Vol. 12, No. 3 Printed in U.S.A.
INJECTION AND IMMUNITY, Sept. 1975, p. 556-563 Copyright © 1975 American Society for Microbiology
Characterization of a Dextranase Produced by an Oral Strain of Actinomyces israelii ROBERT H. STAAT' AND CHARLES F. SCHACHTELE* Microbiology Research Laboratories, School of Dentistry and Department of Microbiology, University of Minnesota, Minneapolis, Minnesota 55455 Received for publication 29 April 1975
A dextranase-producing, gram-positive, anaerobic, rod-shaped bacterium isolated from human dental plaque was identified as Actinomyces israelii. Although the extracellular dextranase (EC 3.2.1.11) formed by this microbe appeared to be constitutively produced, the bacterium did not utilize the reaction products as a carbon source during growth. A striking feature of the dextranase was the formation of two distinct groups of oligosaccharide end products. The two groups presumably correspond to the limit dextran and the released reaction product which appeared to be cleaved from the end(s) of larger dextran molecules. Low levels of dextranase activity were measured by [3H ]NaBH4 reduction and alcohol fixation of the large, tritiated end products on filter paper disks. Of the carbohydrate substrates tested, only a-1,6-linked glucans were cleaved. The enzyme did not exhibit any metal ion requirements, and its pH optimum was 6.3. It is suggested that the A. israelii dextranase may function as a regulatory factor during extracellular in vivo glucan synthesis from sucrose by various plaque microbes.
The unique water-insoluble glucans synthesized from sucrose by the Streptococcus mutans dextransucrase (EC 2.4.1.5) are considered to be responsible for this organism's ability to develop adherent plaques on the tooth surface (16-19, 22, 23). Therefore, it is likely that agents which disrupt the glucans or interfere with their synthesis could reduce the cariogenic potential of S. mutans. The water-insoluble glucans produced by S. mutans are resistant to extensive hydrolysis by dextranases (4, 20, 23, 32, 37). However, if dextranase is present during dextran synthesis from sucrose, the resulting glucans are significantly structurally altered (4, 11, 45; M. D. Dewar and G. J. Walker, Caries Res., in press). These modified glucans reduce the ability of S. mutans to adhere to various surfaces (4, 13, 38) and may suppress the microbe's cariogenic potential in animal models (5, 12). Recently we isolated several distinct groups of dextranase-producing microbes from human dental plaque and suggested that the dextranases released by these microbes could be natural antagonists to the establishment of S. mutans in the oral cavity (38). One group of oral dextranase-producing bacteria was composed of anaerobic gram-positive rods which were tenta'Present address: Department of Community Dentistry, College of Dental Medicine, Medical University of South Carolina, Charleston, S.C. 29401.
556
tively characterized as members of the genus Actinomyces. In the present study, we confirm this classification and describe some of the unusual enzymatic properties of the dextranase produced by this bacterium. MATERIALS AND METHODS Microorganisms. The isolation and preliminary characterization of the bacteria has been previously reported (42). Further taxonomic characterization was performed by the staff of L. V. Holdeman at the Anaerobe Laboratory, Virginia Polytechic Institute, Blacksburg, Va. The procedures used to define the 02 relationship, fermentation capabilities, and fermentation end products analyzed by gas-liquid chromatography have been presented in detail (25). Enzyme preparation. The enzyme was prepared by anaerobically growing the microorganisms in Trypticase soy broth (Difco, Detroit, Mich.) supplemented with 0.5% dextran T40 (Pharmacia, Uppsala, Sweden) and 0.1% yeast extract (Difco) for 48 to 72 h (late log phase) at 37 C in an atmosphere of 10% CO2, 10"/c H2, and 80% N2. The cultures were chilled to 4 C and then centrifuged (8,000 x g, 10 min). The supernatant was concentrated 10 times using an Amicon (Amicon Corp., Lexington, Mass.) ultrafiltration system with PM1O membrane at a pressure of 50 lb/in2. The concentrated preparation was dialyzed in the cold against distilled water for 12 h, followed by dialysis against 0.01 M phosphate buffer (pH 6.0). The crude enzyme preparation was clarified by centrifugation in the cold (10,000 x g, 10 min) and stored at 4 C.
VOL 12, 1975 Dextranase assays. Enzyme activity was measured by monitoring the release of reducing sugar during incubation of the enzyme and substrate at 37 C. Two different techniques were used to quantitate the reducing groups. The first was the method of Somogyi (41) using Nelson's reagent for color development (31). The standard reaction mixture contained either 0.1 or 0.2 ml of enzyme, 0.1 ml of 1.0 M potassium phosphate buffer at pH 6.3, 0.3 ml of dextran T40 (40 mg/ml), and water to a final volume of 1.0 ml. At appropriate intervals, either 0.1- or 0.2-ml samples were removed from the reaction mixture and added to the Somogyi reagent. After color development, the absorbance of the samples was read at 550 nm with a Spectronic 20 (Bausch and Lomb, Rochester, N.Y.). Glucose was used to standardize the colorimetric reaction. One unit of enzyme activity was defined as the amount of enzyme that liberated 1 Ag of glucose equivalent per min in the typical reaction mixture. Since the level of enzyme activity of nonconcentrated preparations obtained from either dextran- or glucose-grown cultures was low (0.2 to 0.3 U/ml) and the reducing power of the residual growth medium was high, it was extremely difficult to accurately quantitate dextran hydrolysis in nonconcentrated preparations by the colorimetric reducing sugar assay. This problem was circumvented with the development of a second reducing sugar assay using [3H]NaBH4 (borohydride). The [3H ]NaBH4 technique has been favored by other researchers as a means to measure reducing sugars because the borohydride specifically reduces aldehydes and ketones. However, in the manufacturing of 3H-labeled borohydride, a significant amount of unidentified acid-stable material is generated (30). The percentage of the contaminant is usually high enough to necessitate further purification of the reduced carbohydrates before accurate data are obtained (7, 35). To overcome the borohydride contamination problem, we used an unusual feature of the Actinomyces dextranase. The dextranase degrades dextrans to saccharides that are 10 to 18 glucose units in length (42). Germaine et al. (15) showed that dextrans larger than 7 glucose units are quantitatively bound to filter paper disks when washed with methanol. NaBH4 is slightly soluble in methanol with increased solubility in acetone. Using this information, the following assay for reducing glucans larger than 7 glucose units was devised: 50 gl of the standard enzyme reaction mix was added to 50 Ml of 0.2 N NaBH4 in 0.1 N NaOH supplemented with 106 counts/min [3H]NaBH4. The reaction was incubated at 37 C for 45 min followed by addition of 100 gl of 2 N HCl. A 25-Al aliquot was then applied to 24-mm Whatman 3MM filter paper disks (Reeve Angel, Clifton, N.J.), and the disks were washed in methanol or acetone (10 ml/disk) for 15 min. The washing was repeated once with fresh solvent. The disks were then air dried, placed in vials containing scintillation fluid (4.0 g of 2,5-diphenyloxazole; 0.1 g
A. ISRAELII DEXTRANASE
557
Downers Grove, Ill.). Protein was measured according to the procedures of Lowry et al. using bovine serum albumin as a standard (29). Time course of enzyme production. The organisms were grown in 50-ml aliquots of glucose-free Trypticase soy broth supplemented with 0.1% yeast extract and either 1% glucose or 1% dextran T40. The Erlenmeyer flasks containing uninoculated medium were septum sealed and preincubated under anaerobic conditions for 24 h with a syringe needle (18 gauge)
inserted through the septum to facilitate gas exchange. The flasks were inoculated with 5 ml of a culture actively growing on the same carbon source. and the needle was removed to seal the system. The cultures were agitated at a rate of 100 cycles/min in a 37 C water bath. Samples (1 ml) were removed at intervals with a syringe and needle. Growth was estimated turbidimetrically at a wavelength of 600 nm. The cells were removed by centrifugation (10,000 x g, 15 min), and 0.5 ml of the supernatant was used in the standard dextranase assay. Reducing sugar was quantitated using the ['H ]NaBH4 assay. Analysis of dextranase reaction products. The products of hydrolysis of dextran by the Actinomyces dextranase were analyzed by chromatography using Bio Gel P10 (Bio-Rad Laboratories, Richmond, Calif.) in a 1.5- by 25-cm column. A 1.0-ml sample of the standard reaction mixture was placed on a column which had been prewashed with 0.1 M sodium acetate (pH 4.5). Carbohydrates were eluted with the same buffer, and the carbohydrate content of each fraction was determined using the phenolsulfuric acid assay (10).
RESULTS Taxonomic characteristics. In a previous report (42), we tentatively placed a dextranaseproducing, anaerobic, gram-positive, rodshaped bacterium from dental plaque in the genus Actinomyces. At that time, it was noted that our preliminary data did not specifically delineate between the genus Bifidobacterium and the genus Actinomyces. One of the major differences between these two genera is the ability of the Bifidobacterium to produce significant amounts of acetic acid, resulting in a ratio of acetic acid to lactic acid of between 2 and 3 to 1 (8, 9, 36). Gas-liquid chromatographic analysis of the end products released during growth of the organisms in a glucose broth indicated that the microbe produced very little acetic acid (Table 1). This confirmed our initial classification of these organisms as Actinomyces spp. The macrocolonies of the dextranase-producing Actinomyces strain Gl are cream-white in color and are similar to the "molar tooth" or "raspberry-like" colonies (40) usually associated with A. israelii (Fig. la). The dextranActinomyces strains G2 and G3 hydrolyzing of of p-bis[2-(5-phenyloxazolyl)lbenzene; 1.0 liter round colonies with a slight convex, form soft, toluene) and counted with a Packard 2425 liquid scintillation spectrometer (Packard Instruments Inc., granular texture to the surface of the white
558
STAAT AND SCHACHTELE
INFECT. IMMUN.
starch supported weak fermentation, whereas arabinose and xylose were not fermented. The organisms failed to hydrolyze starch agar medium. These microbes grew anaerobically in the presence of CO2; however, no growth on Trypticase soy blood agar has been detected for any of the isolates under aerobic conditions. On the basis of the organism's ability to ferment ribose to acid and on its inability to produce observable growth on Trypticase soyTABLE 1. Distribution of fermentation products blood agar when grown aerobically (14, 40, 44), resulting from eInaerobic growth of Actinomyces we suggest that these dextranase-producing strains Gl, G2, and G3 in peptone-yeast microbes be considered strains of A. israelii. extract-glucose broth Enzyme characterization: evaluation of the [8H]NaBH4 dextranase assay. Figure 2 shows % of total acid produceda Acid that incorporation of borohydride derived 3H Gi G2 G3 into nonhydrolyzed dextran T40 is linear with respect to dextran concentration. The borohy9.3 4.2 6.2 Acetic ......... dride reaction is time and temperature dependtb 7.5 Formic ........ 0 ent and often requires extended incubation for 0 0 t Propionic ...... completion (30). Reacting dextran T40 or Tio 0 0 0.5 Butyric ........ 92.6 90.6 Lactic ......... 62.6 with [3H]NaBH4 at 37 C for various time pe0 1.8 Pyruvic ........ 6.5 riods indicated that reduction of these dextrans 3.2 Succinic ....... 14.0 2.7 neared completion after 30 min. Therefore, aMeq of individual acid/meq of total acid x 100 as incubation for 45 min at 37 C was used for the borohydride-reducing dextran assay. Figure 3 determined by gas-liquid chromatography. illustrates the incorporation of 3H into oligosact, Trace.
colonies (Fig. lb), similar to the smooth colonies of both A. israelii and Actinomyces naeslundii (44). Results of the biochemical reactions for species characterization were identical for the three dextranase-producing strains, except for strain G3 which could not hydrolyze esculin. Data from cultures grown in medium supplemented with various carbohydrates indicated that ribose was fermented to acid and soluble
FIG. 1. Colonial morphologies of dextranase-producing Actinomyces; (a) strain Gl, (b) strain G2. Strain G3 is similar to G2.
A. ISRAELII DEXTRANASE
VOL 12, 1975
have expected had the organism catabolized the dextran (42). This suggested that the supplemental dextran did not contribute to the growth of the bacterium. This was confirmed by growing cells in medium devoid of supplemental carbohydrate. The growth curve produced in this experiment was essentially identical to the one in Fig. 4 (data not shown). A. israelii utilizes glucose for growth, and dextranase activity was detectable in the supernatant of glucose-grown cells. Figure 5 illustrates that cells grown in glucose broth produced dextranase during the logarithmic growth phase. Although the total growth and dextranase production was greater for glucose-grown cultures compared to cultures supplemented only with dextran, the shape of the growth curves and the specific enzyme activities were similar in both experiments. To date, no growth conditions have been found which suppress dextranase production by A. israelii strains G1-3.
8 7
6 C
5
0
7t
0.
g)
559
4 3
2
0
250 200 150 pg Dextran T40 FIG. 2. Incorporation of sodium borohydridederived 3H into the reducing ends of dextran T40. Dextran T40 was added to 50 ;,1 of 0.2 N [3HJNaBH4 (106 counts/min) in 0.1 N NaOH and incubated at 37 C for I h. 50
100
6 5
charides cleaved from dextran T40 by the A. I) 4 israelii dextranases (open circles) compared to heat-inactivated enzyme (closed circles). This 0 curve is similar to that obtained with the colorimetric reducing sugar assay using the same enzyme (42). Validation that the [3H]NaNH4 was in sufficient excess for complete carbohydrate reduction was obtained by incubating 1% glucose in the system used to show linearity of the 3H incorporation into dextran T,4, The glucose did not affect the amount of 3H incorporated into the dextran T40 nor was the slope of the line altered (data not shown). Conditions for enzyme production. The effect of different carbohydrate supplements on growth and dextranase production by A. israelii was studied. The growth curve of strain Gl with dextran T40 as the carbohydrate supplement is shown in Fig. 4 (open circles). Dextranase ac60 40 20 0 tivity (closed circles) was greatest as the culture shifted from the logarithmic to the stationary Time phase of growth. The rapid inactivation of the enzyme activity is presumed to be caused by FIG. 3. Incorporation of sodium borohydrideproteolytic activity in the post-log culture derived 3H into the glucans released from dextran T740 by the A. israelii Gl dextranase. Symbols: 0, active supernatant. It was noted previously that A. israelii did not enzyme; 0, heat-inactivated (95 C for 10 min) enferment dextran to an acid pH as one would zyme.
12
(min)
560
INFECT. IMMUN.
STAAT AND SCHACHTELE
.8
6K2 c
_2
E
-2
->O\
4
0
Substrate specificity. The ability of A. israelii Gl dextranase to hydrolyze various carbohydrates was determined, and the results are presented in Table 3. The predominantly a-1,6linked polysaccharide, dextran, was the only substrate hydrolyzed. End product analysis. Complete hydrolysis of dextran T40 by the A. israelii Gl dextranase
L, 0
2
4
-_i6 __-.8 rime
10
0.4
o 2
24
hours)
FIG. 4. Growth and enzyme production curves for A. israelii GJ grown anaerobically with dextran T40 as the supplemental carbon source at a 1% concentration. Symbols: 0, optical density (turbidity) of culture at 600 nm; 0, dextranase activity assayed with the [3H]NaBH4 procedure.
0 QI)
0.3
0.2
0
-o L.. 0 'I,
2.0 z
>
-o
0.1
E
_\X
8.0
'D c
0
_:
D
T
_
.D
pH
Ir
ci 4,I -r
FIG. 6. pH optimum of the crude A. israelii GI dextranase. Dextranase activity was determined in the standard dextranase reaction (0.1 ml of enzyme) as the increased reducing sugar released after 45 min. Buffers: pH 4.0 to 5.5, sodium citrate; pH 6.0 to 7.5,
potassium phosphate. 4
6
8
10
12
24
Time (hours)
FIG. 5. Growth and enzyme production curves for A. israelii GI grown anaerobically with glucose as the supplemental carbon source at a 1% concentration. Symbols: 0, optical density (turbidity) of culture at 600 nm; 0, dextranase activity assayed with the ['H]NaBH4 procedure.
pH optimum. The pH optimum for the A. israelii dextranase centers around 6.3 (Fig. 6). The activity was relatively stable over the pH range 5.6 to 7.0. Inhibition studies. Among the metal ions tested, Ag+ and Hg2+ at 1 mM were the only inhibitors of the A. israelii dextranase (Table 2). Additonally, no metal ions caused activation of the dextranase nor did the metal chelator ethylenediaminetetraacetic acid cause any reduction in dextranase activity. The sulfhydryl reagents, dithiothreitol and iodoacetamide, had no effect at 10 mM concentrations, whereas the anionic detergent, sodium dodecyl sulfate, showed complete inhibition of the enzyme.
TABLE 2. Effect of some metal salts and other chemicals on the A. israelii Gl dextranase Chemicala
Concn (mM)
None
CaCl2 AgNO3
1 1
HgCl2 MnCl2
1 1
Relative 100 105 0 0 101
MgCl2
1
95
ZnSO4 EDTA SDS DTT IA
1
100
10 10 10 10
103 0 103 100
a EDTA, Ethylenediaminetetraacetic acid; SDS, sodium dodecyl sulfate; DTT, dithiothreitol; IA, iodoacetamide. 'The chemical was added to the standard reaction mixture at to, and 0.2-ml aliquots were taken at t20 and t40 and assayed for reducing power with the [3H JNaBH4 procedure. Relative activity = (t40 -
t20)chem/(t40 - t20)none x 100.
A. ISRAELII DEXTRANASE
VOL 12, 1975
resulted in the formation of oligosaccharides containing approximately 10 to 18 glucose units (42). In an attempt to visualize the dextran degradation, typical dextranase reaction mixtures were chromatographed at various stages of hydrolysis on Bio Gel P10 (Fig. 7). After 1 h, a small amount of apparent end product appeared in the area of fraction 38. At the 2-h mark, significant amounts of end product had appeared, and, essentially, none of the original dextran remained intact. Also, after 2 h, the polysaccharides were distributed into two distinct molecular weight ranges presumably corresponding to degradation intermediates and product. The 6-h pattern illustrates the continued movement of the intermediate peak to the right. The double-peaked pattern seen at 24 h remained unchanged through 48 h of incubation. The position of the double peak pattern corresponds to dextrans of approximately 10 to 18 glucose units in size (42).
DISCUSSION Dextranases produced by the oral strains of A. israelii isolated in our laboratories are extracellular endohydrolytic enzymes. Other bacteria capable of producing extracellular dextranases belong to the genera Pseudomonas (34), Bifidobacterium (1, 2, 6), Bacteriodes (24, 39), and Cellvibrio (28). Phylogenetically, the Actinomyces are closely related to the Bifidobacterium (orginally thought to be Lactobacillus [33]); however, comparison of the dextranases produced by these organisms reveals substantial differences. Contrasting the physiological characteristics of the two microbes, one can note the failure of A. israelii to ferment or apparently utilize dextrans in any way during growth, whereas the Bifidobacterium effectively utilize dextrans as the sole carbon source (6). Additionally, the A. israelii dextranase appears TABLE 3. Substrate specificity of concentrated crude A. israelii Gl dextranase
(Cogcn)
Glucan
Dextran T4, ... Amylose Glycogen ..... Pseudonigeran Laminaran ... Isomaltose
Linkage
Activitya
12 6
a-1,6; a-1,3 a-1,4
+
6 4
a-1,4; a-1,6
-
561
E
I
Frocticm I-"ai* FIG. 7. Bio Gel P-10 column chromatography of the reaction products produced by A. israelii Gl dextranase at various stages of dextran T40 hydrolysis.
a constitutive enzyme, whereas the Bifidobacterium dextranase is an inducible enzyme (1, 6). Comparative enzymology of the dextranases from the two different genera indicates that I both retain activity over the same broad pH was noted after 90 min of incubation of the standard range (5.5 to 7.0), and both appear to hydrolyze reaction system compared to boiled (95 C, 10 min) only linear a-1,6-linked glucans. The Bifidobacterium dextranase hydrolyzes dextrans to oligoenzyme preparations.
a-1,3 6 ,-1,3 0.6 a-1,6 Activity was considered positive (+) if an increase of 10% in the reducing sugar assay (Nelson-Somogyi) ....
to be
562
INFECT. IMMUN.
STAAT AND SCHACHTELE
saccharides 3 to 5 glucose units in length (2, 3), whereas the A. israelii dextranase apparently hydrolyzes dextran T40 into at least two types of oligosaccharides 10 to 18 glucose units in size. The progressive development of the doublepeaked, end product curve by the A. israelii dextranase (Fig. 7) can be explained as a nonrandom enzymatic recognition of the end of the dextran molecule with subsequent release of oligosaccharides of relatively specific length from the dextran. This could imply that the larger end product oligosaccharides are limit dextrans and that a dextran molecule of possibly 20 glucose units is needed before hydrolysis can be initiated by the A. israelii dextranase. The physiological significance of the A. israelii dextranase is not understood at the present time, and the production of dextranase by this microbe is inconsistent with the overview of extracellular microbial hydrolase function. The dextranase apparently does not contribute to the accumulation of metabolic requirements for the cells, since growth and dextranase production were equivalent in the presence or absence of the substrate (Fig. 4). Additionally, glucose did not suppress dextranase production by A. israelii (Fig. 5). These data can be explained by suggesting that an as yet unidentified molecule is the primary substrate for the enzyme and that cleavage of large oligosaccharides from dextrans is a fortuitous side reaction. An alternate explanation is that dextranase production by A. israelii constitutes a contribution to a symbiotic relationship established in the complex milieu of dental plaque. A. israelii is often recovered from the plaque of healthy individuals (26, 27, 40), and production of dextranase by selected strains may enhance the microbes' ability to maintain itself in plaque through association with- the dextran-forming microorganisms. In preliminary experiments, we recovered dextranase-producing microbes resembling A. israelii about 60% of the time from the plaque of elementrary school children (R. Staat and C. Schachtele, unpublished data). The effects of indigenous dextranase activities on the structure or composition of dental plaque can manifest themselves in several ways. Oral microbes such as Bacteroides ochraceus induce dextranases in response to the presence of the substrate and ferment the released glucose to acid (R. Staat and C. Schachtele, submitted for publication). The net result of this activity would be a reduced dextran pool in plaque. However, organisms such as A. israelii arid S. mutans produce dextranases, but do not metabolize the reaction products to acid; thus,
they apparently do not by themselves decrease the amount of dextran in plaque. This is not to suggest that the dextranases do not contribute to the metabolism of dental plaque. Walker proposed that oligosaccharides produced by endohydrolytic dextranase activity acted as alternate acceptor molecules in the S. mutans dextransucrase reaction (45). This concept was extended to account for the possible function of the S. mutans dextranase as a regulatory mechanism for the production of highly branched, dextranase-resistant glucans by that organism's dextransucrase (21, 43). Recently, our laboratory presented data indicating that the endohydrolytic dextranase produced by A. israelii reduces both the production of water-insoluble glucans by S. mutans 6715 dextransucrase and the adherence of whole organisms to glass surfaces (38). It should be noted that the A. israelii dextranase does not reduce the rate of total polysaccharide synthesis by S. mutans 6715 dextransucrase (R. Staat, unpublished data). Therefore, it can be proposed that the ability of S. mutans to establish as a member of the plaque flora is regulated in part by the qualitative structure of the glucan rather than the quantity of glucan produced. If this proposal is valid, then extracellular dextranases produced by the indigenous plaque flora have the potential for controlling the establishment of S. mutans by structural modifications of glucans and that elimination of the glucans from dental plaque need not be a prerequisite for ecological control of this bacterium. ACKNOWLEDGMENTS We thank L. V. Holdeman and her staff at the Virginia Polytechnic Institute Anaerobe Laboratory for the taxonomic data. The technical assistance of K. Cummings and WoonLam S. Leung is acknowledged. This work was supported by Public Health Service grant DE 02654 from the National Institute of Dental Research. R.H.S. was the recipient of National Institute of Dental Research Individual Research Fellowship Award 1 F22 DE00709. C.F.S. was the recipient of Public Health Service Career Development Award K4-DE-42,859.
LITERATURE CITED 1. Bailey, R. W., and R. T. J. Clarke. 1959. A bacterial dextranase. Biochem. J. 72:49-54. 2. Bailey, R. W., D. H. Hutson, and H. Weigel. 1960. Action of bacterial dextranase on branched dextrans. Nature
(London) 186:553-554. 3. Bailey, R. W., D. H. Hutson, and H. Weigel. 1961. The action of a Lactobbacillus bifidus dextranase on a branched dextran. Biochem. J. 80:514-519. 4. Bowen, W. H. 1968. Effects of dextranase on cariogenic and noncariogenic dextrans. Br. Dent. J. 124:347-349. 5. Bowen, W. H. 1971. The effect of dextranase on caries activity in monkeys (Macaca irus). Br. Dent. J. 131:445-449. 6. Clarke, R. 'F. 1. 1959. A dextran-fermenting organism
VOL 12, 1975 from the rumen closely resembling Lactobacillus bifidus. J. Gen. Microbiol. 20:549-553. 7. Conrad, H. E., E. Varboncouer, and M. E. James. 1973. Qualitative and quantitative analysis of reducing carbohydrates by radiochromatography on ion-exchange papers. Anal. Chem. 51:486-500. 8. de Vries, W., S. J. Gerbrandy, and A. H. Stouthamer. 1967. Carbohydrate metabolism in Bifidobacterium bifidum. Biochim. Biophys. Acta 136:415-425. 9. de Vries, W., and A. H. Stouthamer. 1968. Fermentation of glucose, lactose, galactose, mannitol, and xylose by bifidobacteria. J. Bacteriol. 96:472-478. 10. Dubois, M., K. A. Gilles, J. K. Hamilton, P. A. Rebers, and F. Smith. 1956. Colorimetric method for the determination of sugars and related substances. Anal. Chem. 28:350-356. 11. Ebisu, S., A. Misaki, K. Kato, and S. Kotani. 1974. The structure of water-insoluble glucans of cariogenic Streptococcus mutans, formed in the absence and presence of dextranase. Carbohydr. Res. 38:374-381. 12. Fitzgerald, R. J., P. H. Keyes, T. H. Stoudt, and D. Spinell. 1968. The effects of a dextranase preparation on plaque and caries in hamsters, a preliminary report. J. Am. Dent. Assoc. 76:301-304. 13. Fitzgerald, R. J., D. M. Spinell, and T. H. Stoudt. 1968. Enzymatic removal of artificial plaques. Arch. Oral Biol. 13:125-128. 14. Georg, L. K., G. W. Roberstad, and S. A. Brinkman. 1964. Identification of species of Actinomyces. J. Bacteriol. 88:477-490. 15. Germaine, G. R., C. F. Schachtele, and A. M. Chludzinski. 1974. Rapid filter paper assay for the dextransucrase activity from Streptococcus mutans. J. Dent. Res. 53:1355-1360. 16. Gibbons, R. J. 1968. Formation and significance of bacterial polysaccharides in caries etiology. Caries Res. 2:164-171. 17. Gibbons, R. J., and S. B. Banghart. 1967. Synthesis of extracellular dextran by cariogenic bacteria and its presence in human dental plaque. Arch. Oral Biol. 12:11-24. 18. Gibbons, R. J., and R. J. Fitzgerald. 1969. Dextraninduced agglutination of Streptococcus mutans and its potential role in the formation of microbial dental plaques. J. Bacteriol. 98:341-346. 19. Gibbons, R. J., and M. Nygaard. 1968. Synthesis of insoluble dextran and its significance in the formation of gelatinous deposits by plaque-forming streptococci. Arch. Oral Biol. 13:1249-1262. 20. Guggenheim, B. 1970. Enzymatic hydrolysis and structure of water-insoluble glucan produced by glycosyltransferases from a strain of Streptococcus mutans. Helv. Odontol. Acta 5 (Suppl. 14):89-108. 21. Guggenheim, B., and J. J. Burckhardt. 1974. Isolation and properties of a dextranase from Streptococcus mutans OMZ 176. Helv. Odontol. Acta 18:101-113. 22. Guggenheim, B., and E. Newbrun. 1969. Extracellular glucosyltransferase activity of an HS strain of Streptococcus mutans. Helv. Odontol. Acta 13:84-97. 23. Guggenheim, B., and H. E. Schroeder. 1968. Biochemical and morphological aspects of extracellular polysaccharides produced by cariogenic streptococci. Helv. Odont. Acta 11:131-152. 24. Hehre, E. J., and T. W. Sery. 1952. Dextran-splitting anaerobic bacteria from the human intestine. J. Bacteriol. 63:424-426. 25. Holdeman, L. V., and W. E. C. Moore. 1973. Anaerobe laboratory manual, 2nd ed. Virginia Polytechnic Insti-
A. ISRAELII DEXTRANASE
563
tute and State University, Blacksburg. 26. Howell, A., W. C. Murphy, F. Paul, and R. M. Stephan. 1959. Oral strains of Actinomyces. J. Bacteriol. 78:82-95. 27. Howell, A., R. M. Stephan, and F. Paul. 1962. Prevalence of Actinomyces israelii, A. naeslundii, Bacterionema matruchotti and Candida albicans in selected areas of the oral cavity and saliva. J. Dent. Res. 41:1050-1059. 28. Ingelman, G. 1948. Enzymatic breakdown of dextran. Acta Chem. Scand. 2:803-812. 29. 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. 30. McLean, C., D. A. Werner, and D. Aminoff. 1973. Quantitative determination of reducing sugars, oligosaccharides, and glycoproteins with ('H) borohydride. Anal. Biochem. 55:72-84. 31. Nelson, N. 1944. A photometric adaptation of the Somogyi method for the determination of glucose. J. Biol. Chem. 153:375-380. 32. Newbrun, E. 1972. Extracellular polysaccharides synthesized by glucosyltransferases of oral streptococci. Caries Res. 6:132-147. 33. Poupard, J. A., I. Husain, and R. F. Norris. 1973. Biology of the bifidobacteria. Bacteriol. Rev. 37:136-165. 34. Richards, G. N., and M. Streamer. 1972. Studies on dextranases. I. Isolation of extracellular, bacterial dextranases. Carbohydr. Res. 25:323-332. 35. Richards, G. N., and W. J. Whelan. 1973. Determination of the number-average molecular weight of polysaccharides by end-group reduction with borohydride-t. Carbohydr. Res. 27:185-191. 36. Scardovi, V., and L. D. Trovatelli. 1965. The fructose6-phosphate shunt as a peculiar pattern of hexose degradation in the genus Bifidobacterium. Ann. Microbiol. Enzymol: 15:19-29. 37. Schachtele, C. F., A. E. Loken, and M. K. Schmitt. 1972. Use of specifically labeled sucrose for comparison of extracellular glucan and fructan metabolism by oral streptococci. Infect. Immun. 5:263-266. 38. Schachtele, C. F., R. H. Staat, and S. K. Harlander. 1975. Dextranases from oral bacteria: inhibition of water-insoluble glucan production and adherence to smooth surfaces by Streptococcus mutans. Infect. Immun. 12:309-317. 39. Sery, T. W., and E. J. Hehre. 1956. Degradation of dextrans by enzymes of intestinal bacteria. J. Bacteriol. 71:373-380. 40. Slack, J. M. 1974. Actinomyces, p. 660-667. in R. E. Buchanan and N. E. Gibbons. (ed.), Bergey's manual of determinative bacteriology, 8th ed. The Williams & Wilkins Co., Baltimore. 41. Somogyi, M. 1945. A new reagent for the determination of sugars. J. Biol. Chem. 160:61-68. 42. Staat, R. H., T. H. Gawronski, and C. F. Schachtele. 1973. Detection and preliminary studies on dextranaseproducing microorganisms from human dental plaque. Infect. Immun. 8:1009-1016. 43. Staat, R. H., and C. F. Schachtele. 1974. Evaluation of dextranase production by the cariogenic bacterium Streptococcus mutans. Infect. Immun. 9:467-469. 44. Thompson, L., and S. A. Lovestedt. 1951. An Actinomyces-like organism obtained from the human mouth. Proc. Staff Meet. Mayo Clin. 26:169-175. 45. Walker, G. J. 1972. Some properties of a dextranglucosidase isolated from streptococci and its use in studies on dextran synthesis. J. Dent. Res. 51:409-414.
INJECTION AND IMMUNITY, Sept. 1975, p. 556-563 Copyright © 1975 American Society for Microbiology
Characterization of a Dextranase Produced by an Oral Strain of Actinomyces israelii ROBERT H. STAAT' AND CHARLES F. SCHACHTELE* Microbiology Research Laboratories, School of Dentistry and Department of Microbiology, University of Minnesota, Minneapolis, Minnesota 55455 Received for publication 29 April 1975
A dextranase-producing, gram-positive, anaerobic, rod-shaped bacterium isolated from human dental plaque was identified as Actinomyces israelii. Although the extracellular dextranase (EC 3.2.1.11) formed by this microbe appeared to be constitutively produced, the bacterium did not utilize the reaction products as a carbon source during growth. A striking feature of the dextranase was the formation of two distinct groups of oligosaccharide end products. The two groups presumably correspond to the limit dextran and the released reaction product which appeared to be cleaved from the end(s) of larger dextran molecules. Low levels of dextranase activity were measured by [3H ]NaBH4 reduction and alcohol fixation of the large, tritiated end products on filter paper disks. Of the carbohydrate substrates tested, only a-1,6-linked glucans were cleaved. The enzyme did not exhibit any metal ion requirements, and its pH optimum was 6.3. It is suggested that the A. israelii dextranase may function as a regulatory factor during extracellular in vivo glucan synthesis from sucrose by various plaque microbes.
The unique water-insoluble glucans synthesized from sucrose by the Streptococcus mutans dextransucrase (EC 2.4.1.5) are considered to be responsible for this organism's ability to develop adherent plaques on the tooth surface (16-19, 22, 23). Therefore, it is likely that agents which disrupt the glucans or interfere with their synthesis could reduce the cariogenic potential of S. mutans. The water-insoluble glucans produced by S. mutans are resistant to extensive hydrolysis by dextranases (4, 20, 23, 32, 37). However, if dextranase is present during dextran synthesis from sucrose, the resulting glucans are significantly structurally altered (4, 11, 45; M. D. Dewar and G. J. Walker, Caries Res., in press). These modified glucans reduce the ability of S. mutans to adhere to various surfaces (4, 13, 38) and may suppress the microbe's cariogenic potential in animal models (5, 12). Recently we isolated several distinct groups of dextranase-producing microbes from human dental plaque and suggested that the dextranases released by these microbes could be natural antagonists to the establishment of S. mutans in the oral cavity (38). One group of oral dextranase-producing bacteria was composed of anaerobic gram-positive rods which were tenta'Present address: Department of Community Dentistry, College of Dental Medicine, Medical University of South Carolina, Charleston, S.C. 29401.
556
tively characterized as members of the genus Actinomyces. In the present study, we confirm this classification and describe some of the unusual enzymatic properties of the dextranase produced by this bacterium. MATERIALS AND METHODS Microorganisms. The isolation and preliminary characterization of the bacteria has been previously reported (42). Further taxonomic characterization was performed by the staff of L. V. Holdeman at the Anaerobe Laboratory, Virginia Polytechic Institute, Blacksburg, Va. The procedures used to define the 02 relationship, fermentation capabilities, and fermentation end products analyzed by gas-liquid chromatography have been presented in detail (25). Enzyme preparation. The enzyme was prepared by anaerobically growing the microorganisms in Trypticase soy broth (Difco, Detroit, Mich.) supplemented with 0.5% dextran T40 (Pharmacia, Uppsala, Sweden) and 0.1% yeast extract (Difco) for 48 to 72 h (late log phase) at 37 C in an atmosphere of 10% CO2, 10"/c H2, and 80% N2. The cultures were chilled to 4 C and then centrifuged (8,000 x g, 10 min). The supernatant was concentrated 10 times using an Amicon (Amicon Corp., Lexington, Mass.) ultrafiltration system with PM1O membrane at a pressure of 50 lb/in2. The concentrated preparation was dialyzed in the cold against distilled water for 12 h, followed by dialysis against 0.01 M phosphate buffer (pH 6.0). The crude enzyme preparation was clarified by centrifugation in the cold (10,000 x g, 10 min) and stored at 4 C.
VOL 12, 1975 Dextranase assays. Enzyme activity was measured by monitoring the release of reducing sugar during incubation of the enzyme and substrate at 37 C. Two different techniques were used to quantitate the reducing groups. The first was the method of Somogyi (41) using Nelson's reagent for color development (31). The standard reaction mixture contained either 0.1 or 0.2 ml of enzyme, 0.1 ml of 1.0 M potassium phosphate buffer at pH 6.3, 0.3 ml of dextran T40 (40 mg/ml), and water to a final volume of 1.0 ml. At appropriate intervals, either 0.1- or 0.2-ml samples were removed from the reaction mixture and added to the Somogyi reagent. After color development, the absorbance of the samples was read at 550 nm with a Spectronic 20 (Bausch and Lomb, Rochester, N.Y.). Glucose was used to standardize the colorimetric reaction. One unit of enzyme activity was defined as the amount of enzyme that liberated 1 Ag of glucose equivalent per min in the typical reaction mixture. Since the level of enzyme activity of nonconcentrated preparations obtained from either dextran- or glucose-grown cultures was low (0.2 to 0.3 U/ml) and the reducing power of the residual growth medium was high, it was extremely difficult to accurately quantitate dextran hydrolysis in nonconcentrated preparations by the colorimetric reducing sugar assay. This problem was circumvented with the development of a second reducing sugar assay using [3H]NaBH4 (borohydride). The [3H ]NaBH4 technique has been favored by other researchers as a means to measure reducing sugars because the borohydride specifically reduces aldehydes and ketones. However, in the manufacturing of 3H-labeled borohydride, a significant amount of unidentified acid-stable material is generated (30). The percentage of the contaminant is usually high enough to necessitate further purification of the reduced carbohydrates before accurate data are obtained (7, 35). To overcome the borohydride contamination problem, we used an unusual feature of the Actinomyces dextranase. The dextranase degrades dextrans to saccharides that are 10 to 18 glucose units in length (42). Germaine et al. (15) showed that dextrans larger than 7 glucose units are quantitatively bound to filter paper disks when washed with methanol. NaBH4 is slightly soluble in methanol with increased solubility in acetone. Using this information, the following assay for reducing glucans larger than 7 glucose units was devised: 50 gl of the standard enzyme reaction mix was added to 50 Ml of 0.2 N NaBH4 in 0.1 N NaOH supplemented with 106 counts/min [3H]NaBH4. The reaction was incubated at 37 C for 45 min followed by addition of 100 gl of 2 N HCl. A 25-Al aliquot was then applied to 24-mm Whatman 3MM filter paper disks (Reeve Angel, Clifton, N.J.), and the disks were washed in methanol or acetone (10 ml/disk) for 15 min. The washing was repeated once with fresh solvent. The disks were then air dried, placed in vials containing scintillation fluid (4.0 g of 2,5-diphenyloxazole; 0.1 g
A. ISRAELII DEXTRANASE
557
Downers Grove, Ill.). Protein was measured according to the procedures of Lowry et al. using bovine serum albumin as a standard (29). Time course of enzyme production. The organisms were grown in 50-ml aliquots of glucose-free Trypticase soy broth supplemented with 0.1% yeast extract and either 1% glucose or 1% dextran T40. The Erlenmeyer flasks containing uninoculated medium were septum sealed and preincubated under anaerobic conditions for 24 h with a syringe needle (18 gauge)
inserted through the septum to facilitate gas exchange. The flasks were inoculated with 5 ml of a culture actively growing on the same carbon source. and the needle was removed to seal the system. The cultures were agitated at a rate of 100 cycles/min in a 37 C water bath. Samples (1 ml) were removed at intervals with a syringe and needle. Growth was estimated turbidimetrically at a wavelength of 600 nm. The cells were removed by centrifugation (10,000 x g, 15 min), and 0.5 ml of the supernatant was used in the standard dextranase assay. Reducing sugar was quantitated using the ['H ]NaBH4 assay. Analysis of dextranase reaction products. The products of hydrolysis of dextran by the Actinomyces dextranase were analyzed by chromatography using Bio Gel P10 (Bio-Rad Laboratories, Richmond, Calif.) in a 1.5- by 25-cm column. A 1.0-ml sample of the standard reaction mixture was placed on a column which had been prewashed with 0.1 M sodium acetate (pH 4.5). Carbohydrates were eluted with the same buffer, and the carbohydrate content of each fraction was determined using the phenolsulfuric acid assay (10).
RESULTS Taxonomic characteristics. In a previous report (42), we tentatively placed a dextranaseproducing, anaerobic, gram-positive, rodshaped bacterium from dental plaque in the genus Actinomyces. At that time, it was noted that our preliminary data did not specifically delineate between the genus Bifidobacterium and the genus Actinomyces. One of the major differences between these two genera is the ability of the Bifidobacterium to produce significant amounts of acetic acid, resulting in a ratio of acetic acid to lactic acid of between 2 and 3 to 1 (8, 9, 36). Gas-liquid chromatographic analysis of the end products released during growth of the organisms in a glucose broth indicated that the microbe produced very little acetic acid (Table 1). This confirmed our initial classification of these organisms as Actinomyces spp. The macrocolonies of the dextranase-producing Actinomyces strain Gl are cream-white in color and are similar to the "molar tooth" or "raspberry-like" colonies (40) usually associated with A. israelii (Fig. la). The dextranActinomyces strains G2 and G3 hydrolyzing of of p-bis[2-(5-phenyloxazolyl)lbenzene; 1.0 liter round colonies with a slight convex, form soft, toluene) and counted with a Packard 2425 liquid scintillation spectrometer (Packard Instruments Inc., granular texture to the surface of the white
558
STAAT AND SCHACHTELE
INFECT. IMMUN.
starch supported weak fermentation, whereas arabinose and xylose were not fermented. The organisms failed to hydrolyze starch agar medium. These microbes grew anaerobically in the presence of CO2; however, no growth on Trypticase soy blood agar has been detected for any of the isolates under aerobic conditions. On the basis of the organism's ability to ferment ribose to acid and on its inability to produce observable growth on Trypticase soyTABLE 1. Distribution of fermentation products blood agar when grown aerobically (14, 40, 44), resulting from eInaerobic growth of Actinomyces we suggest that these dextranase-producing strains Gl, G2, and G3 in peptone-yeast microbes be considered strains of A. israelii. extract-glucose broth Enzyme characterization: evaluation of the [8H]NaBH4 dextranase assay. Figure 2 shows % of total acid produceda Acid that incorporation of borohydride derived 3H Gi G2 G3 into nonhydrolyzed dextran T40 is linear with respect to dextran concentration. The borohy9.3 4.2 6.2 Acetic ......... dride reaction is time and temperature dependtb 7.5 Formic ........ 0 ent and often requires extended incubation for 0 0 t Propionic ...... completion (30). Reacting dextran T40 or Tio 0 0 0.5 Butyric ........ 92.6 90.6 Lactic ......... 62.6 with [3H]NaBH4 at 37 C for various time pe0 1.8 Pyruvic ........ 6.5 riods indicated that reduction of these dextrans 3.2 Succinic ....... 14.0 2.7 neared completion after 30 min. Therefore, aMeq of individual acid/meq of total acid x 100 as incubation for 45 min at 37 C was used for the borohydride-reducing dextran assay. Figure 3 determined by gas-liquid chromatography. illustrates the incorporation of 3H into oligosact, Trace.
colonies (Fig. lb), similar to the smooth colonies of both A. israelii and Actinomyces naeslundii (44). Results of the biochemical reactions for species characterization were identical for the three dextranase-producing strains, except for strain G3 which could not hydrolyze esculin. Data from cultures grown in medium supplemented with various carbohydrates indicated that ribose was fermented to acid and soluble
FIG. 1. Colonial morphologies of dextranase-producing Actinomyces; (a) strain Gl, (b) strain G2. Strain G3 is similar to G2.
A. ISRAELII DEXTRANASE
VOL 12, 1975
have expected had the organism catabolized the dextran (42). This suggested that the supplemental dextran did not contribute to the growth of the bacterium. This was confirmed by growing cells in medium devoid of supplemental carbohydrate. The growth curve produced in this experiment was essentially identical to the one in Fig. 4 (data not shown). A. israelii utilizes glucose for growth, and dextranase activity was detectable in the supernatant of glucose-grown cells. Figure 5 illustrates that cells grown in glucose broth produced dextranase during the logarithmic growth phase. Although the total growth and dextranase production was greater for glucose-grown cultures compared to cultures supplemented only with dextran, the shape of the growth curves and the specific enzyme activities were similar in both experiments. To date, no growth conditions have been found which suppress dextranase production by A. israelii strains G1-3.
8 7
6 C
5
0
7t
0.
g)
559
4 3
2
0
250 200 150 pg Dextran T40 FIG. 2. Incorporation of sodium borohydridederived 3H into the reducing ends of dextran T40. Dextran T40 was added to 50 ;,1 of 0.2 N [3HJNaBH4 (106 counts/min) in 0.1 N NaOH and incubated at 37 C for I h. 50
100
6 5
charides cleaved from dextran T40 by the A. I) 4 israelii dextranases (open circles) compared to heat-inactivated enzyme (closed circles). This 0 curve is similar to that obtained with the colorimetric reducing sugar assay using the same enzyme (42). Validation that the [3H]NaNH4 was in sufficient excess for complete carbohydrate reduction was obtained by incubating 1% glucose in the system used to show linearity of the 3H incorporation into dextran T,4, The glucose did not affect the amount of 3H incorporated into the dextran T40 nor was the slope of the line altered (data not shown). Conditions for enzyme production. The effect of different carbohydrate supplements on growth and dextranase production by A. israelii was studied. The growth curve of strain Gl with dextran T40 as the carbohydrate supplement is shown in Fig. 4 (open circles). Dextranase ac60 40 20 0 tivity (closed circles) was greatest as the culture shifted from the logarithmic to the stationary Time phase of growth. The rapid inactivation of the enzyme activity is presumed to be caused by FIG. 3. Incorporation of sodium borohydrideproteolytic activity in the post-log culture derived 3H into the glucans released from dextran T740 by the A. israelii Gl dextranase. Symbols: 0, active supernatant. It was noted previously that A. israelii did not enzyme; 0, heat-inactivated (95 C for 10 min) enferment dextran to an acid pH as one would zyme.
12
(min)
560
INFECT. IMMUN.
STAAT AND SCHACHTELE
.8
6K2 c
_2
E
-2
->O\
4
0
Substrate specificity. The ability of A. israelii Gl dextranase to hydrolyze various carbohydrates was determined, and the results are presented in Table 3. The predominantly a-1,6linked polysaccharide, dextran, was the only substrate hydrolyzed. End product analysis. Complete hydrolysis of dextran T40 by the A. israelii Gl dextranase
L, 0
2
4
-_i6 __-.8 rime
10
0.4
o 2
24
hours)
FIG. 4. Growth and enzyme production curves for A. israelii GJ grown anaerobically with dextran T40 as the supplemental carbon source at a 1% concentration. Symbols: 0, optical density (turbidity) of culture at 600 nm; 0, dextranase activity assayed with the [3H]NaBH4 procedure.
0 QI)
0.3
0.2
0
-o L.. 0 'I,
2.0 z
>
-o
0.1
E
_\X
8.0
'D c
0
_:
D
T
_
.D
pH
Ir
ci 4,I -r
FIG. 6. pH optimum of the crude A. israelii GI dextranase. Dextranase activity was determined in the standard dextranase reaction (0.1 ml of enzyme) as the increased reducing sugar released after 45 min. Buffers: pH 4.0 to 5.5, sodium citrate; pH 6.0 to 7.5,
potassium phosphate. 4
6
8
10
12
24
Time (hours)
FIG. 5. Growth and enzyme production curves for A. israelii GI grown anaerobically with glucose as the supplemental carbon source at a 1% concentration. Symbols: 0, optical density (turbidity) of culture at 600 nm; 0, dextranase activity assayed with the ['H]NaBH4 procedure.
pH optimum. The pH optimum for the A. israelii dextranase centers around 6.3 (Fig. 6). The activity was relatively stable over the pH range 5.6 to 7.0. Inhibition studies. Among the metal ions tested, Ag+ and Hg2+ at 1 mM were the only inhibitors of the A. israelii dextranase (Table 2). Additonally, no metal ions caused activation of the dextranase nor did the metal chelator ethylenediaminetetraacetic acid cause any reduction in dextranase activity. The sulfhydryl reagents, dithiothreitol and iodoacetamide, had no effect at 10 mM concentrations, whereas the anionic detergent, sodium dodecyl sulfate, showed complete inhibition of the enzyme.
TABLE 2. Effect of some metal salts and other chemicals on the A. israelii Gl dextranase Chemicala
Concn (mM)
None
CaCl2 AgNO3
1 1
HgCl2 MnCl2
1 1
Relative 100 105 0 0 101
MgCl2
1
95
ZnSO4 EDTA SDS DTT IA
1
100
10 10 10 10
103 0 103 100
a EDTA, Ethylenediaminetetraacetic acid; SDS, sodium dodecyl sulfate; DTT, dithiothreitol; IA, iodoacetamide. 'The chemical was added to the standard reaction mixture at to, and 0.2-ml aliquots were taken at t20 and t40 and assayed for reducing power with the [3H JNaBH4 procedure. Relative activity = (t40 -
t20)chem/(t40 - t20)none x 100.
A. ISRAELII DEXTRANASE
VOL 12, 1975
resulted in the formation of oligosaccharides containing approximately 10 to 18 glucose units (42). In an attempt to visualize the dextran degradation, typical dextranase reaction mixtures were chromatographed at various stages of hydrolysis on Bio Gel P10 (Fig. 7). After 1 h, a small amount of apparent end product appeared in the area of fraction 38. At the 2-h mark, significant amounts of end product had appeared, and, essentially, none of the original dextran remained intact. Also, after 2 h, the polysaccharides were distributed into two distinct molecular weight ranges presumably corresponding to degradation intermediates and product. The 6-h pattern illustrates the continued movement of the intermediate peak to the right. The double-peaked pattern seen at 24 h remained unchanged through 48 h of incubation. The position of the double peak pattern corresponds to dextrans of approximately 10 to 18 glucose units in size (42).
DISCUSSION Dextranases produced by the oral strains of A. israelii isolated in our laboratories are extracellular endohydrolytic enzymes. Other bacteria capable of producing extracellular dextranases belong to the genera Pseudomonas (34), Bifidobacterium (1, 2, 6), Bacteriodes (24, 39), and Cellvibrio (28). Phylogenetically, the Actinomyces are closely related to the Bifidobacterium (orginally thought to be Lactobacillus [33]); however, comparison of the dextranases produced by these organisms reveals substantial differences. Contrasting the physiological characteristics of the two microbes, one can note the failure of A. israelii to ferment or apparently utilize dextrans in any way during growth, whereas the Bifidobacterium effectively utilize dextrans as the sole carbon source (6). Additionally, the A. israelii dextranase appears TABLE 3. Substrate specificity of concentrated crude A. israelii Gl dextranase
(Cogcn)
Glucan
Dextran T4, ... Amylose Glycogen ..... Pseudonigeran Laminaran ... Isomaltose
Linkage
Activitya
12 6
a-1,6; a-1,3 a-1,4
+
6 4
a-1,4; a-1,6
-
561
E
I
Frocticm I-"ai* FIG. 7. Bio Gel P-10 column chromatography of the reaction products produced by A. israelii Gl dextranase at various stages of dextran T40 hydrolysis.
a constitutive enzyme, whereas the Bifidobacterium dextranase is an inducible enzyme (1, 6). Comparative enzymology of the dextranases from the two different genera indicates that I both retain activity over the same broad pH was noted after 90 min of incubation of the standard range (5.5 to 7.0), and both appear to hydrolyze reaction system compared to boiled (95 C, 10 min) only linear a-1,6-linked glucans. The Bifidobacterium dextranase hydrolyzes dextrans to oligoenzyme preparations.
a-1,3 6 ,-1,3 0.6 a-1,6 Activity was considered positive (+) if an increase of 10% in the reducing sugar assay (Nelson-Somogyi) ....
to be
562
INFECT. IMMUN.
STAAT AND SCHACHTELE
saccharides 3 to 5 glucose units in length (2, 3), whereas the A. israelii dextranase apparently hydrolyzes dextran T40 into at least two types of oligosaccharides 10 to 18 glucose units in size. The progressive development of the doublepeaked, end product curve by the A. israelii dextranase (Fig. 7) can be explained as a nonrandom enzymatic recognition of the end of the dextran molecule with subsequent release of oligosaccharides of relatively specific length from the dextran. This could imply that the larger end product oligosaccharides are limit dextrans and that a dextran molecule of possibly 20 glucose units is needed before hydrolysis can be initiated by the A. israelii dextranase. The physiological significance of the A. israelii dextranase is not understood at the present time, and the production of dextranase by this microbe is inconsistent with the overview of extracellular microbial hydrolase function. The dextranase apparently does not contribute to the accumulation of metabolic requirements for the cells, since growth and dextranase production were equivalent in the presence or absence of the substrate (Fig. 4). Additionally, glucose did not suppress dextranase production by A. israelii (Fig. 5). These data can be explained by suggesting that an as yet unidentified molecule is the primary substrate for the enzyme and that cleavage of large oligosaccharides from dextrans is a fortuitous side reaction. An alternate explanation is that dextranase production by A. israelii constitutes a contribution to a symbiotic relationship established in the complex milieu of dental plaque. A. israelii is often recovered from the plaque of healthy individuals (26, 27, 40), and production of dextranase by selected strains may enhance the microbes' ability to maintain itself in plaque through association with- the dextran-forming microorganisms. In preliminary experiments, we recovered dextranase-producing microbes resembling A. israelii about 60% of the time from the plaque of elementrary school children (R. Staat and C. Schachtele, unpublished data). The effects of indigenous dextranase activities on the structure or composition of dental plaque can manifest themselves in several ways. Oral microbes such as Bacteroides ochraceus induce dextranases in response to the presence of the substrate and ferment the released glucose to acid (R. Staat and C. Schachtele, submitted for publication). The net result of this activity would be a reduced dextran pool in plaque. However, organisms such as A. israelii arid S. mutans produce dextranases, but do not metabolize the reaction products to acid; thus,
they apparently do not by themselves decrease the amount of dextran in plaque. This is not to suggest that the dextranases do not contribute to the metabolism of dental plaque. Walker proposed that oligosaccharides produced by endohydrolytic dextranase activity acted as alternate acceptor molecules in the S. mutans dextransucrase reaction (45). This concept was extended to account for the possible function of the S. mutans dextranase as a regulatory mechanism for the production of highly branched, dextranase-resistant glucans by that organism's dextransucrase (21, 43). Recently, our laboratory presented data indicating that the endohydrolytic dextranase produced by A. israelii reduces both the production of water-insoluble glucans by S. mutans 6715 dextransucrase and the adherence of whole organisms to glass surfaces (38). It should be noted that the A. israelii dextranase does not reduce the rate of total polysaccharide synthesis by S. mutans 6715 dextransucrase (R. Staat, unpublished data). Therefore, it can be proposed that the ability of S. mutans to establish as a member of the plaque flora is regulated in part by the qualitative structure of the glucan rather than the quantity of glucan produced. If this proposal is valid, then extracellular dextranases produced by the indigenous plaque flora have the potential for controlling the establishment of S. mutans by structural modifications of glucans and that elimination of the glucans from dental plaque need not be a prerequisite for ecological control of this bacterium. ACKNOWLEDGMENTS We thank L. V. Holdeman and her staff at the Virginia Polytechnic Institute Anaerobe Laboratory for the taxonomic data. The technical assistance of K. Cummings and WoonLam S. Leung is acknowledged. This work was supported by Public Health Service grant DE 02654 from the National Institute of Dental Research. R.H.S. was the recipient of National Institute of Dental Research Individual Research Fellowship Award 1 F22 DE00709. C.F.S. was the recipient of Public Health Service Career Development Award K4-DE-42,859.
LITERATURE CITED 1. Bailey, R. W., and R. T. J. Clarke. 1959. A bacterial dextranase. Biochem. J. 72:49-54. 2. Bailey, R. W., D. H. Hutson, and H. Weigel. 1960. Action of bacterial dextranase on branched dextrans. Nature
(London) 186:553-554. 3. Bailey, R. W., D. H. Hutson, and H. Weigel. 1961. The action of a Lactobbacillus bifidus dextranase on a branched dextran. Biochem. J. 80:514-519. 4. Bowen, W. H. 1968. Effects of dextranase on cariogenic and noncariogenic dextrans. Br. Dent. J. 124:347-349. 5. Bowen, W. H. 1971. The effect of dextranase on caries activity in monkeys (Macaca irus). Br. Dent. J. 131:445-449. 6. Clarke, R. 'F. 1. 1959. A dextran-fermenting organism
VOL 12, 1975 from the rumen closely resembling Lactobacillus bifidus. J. Gen. Microbiol. 20:549-553. 7. Conrad, H. E., E. Varboncouer, and M. E. James. 1973. Qualitative and quantitative analysis of reducing carbohydrates by radiochromatography on ion-exchange papers. Anal. Chem. 51:486-500. 8. de Vries, W., S. J. Gerbrandy, and A. H. Stouthamer. 1967. Carbohydrate metabolism in Bifidobacterium bifidum. Biochim. Biophys. Acta 136:415-425. 9. de Vries, W., and A. H. Stouthamer. 1968. Fermentation of glucose, lactose, galactose, mannitol, and xylose by bifidobacteria. J. Bacteriol. 96:472-478. 10. Dubois, M., K. A. Gilles, J. K. Hamilton, P. A. Rebers, and F. Smith. 1956. Colorimetric method for the determination of sugars and related substances. Anal. Chem. 28:350-356. 11. Ebisu, S., A. Misaki, K. Kato, and S. Kotani. 1974. The structure of water-insoluble glucans of cariogenic Streptococcus mutans, formed in the absence and presence of dextranase. Carbohydr. Res. 38:374-381. 12. Fitzgerald, R. J., P. H. Keyes, T. H. Stoudt, and D. Spinell. 1968. The effects of a dextranase preparation on plaque and caries in hamsters, a preliminary report. J. Am. Dent. Assoc. 76:301-304. 13. Fitzgerald, R. J., D. M. Spinell, and T. H. Stoudt. 1968. Enzymatic removal of artificial plaques. Arch. Oral Biol. 13:125-128. 14. Georg, L. K., G. W. Roberstad, and S. A. Brinkman. 1964. Identification of species of Actinomyces. J. Bacteriol. 88:477-490. 15. Germaine, G. R., C. F. Schachtele, and A. M. Chludzinski. 1974. Rapid filter paper assay for the dextransucrase activity from Streptococcus mutans. J. Dent. Res. 53:1355-1360. 16. Gibbons, R. J. 1968. Formation and significance of bacterial polysaccharides in caries etiology. Caries Res. 2:164-171. 17. Gibbons, R. J., and S. B. Banghart. 1967. Synthesis of extracellular dextran by cariogenic bacteria and its presence in human dental plaque. Arch. Oral Biol. 12:11-24. 18. Gibbons, R. J., and R. J. Fitzgerald. 1969. Dextraninduced agglutination of Streptococcus mutans and its potential role in the formation of microbial dental plaques. J. Bacteriol. 98:341-346. 19. Gibbons, R. J., and M. Nygaard. 1968. Synthesis of insoluble dextran and its significance in the formation of gelatinous deposits by plaque-forming streptococci. Arch. Oral Biol. 13:1249-1262. 20. Guggenheim, B. 1970. Enzymatic hydrolysis and structure of water-insoluble glucan produced by glycosyltransferases from a strain of Streptococcus mutans. Helv. Odontol. Acta 5 (Suppl. 14):89-108. 21. Guggenheim, B., and J. J. Burckhardt. 1974. Isolation and properties of a dextranase from Streptococcus mutans OMZ 176. Helv. Odontol. Acta 18:101-113. 22. Guggenheim, B., and E. Newbrun. 1969. Extracellular glucosyltransferase activity of an HS strain of Streptococcus mutans. Helv. Odontol. Acta 13:84-97. 23. Guggenheim, B., and H. E. Schroeder. 1968. Biochemical and morphological aspects of extracellular polysaccharides produced by cariogenic streptococci. Helv. Odont. Acta 11:131-152. 24. Hehre, E. J., and T. W. Sery. 1952. Dextran-splitting anaerobic bacteria from the human intestine. J. Bacteriol. 63:424-426. 25. Holdeman, L. V., and W. E. C. Moore. 1973. Anaerobe laboratory manual, 2nd ed. Virginia Polytechnic Insti-
A. ISRAELII DEXTRANASE
563
tute and State University, Blacksburg. 26. Howell, A., W. C. Murphy, F. Paul, and R. M. Stephan. 1959. Oral strains of Actinomyces. J. Bacteriol. 78:82-95. 27. Howell, A., R. M. Stephan, and F. Paul. 1962. Prevalence of Actinomyces israelii, A. naeslundii, Bacterionema matruchotti and Candida albicans in selected areas of the oral cavity and saliva. J. Dent. Res. 41:1050-1059. 28. Ingelman, G. 1948. Enzymatic breakdown of dextran. Acta Chem. Scand. 2:803-812. 29. 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. 30. McLean, C., D. A. Werner, and D. Aminoff. 1973. Quantitative determination of reducing sugars, oligosaccharides, and glycoproteins with ('H) borohydride. Anal. Biochem. 55:72-84. 31. Nelson, N. 1944. A photometric adaptation of the Somogyi method for the determination of glucose. J. Biol. Chem. 153:375-380. 32. Newbrun, E. 1972. Extracellular polysaccharides synthesized by glucosyltransferases of oral streptococci. Caries Res. 6:132-147. 33. Poupard, J. A., I. Husain, and R. F. Norris. 1973. Biology of the bifidobacteria. Bacteriol. Rev. 37:136-165. 34. Richards, G. N., and M. Streamer. 1972. Studies on dextranases. I. Isolation of extracellular, bacterial dextranases. Carbohydr. Res. 25:323-332. 35. Richards, G. N., and W. J. Whelan. 1973. Determination of the number-average molecular weight of polysaccharides by end-group reduction with borohydride-t. Carbohydr. Res. 27:185-191. 36. Scardovi, V., and L. D. Trovatelli. 1965. The fructose6-phosphate shunt as a peculiar pattern of hexose degradation in the genus Bifidobacterium. Ann. Microbiol. Enzymol: 15:19-29. 37. Schachtele, C. F., A. E. Loken, and M. K. Schmitt. 1972. Use of specifically labeled sucrose for comparison of extracellular glucan and fructan metabolism by oral streptococci. Infect. Immun. 5:263-266. 38. Schachtele, C. F., R. H. Staat, and S. K. Harlander. 1975. Dextranases from oral bacteria: inhibition of water-insoluble glucan production and adherence to smooth surfaces by Streptococcus mutans. Infect. Immun. 12:309-317. 39. Sery, T. W., and E. J. Hehre. 1956. Degradation of dextrans by enzymes of intestinal bacteria. J. Bacteriol. 71:373-380. 40. Slack, J. M. 1974. Actinomyces, p. 660-667. in R. E. Buchanan and N. E. Gibbons. (ed.), Bergey's manual of determinative bacteriology, 8th ed. The Williams & Wilkins Co., Baltimore. 41. Somogyi, M. 1945. A new reagent for the determination of sugars. J. Biol. Chem. 160:61-68. 42. Staat, R. H., T. H. Gawronski, and C. F. Schachtele. 1973. Detection and preliminary studies on dextranaseproducing microorganisms from human dental plaque. Infect. Immun. 8:1009-1016. 43. Staat, R. H., and C. F. Schachtele. 1974. Evaluation of dextranase production by the cariogenic bacterium Streptococcus mutans. Infect. Immun. 9:467-469. 44. Thompson, L., and S. A. Lovestedt. 1951. An Actinomyces-like organism obtained from the human mouth. Proc. Staff Meet. Mayo Clin. 26:169-175. 45. Walker, G. J. 1972. Some properties of a dextranglucosidase isolated from streptococci and its use in studies on dextran synthesis. J. Dent. Res. 51:409-414.
