membrane preparations leading to the conclusion that a glc/man PTS, which also phosphorylated 2-D6, and a separate frc PTS were present in these membranes. A third advantage of using Ml-prepared membranes is the resulting clean separation of cell -wall and membrane material one is able to attain with this procedure (75). Presinably, this yields a purer product than that obtained with mechanically broken cells; Siegel et al (75) demonstrated this to be true with respect to contaminating cell wall. A major limitation of this technique can best be understood by discussing the data presented in Tables 13 and 14. Table 13 demonstrates the fractionation of PTS activity during the separation of the cellular extract into the "cytoplasmic" and "membrane" fractions. It is evident that all the activity is retained in the membrane fraction. Theoretically "purified" membranes should not be active since they should lack the soluble proteins. However, the apparent activity found in these membrane preparations brings into question the absolute purity of these membranes.

PAGE 142

133 As stated, it has been demonstrated that membranes prepared by Ml are free of cell wall; however, results presented in Table 14 indicate cytoplasmic contamination, albeit at low levels. These results demonstrate that in the absence of PEP and NaF phosphorylation of glucose still occurs; thus, indicating the presence of an energy reserve in the form of glycolytic intermediates. The inclusion of NaF inhibits this background suggesting the presence of at least one glycolytic enzyme, enolase, since NaF specifically inhibits that enzyme. These data may be interpreted to mean that membranes containing cytoplasmic constituents may be present in these preparations. In order to pursue some of the more intriguing results obtained during the course of these studies, it would be necessary to purify these membranes further, if possible, perhaps by sucrose-density gradient centrifugation or by hydrophobic chromatography. A second disadvantage is the limitation on quantitation inherent in the methodology. The conclusion that Ml -prepared membranes were physiologically similar to the cells from which they were prepared was a qualitative one. It is evident from Table 15 that an appreciable loss of activity occurred during the isolation procedure. Quantification of this loss was not made due to the inability to accurately measure the amount of membrane protein. For instance, the amount of Ml contaminating the cell fractions was unknown; therefore, the amounts of membrane protein employed in these studies reflected relative rather than absolute amounts. One can only speculate as to the reasons for the apparent loss of activity. For example, it may be stereochemical rearrangement of membrane proteins within the lipid matrix, a denaturation of the proteins, or a loss of "soluble" PTS proteins such that HPr and EI are in limiting quantities.

PAGE 143

134 One question yet to be answered is the extent of permeation of these membranes by large protein molecules. One assumes that these membranes form closed vesicles in aqueous suspensions based on thermodynamic considerations. However, the extent of leakiness of such "vesicles" remains to be demonstrated. This is a question which repeatedly presented itself throughout the many aspects of these studies. For example, Table 13 illustrates the failure of cytoplasm to increase the glucose phosphorylating activity of cell-free membranes. The question then can be raised as to whether this failure is due to the impermeability of these "vesicles" to high-molecular weight compounds or the presence of saturating amounts of EI and HPr in such preparations. It is obvious that answering this question would shed some light on the molecular interactions of the individual PTS components in this species. The major advantage of a cell -free system is that it allows the study of the individual PTS proteins. The two proteins examined were q1 c the EI and EII^ The reason these were chosen for study is that assays have been described for each (56,64) that allow direct measurement, thereby obviating the need for the isolation and purification of these protei ns S. mutans does appear to have a typical EII^^^ in that it not only functions in transport utilizing PEP as a donor but also catalyzes the phosphoryl exchange reaction (transphosphorylation; Table 17) between glucose and gl ucose-6-phosphate. It has been reported that the transphosphorylation reaction is one-tenth as active as the PEP-mediated reaction (56). For example, a typical result of 23.5 pmoles product/yg membrane protein when PEP is used is to be compared with 2.3 pmoles

PAGE 144

135 product/yg membrane protein when glucose-6-phosphate is the phosphoryl donor. In both these cases products and reactants are separated by ion exchange chromatography. It may be noted that there is a discrepancy in the results when comparing those obtained by chromatography vs. those obtained by ion-exchange membrane filtration (Table 15). This is due to the greater capacity of the columns. It is for this reason that columns are used in the transphosphorylation assay (Dr. G. Jacobsen, personal communication). Since cold glucose-6-phosphate is in 1000-fold excess of 14 C-glucose, the cold compound occupies many of the binding sites on the filter and thus effectively competes with the ^^C-glucose-6-phosphate formed. This permease (EII^^^) is similar to that described for £. coli (59) in that the transphosphorylation reaction is subject to severe substrate inhibition (Table 17). The exchange reaction between PEP and pyruvate allowed the identification of an El-like protein in both the membrane and cytoplasmic fractions. The observation that membranes alone catalyzed a phosphorylation reaction with PEP as the phosphoryl donor implied the presence of such a protein; the direct demonstration of this reaction demonstrated conclusively that this normally soluble protein is contained in these cell-free membrane preparations. Membrane vesicles prepared from E^. coli by osmotic lysis (Kaback vesicles) have been shown to contain low levels of EI and PEP (12). Indeed Kaback (28) used these vesicles to demonstrate PEP-dependent transport. Results from studies of fractionation of EI activity during membrane isolation reveal that the major activity (65%) fractionates with the cytoplasm. However, a significant amount (35%) fractionated with the

PAGE 145

136 membranes. Elution of EI from membranes using phosphate buffer was uncussessful This simply may be due to an incorrect choice of buffer. Alternatively, it may be due to the trapping of EI (and presimably HPr) in closed vesicles. The trapping may be the result of non-specific events involving the closure of membranes in the presence of cytoplasm or it may be the result of a specific association of EI with membrane receptors (perhaps the EII^^^) if, in fact, EI (HPr) are extrinsic membrane proteins. Singer and Nicholson (79) define extrinsic membrane proteins as those which fractionate with membranes but can be removed by relatively gentle means such as changes in ionic strength or pH. However, this definition does not always allow for conclusive results. The possibility that EI is an extrinsic membrane protein is supported by three observations. The first is the increased lability of EI observed in cytoplasmic vs. membrane preparations. The membrane association may protect EI from oxidation. The second is the significant fraction (35%) of the total EI (cytoplasm plus membrane) isolated in the membrane preparation. However, the possibility exists that much of the cytoplasmic EI content was oxidized before assaying and therefore the amount calculated was greatly underestimated. A third more interesting observation comes from the study of mutant membranes. Membranes prepared from the various classes of glc PTS negative mutants appeared to have normal levels of total cellular EI, however the amount within the membrane was greatly reduced when compared to wildtype membrane (Table 21). It is attractive to suppose that the EII^^^ is a major receptor for EI (or HPr) and the loss of a normal permease results in the inability of EI to maintain its membrane association.

PAGE 146

137 Again, it is difficult to draw definitive conclusions from data obtained using mutant membranes since complete separation of cytoplasm from membranes was not achieved. For instance in Table 21 experiment 2, 77% of the EI was detected in the membrane of wildtype cells which is a much higher amount than had been previously observed. One reason for this higher figure may be related to the distribution of protein, 47% of the total protein was found in the membrane fraction in this experiment as compared to 20% in the first experiment. This is certainly the result of cytoplasmic contamination. The above discussion highlights the necessity for obtaining pure membranes in order to be able to draw reliable conclusions. However, these data do demonstrate the existence of an El-like protein in this organism. Its cellular localization is a more difficult determination. One approach that may shed some light on this question is to determine the sidedness of these presumed membrane vesicles.. If, for instance, 35% of the vesicles are inside-out then 35% of the extrinsic proteins (e.g., EI) would be washed from the membrane with low ionic strength buffer. Of vesicles prepared by osmotic lysis of E. coli approximately 15% are everted, whereas 95% of the vesicles prepared in the French pressure cell are everted (12). Vesicles possibly existing in Ml membrane preparations would be expected to be similar to Kaback vesicles since both are prepared by osmotic lysis. The determination of the absolute amount of everted vesicles would be quite instructive in planning further experimentation. Regulatory functions of the glucose phosphotransferase system In addition to transport, the glc PTS in E. coll Plays a role in regulation

PAGE 147

138 of other transport systems (see Introduction ). One of the main objectives of this study was to assign a regulatory role specifically to the glc PTS as has been done for this enteric microbe. A regulatory role for the glc PTS in S^. mutans has been controversial. Hamilton and Lo (16) observed a diauxie when S^. mutans was grown in a combination of lactose and glucose; however, this diauxie was "leaky" in that glucoseinduced repression was concentration-dependent. The growth history of the cells used in their experiments was not presented so that it is difficult to evaluate the results obtained by these authors. Thompson et al. (90) and Heller and Rdschenthaler (23) have argued in favor of a competition of the various PT-systems for the common components, with certain systems being more efficient (e.g., glucose) than others and causing an inhibition of the less efficient systems (e.g., lactose). The data presented in this dissertation demonstrates the repressive effect the glc PTS has on the synthesis of the lac PTS in cells of S^. mutans (Fig. 14). This regulatory role appears to be specific for the glc PTS. The frc PTS is able to inhibit the transport of lactose but not repress the synthesis of the lac PTS (Fig, 15). It is possible, therefore, that regulation may be dependent on a functional EII^^^. In order to clarify the role in the regulation of sugar transport of the glc PTS in general and the EII^^^ in particular, glc PTS" mutants were utilized. The rationale for taking this approach was to obtain a mutant sub-strain of GS5 showing wild-type PTS activity for all PTS functions except those unique for glucose and in addition was able to transport glucose through an alternate system. This would provide a valuable

PAGE 148

139 tool for the study of the hierarchy of sugar uptake. Before proceeding with a discussion of regulation, I must diverge and discuss the nature of the selected mutants to meet the above criteria. Tables 10 and 11 outline the various PTS activities of selected clones. The clones I chose to study were based not only on these activities but also on their growth characteristics. One of the most promising mutants was 3A since not only did this strain appear to have equivalent or greater levels of the frc and mtl PT-systems when compared with wild-type strains indicating wild-type levels of EI and HPr, but 3A also appeared to have the same growth characteristics as the wild-type in TYE plus mannitol. Other mutants chosen to be examined in detail were 8B, 48 and 2A (data not presented). All mutant strains appeared to contain EI, as measured by the PEPpyruvate exchange reaction, at levels comparable to wild-type. When cell -free membranes of mutants were prepared, they were found to be similar in their lack of functions dependent on EII^^^. All but 8B lacked PEP-dependent glucose phosphorylating activity. Mutant 8B membranes showed 10% of wild-type levels of the latter activity; this result is in agreement with that obtained with whole cells (13%, Table 11). These data substantiated the conclusions reached with studies using the whole cells from which these membranes were derived, namely, that the lesion is in the sugar-specific component(s) of the glc/man PTS. Contradictory, however, were the results obtained when fructose was used as a substrate. Unlike studies with intact cells, all membranes derived from mutants showed reduced levels of PEP-dependent phosphorylation (Table 19). It is difficult to determine the reason for this discrepancy

PAGE 149

140 without fully defining the biochemistry of the isolated mutants, however speculation has led to a number of possibilities. One possible reason for these results is that mutagenesis yielded fragile membranes either because of damage to the EII^^^ or because of unspecified secondary q1 c events. In the former case one could hypothesize that the EIP plays a critical structural role; a hypothesis that has some support in light of the data collected on the distribution of EI in mutant vs. wild-type cell extracts (Table 21). The results of these studies indicate that such a structural alteration in the case of these mutants would only be apparent when the membranes are perturbed much like the situation of the ATPase of E^. coli This enzyme was found to change its orientation upon membrane isolation (69). A simpler explanation is the amount of EI may be below saturating levels in these membranes perhaps because of the damq1 c age to the EII^ (as discussed in the previous section). Complementation experiments with isolated PTS components would allow the evaluation of the validity of the latter point. The conclusion that the major lesion in these mutant cells is in the q1 c EIP was substantiated by the failure to observe a transphcsphorylation reaction; a reaction which requires this membrane-bound protein complex only (59), and on the apparently normal levels of EI (and HPr) as shown by direct measurements of total cellular EI (Table 21). This does not rule out a secondary lesion in the EI or HPr proteins which prevents their specific association with the EII^^^ to block PEP-dependent phosphorylation of this sugar. However, given the non-specificity of HPr and EI found in other genera (21), this would seem unlikely. Glc PTS negative mutants were still able to utilize glucose, most likely through a transport mechanism driven by the proton motive force

PAGE 150

141 (18), however, these cells no longer show the typical diauxic growth pattern demonstrated by wild-type cultures growing in glucose plus q1 c lactose (Fig. 16). The localization of the lesion in the EIP is the first step in elucidating a possible mechanism of regulation. In E. coli regulatory functions of the PTS are carried out by the nonspecific components (EI and HPr) and by the high-affinity glc EI 1 1 (see Introduction ). A definitive determination of regulatory functions in S^. mutans would require biochemical fractionation of the wild-type vs. mutant PTS proteins along the lines advanced by Kundig and Roseman (30, 31) in their studies of E. coli It is an approach such as this that allows the definitive determination of the functionality of the individual PTS proteins. In vitro recombination of mutant and wild-type proteins also would aid in the elucidation of which functions are affected by mutagenesis. The similarity of the well-studied regulatory role of the E. coli glc PTS to the S^. mutans glc PTS extends only to the observation that glucose is the preferred sugar in non-induced cells of both species. In ^. coli regulation is complex involving distinct mechanisms (inducer exclusion vs. catabolite repression) and distinct effector molecules (cAMP). Furthermore, regulation presumably involves the PTS functioning to control non-PTS-mediated transport (e.g., glucose vs. lactose); thus, a role for EI and HPr can be ascribed. In S^. mutans regulation of one PTS over a second occurs. It should be noted here that even though the focus of recent studies in E^. coli has been on the PTS control over nonPTS-mediated transport, diauxie has been observed when cells are grown on two PTS substrates as in the cases of glucose plus fructose (21)

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142 and glucose plus sorbitol (34). Very little is known about this control mechanism and its elucidation would probably aid in our understanding of regulation in S. mutans Because EI and HPr function in all PTS-mediated reactions, it is difficult to envision a regulatory role for these two proteins in the case of glc PTS control over other PTS-mediated transport. However, it is plausible that a phospho-EIII^^^ similar to that of E. coli exists in S^. mutans and is able to inhibit heterologous permeases (EII). Thus, in the presence of glucose, phospho-EIII^^^ is formed primarily to transport glucose and secondarily to inhibit other permeases, thereby causing inducer exclusion. Again, it would be informative to determine if a soluble EIII exists before proceeding with further experimentation. If upon fractionation and isolation of the various PTS components such a protein is found, it would then be of interest to add that protein, in a phosphorylated form, to a cell -free system derived from lactose-grown cells to determine if it inhibits lactose phosphorylation. Again, the permeability barrier may have to be overcome. Though I favor the above model, other proposals must be considered. It has been proposed that various phosphorylated metabolites react with the several permeases, thereby causing inducer exclusion. This has been proposed for the observed diauxie in E. coli between glucose and fructose (two PTS substrates) and has been given some support since the discovery of the transphosphorylation reaction (21). A priori one would expect glucose-6-phosphate to be formed both during glucose and lactose metabolism, therefore it is difficult to imagine how this compound or any of its subsequent metabolic products could have a negative regulatory

PAGE 152

143 function in lactose transport. Also, transphosphorylation involves a sugar and its homologous phosphorylated derivative (59,60), therefore the only plausible control mechanism one could postulate is a feed-back inhibition of that specific transport system. An attractive hypothesis has been put forth by Thompson et al (90). They propose that catabolite inhibition is responsible for the preferential utilization of glucose over other PTS sugars. That is, the affinity for HPr by the glucose-specific PTS proteins is greater than by the other sugar-specific PTS proteins (e.g., lactose). If this were the case then a less stringent diauxie than that observed here (Fig, 14) would be expected. Indeed Hamilton and Lo (17) demonstrated that diauxie in the strain of S^. mutans they were studying was concentration dependent. That is, as the glucose concentration was lowered, induction of the lac PTS occurred. This would be an expected observation if competition between two PT-systems exhibiting differential affinities for a limited amount of HPr occurred, as the catabolite inhibition theory proposes. However, the results presented in this dissertation indicate that repression of synthesis rather than competitive inhibition is responsible since the lac PTS is not induced until all the glucose is exhausted. The necessity for the complete absence of glucose is supported by the definite lag period before lac PTS activity; is evident in a glucoserepressed culture. Furthermore, in cells pre-induced for lactose, bath glucose and lactose are utilized simultaneously (Table 22) suggesting no such competition occurs. Catabolite inhibition may explain the results observed when fructose and lactose were present in the growth medium (Fig. 15).

PAGE 153

144 The lac PTS was partially induced in the presence of fructose but did not function until fructose was exhausted; that is, lactose did not start disappearing from the medium until fructose was completely spent. However, since the lac PTS already had been induced, no diauxie occurred. These are exactly the conditions described by Thompson et al. (90) for the glucose preference over galactose in S^. lactis which they explained as being due to catabolite inhibition. Finally, most investigators believe that regulation in streptococci is the result of inducer exclusion (23,24,90), whether it be due to catabolite repression or as yet some non-defined mechanism. Catabolite repression implies the presence of cAMP and as elucidated in the Introduction the evidence in favor of a role for cAMP in Gram-positive organisms is weak.

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147 23. Heller, K. and R. Rbschenthaler, 1978. 6-D-phosphogalactoside galactohydrolase of Streptococcus faecal is and the inhibition of its synthesis of glucose. Can. J. Microbiol. 24:512-519. 24. Heller, K. and R. Rbschenthaler. 1979. The utilization of lactose by Streptococcus faecal is : Effect of glucose and uptake of methyl-3-D-thiogalactoside. FEMS Microbiol. Letts. 5:115-118. 25. Hengstenberg, W. 1977. Enzymology of carbohydrate transport in bacteria. Curr. Top. Microbiol. Immunol. 77:97-126. 26. Hengstenberg, W. W. K. Penberthy, K. L. Hill, and M. L. Morse. 1969. Phosphotransferase system of Staphylococcus aureus : Its requirement for the accumulation and metabolism of galactosides. J. Bacteriol. 99:383-388. 27. Hudig, H. and W. Hengstenberg. 1980. The bacterial phosphoenolpyruvate-dependent phosphotransferase system (PTS). Solubilization and kinetic parameters of the glucose-specific membrane bound enzyme II component of Streptococcus faecal is FEES 114: 103-106. 28. Kaback, H. R. 1974. Transport studies in bacterial membrane vesicles. Science 186:882-892. 29. Kornberg, H. L. and R. E. Reeves. 1972. Inducible phosphoenolpyruvate-dependent hexose phosphotransferase activities in Excherichia coli Biochem. J. 128:1139-1144. 30. Kundig, W. and S. Roseman. 1971. Sugar transport I. Isolation of a phosphotransferase system from Escherichia coli J. Biol. Chem. 246:1393-1406. 31. Kundig, W. and S. Roseman. 1971. Sugar transport II. Characterization of constitutive membrane-bound enzymes II of the Excherichia coli system. J. Biol. Chem. 246:1407-1418. 32. LeBlanc, D. J., V. L. Crow, L. N. Lee, and C. F. Garon. 1979. Influence of the lactose plasmid on the metabolism of galactose by Streptococcus lactis J. Bacteriol. 137:878-884. 33. Lengeler, J. 1979. Streptozotocin an antibiotic superior to penicillin in the selection of rare bacterial mutations. FEMS Microbiil. Lett. 5:417-419. 34. Lengeler, J., and E. C. C. Lin. 1972. Reversal of the mannitolsorbital diauxie in Escherichia coli J. Bacteriol. 112:840-848. 35. Lengeler, J. W. R. J. Mayer, and K. Schmid. 1982. Phosphoenolpyruvate-dependent phosphotransferase system enzyme III and plasmid-encoded sucrose transport in Escherichia coli K12. J. Bacteriol. 151:468-471.

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148 36. Leonard, J. E. and M, H. Saier, Jr. 1981. Genetic dissection of catalytic activities of the Salmonella typhimurium niannitol enzyme II. J. Bacteriol. 145:1106-1109. 37. Lowry, 0. H. N. J. Rosebrough, A. L. Farr, and R. J. Randall. 1951. Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193:265-275. 38. Magasanik, B. 1970. Glucose effects: Inducer exclusion and repression, p. 189-219. In J. Beckwith, and D. Zipser (ed.) The lactose operon. Cold Spring Harbor Laboratories, Cold Spring Harbor, N.Y. 39. Makeman, R. S. and E. W. Sutherland. 1965. Adenosine 3',5'-phase in Escherichia coli J. Biol. Chem. 240:1309-1314. 40. Maryanski J. H. and C. L. Wi ttenberger. 1975. Mannitol transport in Streptococcus mutans J. Bacteriol. 124:1475-1481. 41. McClatchy, J. K. and E. D. Rosenblum. 1963. Induction of lactose utilization in Staphylococcus aureus J. Bacteriol. 86:1211-1215. 42. McKay, L. L. L. A. Walter, W. E. Sandine, and P. R. Elliker. 1969. Involvement of phosphoenolpyruvate in lactose utilization by group N streptococci. J. Bacteriol. 99:603-610. 43. Morse, M. L. K. L. Hill, J. B. Egan, and W. Hengstenberg. 1968. Metabolism of lactose by Staphylococcus aureus and its genetic basis. J. Bacteriol. 95:2270-2274. 44. Park, Y. H. and L. L. McKay. 1982. Distinct galactose phosphoenolpyruvate-dependent phosphotransferase system in Streptococcus lactis J. Bacteriol. 149:420-425. 45. Pastan, I., and S. Adhya. 1976. Cyclic adenosine 3' ,5' -monophosphate in Escherichia coli Bacteriol. Rev. 40:527-551. 46. Peterkofsky, A., and C. Gazdar. 1973. Measurements of rates of adenosine 3': 5' -cyclic monophosphate synthesis in intact Escherichia coli B. Proc. Nat. Acad. Sci. 70:2149-2152. 47. Peterkofsky, A., and C. Gazdar. 1974. Glucose inhibition of adenylate cyclase in intact cells of Escherichia coli B. Proc. Nat. Acad. Sci. 71:2324-2328. 48. Peterkofsky, A., J. E. Gonzalez, and C. Gazdar. 1978. The Escherichia coli adenylate cyclase complex. Regulation by enzyme I of the phosphoenolpyruvate: sugar phosphotransferase system. Arch. Biochem. and Biophys. 188:47-55. 49. Postma, P. W. and J. B. Stock. 1980. Enzymes II of the phosphotransferase system do not catalyze sugar transport in the absence • of phosphorylation. J. Bacteriol. 141:476-484.

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149 50. Raabo, E. and T. C. Terkildsen. 1960. On the enzymatic determination of blood glucose, Scand. J. Clin. Lab. Invest. 12:402-407. 51. Ratliff, T. L. and S. Stinson. 1980. Effect of prostaglandin E-,induced elevation of cyclic AMP on glucose repression in the lactic streptococci. Can. J. Microbiol. 26:58-63. 52. Reizer, J., and C. Panos. 1980. Regulation of 3-galactoside phosphate accumulation in Streptococcus pyogenes by an expulsion mechanism. Proc. Nat. Acad. Sci 77:5497-5501. 53. Rephaeli, A. W. and M. H. Saier, Jr. 1980. Regulation of genes coding for enzyme constituents of the bacterial phosphotransferase system. J. Bacteriol. 141: 658-663. 54. Romano, A. H. S. J. Eberhard, S. L. Dingle, and T. D. McDowell. 1970. Distribution of the phosphoenolpyruvate: glucose phosphotransferase system in bacteria. J. Bacteriol. 104:808-813. 55. Saier, M. H. Jr. 1977. Bacterial phosphoenolpyruvate: sugar phosphotransferase systems: Structural, functional, and evolutionary interrelationships. Bacteriol. Rev. 41:856-871. 56. Saier, M. H. Jr., D. F. Cox, and E. G. Moczydlowski 1977. Sugar phosphate: sugar transphosphorylation coupled to exchange group translocation catalyzed by enzyme II complexes of the phosphoenolpyruvate: sugar phosphotransferase system in membrane vesicles of Escherichia coli J. Biol. Chem. 252:8908-8916. 57. Saier, M. H. Jr., and B. U. Feucht. 1975. Coordinate regulation of adenylate cyclase and carbohydrate permeases by the phosphoenolpyruvate: sugar phosphotransferase system in Salmonell a typhi murium J. Biol. Chem. 250: 7078-7080. 58. Saier, M. H. Jr., and B. U. Feucht. 1980. Regulation of carbohydrate transport activities in Salmonella typhi murium: Use of the phosphoglycerate transport system to energize solute uptake. J. Bacteriol. 141:611-617. 59. Saier, M. H. Jr., B. U. Feucht, and W. K. Mora. 1977. Sugar phosphate: sugar transphosphorylation and exchange group translocation catalyzed by the enzyme II complexes of the bacterial phosphoenolpyruvate: sugar phosphotransferase system. J. Biol. Chem. 252:8899-8907. 60. Saier, M. H. Jr., and E. G. Moczydlowski. 1978. The regulation of carbohydrate transport in Escherichia coli and Salmonella typhimurium p. 103-125. In B. P. RoserTTed.), Bacterial transport. Marcel Dekker, N.Y. 61. Saier, M. H. Jr., M. J. Newman, and A. W. Rephaeli. 1977. Properties of a phosphoenolpyruvate: mannitol phosphotransferase system in Spirocheata aurantia J. Biol. Chem. 252:8890-8898.

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150 62. Saier, M. H. Jr., and S. Roseman. 1976. Sugar transport. The err mutation: Its effect on repression of enzyme synthesis. J. Biol. Chem. 251:6598-6605. 63. Saier, M. H. Jr., and S. Roseman. 1976. Sugar transport. Inducer exclusion and regulation of the melibiose, maltose, glycerol, and lactose transport systems of the phosphoenol pyruvate: sugar phosphotransferase system, J. Biol. Chem. 251:6606-6615. 64. Saier, M. H. Jr., M. R. Schmidt, and Philip Lin. 1980. Phosphoryl exchange reaction catalyzed by enzyme I of the bacterial phosphoenolpyruvate: sugar phosphotransferase system. Kinetic characterization. J. Biol. Chem. 255:8579-8584. 65. Saier, M. H. Jr., and R. D. Simoni. 1976. Regulation of carbohydrate uptake in Gram positive bacteria. J. Biol. Chem. 251: 893-894. 66. Saier, M. H. Jr., R. D. Simoni, and S. Roseman. 1976. Sugar transport: Properties of mutant bacteria defective in proteins of the phosphoenolpyruvate: sugar phosphotransferase system, J. Biol. Chem. 251:6584-6597. 67. Saier, M. H. Jr., H. Straud, J. S. Massman, J. J. Judice, M. J. Newman, and B. U. Feucht. 1978. Pennease-speci fic mutations in Salmonella typhimurium and Escherichia coli that release the glycerol, maltose, melibiose, and lactose transport systems from regulation by the phosphoenolpyruvate: sugar phosphotransferase system. J. Bacteriol. 133:1358-1367. 68. Sakyoun, N. and I. F. Durr. 1972. Evidence against the presence of 3',5'-cyclic adenosine monophosphate and relevant enzymes in Lactobacillus plantarum J. Bacteriol. 112:421-426. 69. Salton, M. R. J., and P. Owen. 1976. Bacterial membrane structure. Ann. Rev. Microbiol. 30:451-482. 70. Schachtele, C. F. 1975. Glucose transport in Streptococcus mutans : Preparation of cytoplasmic membranes and characteristic of phosphotransferase activity. J. Dent. Res. 54: 330-338. 71. Schachtele, C. F. and J. A. Mayo. 1973. Phosphoenol pyruvatedependent glucose transport in oral streptococci. J. Dent. Res. 52:1209-1215. 72. Schmid, K. M. Schupfner, and R. Schmitt. 1982. Plasmid-mediated uptake and metabolism of sucrose by Escherichia coli K-12. J. Bacteriol. 151:68-76. 73. Scholte, B. J. A. R. J. Schiiitema, and P. W. Postma. 1982. Characterization of factor IIl9lc -jp catabolite repressionresistant (err) mutants of Salmonella typhimurium J. Bacteriol. 149: 576-586.

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151 74. Shanmugam, K. T. and R. C. Valentine. 1980. Nitrogen fixation (nif) mutants of Klebsiella pneumoniae In Methods in Enzymology, Vol. 69, p. 47-52. Academic Press, N.Y. 75. Siegel, J. L. S. F. Hurst, E. S. Libennan, S. E. Coleman, and A. S. Bleiweis. 1981. Mutanolysin-induced spheroplasts of Streptotoccus mutans are true protoplasts. Infect. Immun. 31 :808-815. 76. Simoni R. D. J. B. Hays, T. Nakazawa, and S. Roseman. 1973. Sugar transport VI. Phosphoryl transfer in the lactose phosphotransferase system of Staphylococcus aureus J. Biol. Chem. 248:957-965. 77. Simoni, R. D. T. Nakazawa, J. B. Hays, and S. Rosemen. 1973. Sugar transport IV. Isolation and characterization of the lactose phosphotransferase system in Staphylococcus aureus J. Biol. Chem. 248:932-940. 78. Simoni, R. D. and S. Roseman. 1973. Sugar transport VII. Lactose transport in Staphylococcus aureus J. Biol. Chem. 248:946-976. 79. Singer, S. J. and G. L. Nicholson. 1972. The fluid mosaic model structure of cell membranes. Science 175:720-731. 80. Slee, A. M. and J. M. Tanzer. 1979. Phosphoenolpyruvate-dependent sucrose phosphotransferase activity in Streptococcus mutans NCTC 10449. Infect. Immun. 24:821-828. 81. Slee, A. M. and J. M. Tanzer. 1979. Phosphoenolpyruvatedependent sucrose phosphotransferase activity in five serotypes of Streptococcus mutans Infect. Immun. 26:783-786. 82. Slee, A. M. and J. M. Tanzer. 1980. Effect of growth conditions on sucrose phosphotransferase activity of Streptococcus mutans Infect. Immun. 27:922-927. 83. St. Martin, E. J., and C. L. Wi ttenberger. 1979. Characterization of a phosphoenolpyruvate-dependent sucrose phosphotransferase system in Streptococcus mutans Infect. Immun. 24:865-868. 84. St. Martin, E. J., and E. L. Wi ttenberger. 1979. Regulation and function of sucrose-6-phosphate hydrolase in Streptococcus mutans. Infect. Immun. 26:487-491. 85. Terleckyj, B. N. P. Willett, and G. D. Shockman. 1975. Growth of several cariogenic strains of oral streptococci in a chemically defined medium. Infect. Immun. 11:649-655. 86. Thompson, J. 1978. In vivo regulation of glycolysis and characterization of sugar phosphotransferase systems in Streptococcus lactis. J. Bacteriol. 136:465-476.

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151 74. Shanmugam, K. T. and R. C. Valentine. 1980. Nitrogen fixation (nif) mutants of Klebisiella pneumoniae Vol. 69, p. 47-52. In A. San Pietro (ed. ) Methods in Enzymology. Adademic Press, f^w York. 75. Siegel, J. L. S. F. Hurst, E. S. Liberman, S. E. Coleman, and A. S. Bleiweis. 1981. Mutanolysi n-i nduced spheroplasta of Streptococcus mutants are true protoplasts. Infect. Immun. 31:808-815. 76. Simoni R. D. J. B. Hays, T. Nakazawa and S. Roseman. 1973. Sugar transport VI. Phosphoryl transfer in the lactose phosphotransferase system of Staphylococcus aureus J. Biol. Chem. 248:957-965. 77. Simoni, R. D. T. Nakazawa, J. B. Hays, and S. Roseman. 1973. Sugar transport IV. Isolation and characterization of the lactose phosphotransferase system in Staphylococcus aureus J. Biol. Chem. 248:932-940. 78. Simoni, R. D. and S. Roseman. 1973. Sugar transport VII. Lactose transport in Staphylococcus aureus J. Biol. Chem. 248:946-976. 79. Singer, S. J. and G. L. Nicholson. 1972. The fluid mosaic model structure of cell membranes. Science 175:720-731. 80. Slee, A. M. and J. M. Tanzer. 1979. Phosphoenolpyruvate-dependent sucrose phosphotransferase activity in Streptococcus mutans NCTC 10449. Infect. Immun. 24:821-828. 81. Slee, A. M. and J. M. Tanzer. 1979. Phosphoenolpyruvatedependent sucrose phosphotransferase activity in five serotypes of Streptococcus mutans Infect. Immun. 26:783-786. 82. Slee, A. M. and J. M. Tanzer. 1980. Effect of growth conditions on sucrose phosphotransferase activity of Streptococcus mutans Infect. Immun. 27:922-927. 83. St. Martin, E. J., and C. L. Hi ttenberger. 1979. Characterization of a phosphoenolpyruvate-dependent sucrose phosphotransferase system in Streptococcus mutans Infact. Immun. 24:865-868. 84. St. Martin, E. J., and C. L. Wittenberger. 1979. Regulation and function of sucrose-6-phosphate hydrolase in Streptococcus mutans Infect. Immun. 26:487-491. 85. Terleckyj, B. N, P. Willett, and G. D. Shockman. 1975. Growth of several cariogenic strains of oral streptococci in a chemically defined medium. Infect. Immun. 11:649-655. 86. Thompson, J. 1978. In vivo regulation of glycolysis and characterization of sugar phosphotransferase systems in Streptococcus lactis J. Bacteriol. 136:465-476.

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BIOGRAPHICAL SKETCH Ellen S. Liberman was born on July 20, 1950, in Everett, Massachusetts. She was graduated from Everett High School and then attended Northeastern University, Boston, Massachusetts, from which she received the Bachelor of Arts degree in 1973, and the Master of Science degree in 1976. Before pursuing work leading to a Doctor of Philosophy degree, she was employed at Forsyth Dental Center, Boston, Massachusetts, as a research assistant. In 1978, she began her course of study for a Doctor of Philosophy degree in the Department of Microbiology and Cell Science at the University of Florida, Gainesville, Florida. 153

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I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. A. S. Bleiweis, Chairman Professor of Microbiology and Cell Science I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. F. C. Davis Associate Professor of Microbiology and Cell Science I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. R. P. Boyce ^ Professor of Biochemistry and Molecular Biology This dissertation was submitted to the Graduate Faculty of the College of Agriculture and to the Graduate Council, and was accepted as partial fulfillment of the requirements for the degree of Doctor of Philosophy. December 1982 Dean, /Cdllege of Agricult Dean for Graduate Studies and Research


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