INFECTION AND IMMUNITY, Sept. 1974, p. 597-604Copyright 0 1974 American Society for Microbiology

Vol. 10, No. 3Printed in U.S.A.

Measurement of Intracellular Jodophilic Polysaccharide in TwoCariogenic Strains of Streptococcus mutans by Cytochemical

and Chemical MethodsJ. R. DiPERSIO, S. J. MATTINGLY, M. L. HIGGINS, AND G. D. SHOCKMAN

Department of Microbiology and Immunology, Temple University School of Medicine, Philadelphia,Pennsylvania 19140

Received for publication 24 May 1974

Quantitative cytological and chemical methods have been developed to studythe intracellular iodophilic polysaccharide (IPS) content of two strains ofStreptococcus mutans. The cytological method uses a periodic acid-chloritetreatment of thin sections to increase the affinity of IPS for uranyl and lead salts.This results in the IPS appearing as individual electron-dense granules which canbe counted for quantitative studies. As a basis for these quantitative studies, IPSwas measured chemically by dissolving whole cells with hot KOH and quantitat-ing spectrophotometrically the amount of iodine-polysaccharide complexformed.

Ultrastructural observations of thin sectionsof bacteria known to contain intracellular poly-saccharides have shown the presence of dif-fusely outlined, relatively electron-transparentcytoplasmic areas about 60 to 100 nm in diame-ter (2, 4). The appearance of these areas reflectsin general the low electron-scattering character-istics common to most polysaccharides. Unlikethe glycogen particles seen in thin sections ofvarious animal tissues, bacterial glycogen gran-ules usually stain only very weakly or not at allwith uranyl and lead salts. Attempts to specifi-cally stain intracellular polysaccharide granulesin thin sections of bacteria, using carbohydrate-specific stains such as periodate-Schiff reagentor thiosemicarbazide, have met with some suc-cess, but clear morphological differentiation ofindividual granules has not been attained.A method involving the treatment of thin

sections with periodic acid and an acidifiedsolution of sodium chlorite, followed by uranylacetate and lead citrate staining, was previouslyused to stain glycogen granules in thin sectionsof Nocardia asteroides (J. R. DiPersio and S. J.Deal, J. Gen. Microbiol., in press). Althoughthe granules appear deeply stained and mor-phologically distinct, the treatment damagedother cellular components. The present workdescribes an improved method for stainingintracellular iodophilic polysaccharide (IPS)granules present in two strains of the cariogenicoral bacterium, Streptococcus mutans.To evaluate the quantitative potential of this

cytochemical method, the amounts of IPS pres-

ent in cells were quantitated chemically, using amethod based on the formation of an 12-polysac-charide complex. We have used both the cyto-chemical and chemical methods in concert tostudy the amount of IPS found in 24-h station-ary-phase cultures of two strains of S. mutans.

MATERIALS AND METHODSOrganism. S. mutans strain FA-1 was kindly

supplied by A. Bleiweis, Department of Microbiology,University of Florida, Gainesville. S. mutansOMZ-176 was obtained from B. Guggenheim, Univer-sity of Zurich, Zurich, Switzerland. Stock cultureswere maintained on Todd-Hewitt 2% (wt/vol) agarslants and stored at 4 C.Medium. The chemically defined medium em-

ployed in all experiments was a modification of themedium previously described for the growth of Strep-tococcus faecalis (S. faecium) 9790 (15a). A prelimi-nary description of this medium for the growth of oralstreptococci has been given (B. Terleckyj, Mastersthesis, Temple University, Philadelphia, Pa., 1974).The major differences between the medium used inthis study and that used for S. faecalis are the use ofa lower phosphate concentration (0.043 M instead of0.3 M), the addition of sodium citrate (0.01 M), anda doubling of the concentration of all vitamins. Theaddition of sodium bicarbonate (0.01 M) was foundto stimulate growth of strain FA-1 and was addedaseptically to the growth medium just before use.Inocula were prepared by growing cells obtained fromTodd-Hewitt 2% agar slants in 10 ml of the chemicallydefined medium for 24 h.

Growth determination. Growth was measuredturbidimetrically at 675 nm in a Coleman model 14spectrophotometer. Turbidity was expressed in ad-justed optical density (AOD) units (14). One AOD

597

on May 14, 2018 by guest

http://iai.asm.org/

Dow

nloaded from

598 DIPERSI

unit is the equivalent of 0.39 Asg cellular dry weight perml. The dry weight of cell suspensions was measuredgravimetrically with drying in vacuo over H2SO4 andP206. Routinely, growth was measured in either500-ml side-arm flasks containing 300 ml of mediumor 18- by 100-mm optically calibrated tubes contain-ing 10 ml of medium. The time of entrance of thecultures into stationary-phase growth was defined asthe initial point in a semilogarithmic plot of AODversus time at which the rate of growth deviated fromlinearity. Samples for electron microscopy were col-lected 16 h after this deviation had taken place.

Chemical determination of IPS content. Samplesof whole cells removed from cultures were heated for 5min at 100 C and collected by centrifugation at 2,000x g for 10 min. After two washes with cold double-dis-tilled water, the cell suspensions were adjusted to1,000- AOD units. One-milliliter amounts were addedto calibrated 13- by 100-mm optically calibratedtubes, and 0.3 ml of KOH (5.3 M) was added to eachtube. Tubes were covered and placed in a boiling-water bath for 90 min. After cooling, the now clearsolutions were neutralized by the addition of 0.3 ml ofHCl (5.3 M). The tubes were vortexed and 1.0 ml of1.0 M potassium phosphate, pH 7.0, was added. Aftervortexing, 0.6 ml of a freshly prepared 0.2% (wt/vol)iodine in 2.0% (wt/vol) potassium iodide solution wasadded with mixing. Tubes were read at 520 nm in aBausch and Lomb spectronic 20 colorimeter. Theyellow-brown iodine-polysaccharide complex was sta-ble for 2 h. In the determination of the optimalwavelength for the assay, 0.1 ml of the iodine-potas-sium iodide solution was added to avoid a highbackground blank at wavelengths below 470 nm.

Glycogen isolation and purification. Cells froman overnight culture (grown in a 2-liter flask contain-ing 1 liter of chemically defined medium) of S.mutans FA-1 were harvested by centrifugation andwashed twice with cold (4 C) double-distilled water. Asolution of 30% (wt/vol) KOH was added to the cells(2 g wet weight) to bring the final volume to 100 ml.The suspension was placed in a 100-C water bath for90 min. After cooling in an ice bath for 30 min, theextracts were centrifuged (2,000 x g, 20 min) andinsoluble material was discarded. An equal volume of95% (wt/vol) ethanol was added to the clear yellowsupematant solution, and the precipitated polysac-charides was collected by centrifugation. Polysaccha-ride was redissolved in 50 ml of water, further purifiedby two additional alcohol precipitations, and lyophi-lized for future use.

Fixation and embedding. Glutaraldehyde wasadded to 25-ml samples of 16-h stationary-phasecultures to give a final concentration of 3% (vol/vol).After fixation at room temperature for 30 min, cellswere pelleted by centrifugation (2,000 x g, 15 min)and fixed overnight at 4 C with 3% (wt/vol) glutaral-dehyde in 0.01 M phosphate, pH 6.2, containing 0.8Mpotassium chloride and 0.01 M magnesium acetate.The next day, cells were washed four times with theabove buffer at 4 C and allowed to remain overnight inthe wash buffer in the cold. The following morningcells were washed four additional times with coldbuffer and suspended for 30 min in the Veronal-ace-tate buffer of Kellenberger et al. (11). Cells were

) ET AL. INFECT. IMMUNITY

postfixed overnight a room temperature with 1%(wt/vol) osmium tetroxide, treated with uranyl ace-tate by the Kellenberger et al. procedure, and thenplaced into 2% (wt/vol) agar made with Kellenbergerbuffer.Hardened blocks approximately 0.05-mm square

were embedded in Epon 812 according to the methodof Luft (12), with the following modifications. Afterethanol dehydration, cell-containing agar blocks werewashed with four 15-min changes of propylene oxide,with care not to exceed 1 h. The cells were suspendedin a mixture of propylene oxide and Epon 812 (50:50)in capped vials, placed on a rotator at six rotations permin for 2 h, and left uncapped overnight. The nextmorning the agar blocks were embedded in fresh Epon812.

Staining. Silver-grey sections were cut on a Rei-chert OMU-2 ultramicrotome using a diamond knifeand placed on Formvar-carbon-coated copper grids.Sections were stained for 20 min at 45 C in 50%(wt/vol) ethanol-saturated uranyl acetate and for 10min in lead citrate (13). Specimens were examined ina Siemens Elmiskop IA electron microscope fre-quently calibrated with a carbon grating replica (E. F.Fullam Inc., Schenectady, N.Y.) at magnificationsranging from 10,000 to 30,000.

Intracellular polysaccharide granules were stainedby a modification of the original procedure of Reveland Karnovsky (In DiPersio and Deal, J. Gen. Mi-crobiol., in press), which was designed to stain glyco-gen in thin sections of animal tissue. Thin sectionswere mounted on Formvar-carbon-coated nickel gridsand treated with a 1% (wt/vol) solution of periodicacid for 30 min at room temperature. After threerinses in distilled water, grids were treated with asolution of sodium chlorite prepared as follows: 500mg of sodium chlorite was dissolved in 18 ml ofdistilled water just prior to use. To this was added 2.0ml of 10 N acetic acid. A 1 to 50 dilution of theacidified solution was used to treat grids for 10 min.After three rinses in distilled water, grids were post-stained with alcoholic uranyl acetate and lead citrateas previously described. Omission of the periodic acidoxidation step in the glycogen staining procedureserved as a control.

Quantitative electron microscopy. Only thosesections with distinct trilaminar membrane profilesaround the entire cell perimeter were used for quanti-tation of IPS granules. Previous studies of otherstreptococci have shown that cell sections with suchmembrane profiles are both central and axial (9). Theaverage number of IPS granules per cell section wasdetermined by studying at least 40 cell sections whichhad previously been treated with the periodic acid-chlorite procedure and stained with uranyl and leadsalts.Direct comparison of the results obtained from

enumerating numbers of granules per cell section andthe percent content of IPS was not possible becausecells of strain FA-1 were considerably larger thanthose of OMZ-176. Therefore, it was necessary todetermine with an electronic planimeter (NumonicsCorp., North Wales, Pa.) the average area of the cellsections found in the two cell populations.

IC

on May 14, 2018 by guest

http://iai.asm.org/

Dow

nloaded from

POLYSACCHARIDE STORAGE IN S. MUTANS

RESULTSElectron microscopy. Cells of two strains of

S. mutans (FA-1 and OMZ-176) were obtainedfrom 24-h stationary-phase cultures for study bythe thin section technique. After the usualuranyl acetate-lead citrate staining procedure,both strains showed round, diffusely outlinedcytoplasmic areas of low electron density about35 to 60 nm in diameter (Fig. 1A and 3A). Suchsections served as controls for the stainingprocedure.

In addition to having many more of theseareas, the cell wall margins of strain FA-1 weremuch more tribanded and less "fuzzy" thanthose of OMZ-176.When such sections were treated with 1%

(wt/vol) periodic acid and acid chlorite andthen stained with uranyl and lead salts, theelectron-transparent zones were seen as elec-

tron-dense and individually distinct entities inthe cytoplasm (Fig. 1B and 3B). The size of theelectron-dense zones appeared to be no differentthan the areas of low density seen in the controlcells (35 to 60 nm). Cells were not markedlydamaged. The presence of polysaccharide gran-ules completely filling the cytoplasmic area ofstrain FA-1 cells is not a scattered phenomenonbut is rather constant throughout the popula-tion of cells (Fig. 2).The use of undiluted chlorite reagent pro-

duced very dark granules but also resulted insignificant damage to walls and other cell com-ponents. Dilutions of the chlorite reagent above1: 50 resulted in poor staining of the polysaccha-ride granules, probably as a result of an insuffi-cient number of free carboxyl groups producedby oxidation with sodium chlorite. The use ofthe chlorite reagent at a 1:50 dilution produced

BFIG. 1. Thin sections of S. mutans strain FA-1 taken from a 24-h stationary-phase culture without (A) and

with (B) periodic acid-chlorite treatment before staining with uranyl acetate-lead citrate. Bar equals 100 nmand applies to (B) as well.

VOL. 10, 1974 599

on May 14, 2018 by guest

http://iai.asm.org/

Dow

nloaded from

DIPERSIO ET AL.

adequately stained granules with little damageto other cellular components. The omission ofthe periodic acid oxidation step from the proce-dure resulted in an absence of electron-densegranules, indicating the importance of free alde-hyde groups prior to further oxidation withchlorite (Fig. 4).The average number of granules found per

central axial section was determined by study-ing at least 40 sections taken from stationary-phase cultures of FA-1 and OMZ-176. Theperiodic acid-chlorite procedure was found to beessential for quantitation, for in its absence itwas impossible to determine the number ofgranules found in unstained aggregates (com-pare Fig. 1A with 1B). The number of denselystained IPS granules observed in sections ofstrain FA-1 averaged 282 ± 97 granules persection, whereas strain OMZ-176 showed only18.6 i 13 granules per section (Table 1).Chemical quantitation of intracellular

polysaccharide. Methods for the quantitationof bacterial glycogen-like polysaccharides usu-ally require chemical differentiation of homo-polymers containing glucose from other storageand structural polysaccharides. van Houte andJansen (16) used the reaction of glycogen-likepolysaccharides with I2 as a basis for spectro-photometrically quantitating iodophilic poly-saccharides in suspensions of intact cells. Thisprocedure required the use of an untreated cellsuspension of the same density as a blank. In

our hands, this procedure was useful with somestrains under some growth conditions. However,with other strains and growth conditions resultsobtained were not always reproducible. There-fore, the I2 method was modified as describedunder Materials and Methods to produce con-sistent and reproducible results. The basis ofthe method is to first dissolve the cells in hotKOH, neutralize, and add I2 in KI to thewater-clear solution to produce a brownish-yel-low color with an absorption maxima at 520 nm(Fig. 5) characteristic of glycogen-like polysac-charides. This color was stable for at least 2 h atroom temperature. Commercial rabbit liver gly-cogen (Sigma) and intracellular polysaccharideisolated and purified from S. mutans FA-1produced colors with similar absorption max-ima at 520 nm when treated in the same manner(Fig. 5).IPS isolated and purified from S. mutans

FA-1 was found to contain 90% carbohydrate(wt/wt) by the phenol-sulfuric acid method (1).Well over 90% of the carbohydrate was glucose,as determined by glucose oxidase and a differ-ential anthrone procedure (15). Purified IPSwas used to obtain a standard curve relatingIPS concentration (corrected for impurities) toabsorbance at 520 nm (Fig. 6). The color pro-duced was proportional to the IPS concentra-tion to well over 1.1 OD units. Suspensionscontaining increasing amounts of cells wereprepared from 24-h cultures of strain FA-1

-'i - :...

f "11 !.. .1

It

Le,1.4.Ol

FIG. 2. Thin section of a field of cells of S. mutans strain FA-1 taken from a 24-h stationary-phase culture withperiodic acid-chlorite treatment before staining with uranyl acetate-lead citrate. Bar equals 100 nm.

600 INFECT. IMMUNITY

W.1

4.4

on May 14, 2018 by guest

http://iai.asm.org/

Dow

nloaded from

POLYSACCHARIDE STORAGE IN S.MUTANS6

treated with hot KOH and then with the I2-KIreagent. The observed absorbance at 520 nmwas proportional to cell concentration up toabout 1,500 AOD of cells (equivalent to 0.59 mgcellular dry weight). This proportionality waslost when larger amounts of cells were used,presumably due to the presence of increasedamounts of organic material which requiredmore hydroxyl groups for polysaccharide extrac-tion. To ensure that all determinations werewithin the range of the assay, all subsequent

IPS determinations used 1,000 AOD (0.39 mgcellular dry weight per ml) of cells in each assaytube.The amounts of IPS present in the same 16-h

cultures of strains of FA-1 and OMZ-176 stud-ied by electron microscopy are shown in Table1.Comparison of chemical and cytochemical

estimations of cellular IPS. Table 1 shows acomparison of the estimates of IPS found in thetwo strains of S. mutans by the cytochemical

BFIG. 3. Thin sections of S. mutans strain OMZ-176 taken from a 24-h stationary-phase culture without (A)

and with (B) periodic acid-chlorite treatment before uranyl acetate-lead citrate staining. Bar in (A) equals 100nm and also applies to (B).

601VOL. 10, 1974

on May 14, 2018 by guest

http://iai.asm.org/

Dow

nloaded from

602 DIPERSI

and chemical procedures. The cytochemicaldata based on granule counts of cell sectionsindicates that strain FA-1 possesses about 15times more IPS than does strain OMZ-176,whereas the chemical determinations based onthe amount of IPS found per unit of cell masssuggests only an eightfold enrichment of iodo-philic material in strain FA-1. However, quanti-tative comparisons were distorted by the factthat cells of strain FA-1 are considerably larger

, 2:

J'.

) ET AL. INFECT. IMMUNITY

than those of strain OMZ-176 (compare Fig. 1Awith Fig. 3A). When the area of each cell sec-tion was measured with a planimeter and thenumber of granules was expressed as numberper unit area (Table 1), the cytochemical andchemical determinations were in good agree-ment. With both methods strain FA-1 con-tained about 7 to 8 times more IPS than didstrain OMZ-176. The observed close correlationbetween the chemical and cytological methods

FIG. 4. Thin section of S. mutans FA-1 taken from a 24-h stationary-phase culture. This section was treatedwith chlorite, but not with periodic acid, before being exposed to uranyl acetate-lead citrate stains. Note that,without periodate treatment, cytoplasmic granules are not rendered electron dense.

IC

on May 14, 2018 by guest

http://iai.asm.org/

Dow

nloaded from

POLYSACCHARIDE STORAGE IN S. MUTANS

TABLE 1. Comparison of IPS as determined by cytochemical and chemical methods

S. mutans 1 No. of granules Area per cell section No. of granules Content of IPSstrain per cell section (Um2) found per unit area (%)

______________________I ~~~~~~~~~~~~ofcell section

FA-1 282.0 i 97a 1.01 i 0.13" 279.2 40-42

OMZ-176 18.6 ± 13a 0.51 ± 0.13b 37.2 5

FA-1/OMZ-176 15.2 1.98 7.5 8

a Numbers represent the average number of granules per cell section plus or minus the standard deviation.b Numbers represent the average area per cell section plus or minus the standard deviation.c This result lies at or near the lower useful limit of the method.

WAVELENGTH - A (nm)FIG. 5. Comparison of the absorption spectra of

rabbit liver glycogen (A), isolated and purified IPSfrom S. mutans FA-I (-), and hydrolyzed whole cellsof S. mutans FA-I (-). Glycogen and whole cells were

treated with alkali as described in Materials andMethods. An iodine (0.2% in 2.0% KI) solution was

added, and the absorbance was determined at varyingwavelengths.

was obtained despite the fact that the relativelylow concentration of IPS in cells from overnightcultures of strain OMZ-176 was at the lowerlimit of the useful range of the chemicalmethod. It would seem that the cytochemicalmethod is capable of detecting lower levels ofIPS than is the chemical procedure.

DISCUSSION

Glycogen-like polysaccharide granules havebeen observed in thin sections of a number ofbacterial species, among them Streptococcusmitis (2), Escherichia coli (4), Arthrobacter

E 102C0I'.)

A0.8z83 ~~~~~~AOD

0m0.40

00 100 200 300 400IPS (jig/ml)

0 500 1000 1500 2000

AOD0/mIFIG. 6. Quantitation of alkali-extracted and al-

cohol-precipitated intracellular polysaccharide fromS. mutans FA-I (a) and alkali-treated whole cells ofS. mutans FA-I (0). The isolation and purification ofS. mutans FA-I IPS is described in Materials andMethods. Stationary-phase cells of S. mutans FA-Iwere suspended at various dry weights (1 AOD =0.39 ug/ml) and treated with KOH. An iodine (0.2% in2.0% KI) solution was added, and the absorbance ofthe cell hydrolysate and purified IPS was determinedat 520 nm.

crystallopoietes (3) and N. asteroides (DiPersioand Deal, J. Gen. Microbiol., in press). Theglycogen granules seen in these and other bacte-rial species have been quite comparable in bothsize and shape, appearing as round, diffuselyoutlined bodies about 60 to 100 nm in diameter.They differ, however, from the glycogen parti-cles seen in many animal tissue sections in thatthe bacterial granules do not readily stain withuranyl acetate and lead citrate. Cedergren andHolme (4) first described intracellular storagepolysaccharide granules as appearing as cyto-

VOL. 10, 1974 603

on May 14, 2018 by guest

http://iai.asm.org/

Dow

nloaded from

DIPERSIO ET AL.

plasmic "holes" in thin sections of E. coli.These "holes" were later found to stain weaklywith periodate-Schiff (10).

Several investigators have used various in-volved modifications of the thiosemicarbazide-osmium method of Hanker et al. (8) to stainintracellular as well as extracellular polysaccha-ride material of several oral species of bacteria(5, 7). With the use of this method, individualisolated glycogen granules appeared very darklystained; however, areas where granules wereconcentrated stained with such black intensityas to obscure the morphology of individualpolysaccharide granules.An adaptation of a method developed by

Revel and Kamofsky for staining glycogen par-ticles in thin sections of various animal tissueswas used effectively by DiPersio and Deal (J.Gen. Microbiol., in press) to stain intracellularglycogen granules in N. asteroides. A modifica-tion improving the overall staining action of thismethod for use with oral streptococci was suc-cessfully developed in the present work. Intra-cellular glycogen granules were preferentiallystained in such a way as to preserve theirmorphological identity even in areas of the cellwhere granules were densely clustered.

Application of these methods to stationary-phase cultures of S. mutans strains FA-1 andOMZ-176 suggests that when differences in cellsize are considered and the data are expressedas granules per unit area of cell, rather than on aper cell basis, both the chemical and cytochemi-cal methods yield comparable results. Obvi-ously, more extensive comparisons of resultsobtained by the two methods must be made.Both methods are currently being used to studyIPS storage and degradation in strain FA-1.Additionally, the chemical method is beingused to determine the effect of various environ-mental factors on IPS content of a variety of S.mutans strains representative of the variousserological types.

ACKNOWLEDGMENTSWe thank Sandra Sonntag and Mary O'Connor for their

assistance.This investigation was supported by Public Health Service

research grant DE-03487 from the National Institute ofDental Research. J. DiPersio was a postdoctoral traineesupported by Public Health Service training grant no.

AI-00233 from the National Institute of Allergy and InfectiousDiseases.

LITERATURE CITED

1. Ashwell, G. 1966. New colorimetric methods of sugaranalysis, p. 93. In E. F. Newfield and V. Ginsburg (ed.),Methods in enzymology, vol. 8. Academic Press Inc.,New York.

2. Berman, K. S., R. J. Gibbons, and J. Nalbandian. 1967.Localization of intracellular polysaccharide granules inStreptococcus mitis. Arch. Oral Biol. 12:1133-1138.

3. Boylen, C. W., and J. L. Pate. 1973. Fine structure ofArthrobacter crystallopoietes during long term starva-tion of rod and spherical stage cells. Can. J. Microbiol.19:1-5.

4. Cedergren, B., and T. Holme. 1959. On the glycogen of E.coli B. Electron microscopy of ultrathin sections ofcells. J. Ultrastruct. Res. 3:70-73.

5. Critchley, P., C. A. Saxton, and A. B. Kolendo. 1968. Thehistology and histochemistry of dental plaque. CariesRes. 2:115-129.

6. Gibbons, R. J., and S. S. Socransky. 1962. Intracellularpolysaccharide storage by organisms in dental plaques.Arch. Oral Biol. 7:73-80.

7. Guggenheim, B., and H. E. Schroeder. 1967. Biochemicaland morphological aspects of extracellular polysaccha-rides produced by cariogenic streptococci. Helv. Odont.Acta 11:131-152.

8. Hanker, J. S., A. R. Seaman, L. P. Weiss, H. Ueno, R. A.Bergman, and A. M. Seligman. 1964. Osmiophilicreagents: new cytochemical principle for light andelectron microscopy. Science 146:1039-1043.

9. Higgins, M. L., and G. D. Shockman. 1970. Early changesin ultrastructure of Streptococcus faecalis after aminoacid starvation. J. Bacteriol. 103:244-254.

10. Holme, T., and B. Cedergren. 1961. Demonstration ofintracellular polysaccharide in E. coli by electronmicroscopy and by cytochemical methods. Acta Pa-thol. Microbiol. Scand. 51:170-186.

11. Kellenberger, E., A. Ryter, and J. Sechaud. 1958. Elec-tron microscopic study of DNA-containing plasms. II.Vegetative and mature phage DNA as compared withnormal bacterial nucleoids in different physiologicalstates. J. Biophys. Biochem. Cytol. 4:671-678.

12. Luft, J. H. 1961. Improvements in epoxy resin embeddingmethods. J. Biophys. Biochem. Cytol. 9:409-414.

13. Reynolds, E. S. 1963. The use of lead citrate at high pH asan electron opaque stain in electron microscopy. J. CellBiol. 17:208-212.

14. Toennies, G., and D. L. Gallant. 1949. The relationshipbetween photometric turbidity and bacterial concen-tration. Growth 13:7-20.

15. Toennies, G., and J. J. Kolb. 1964. Carbohydrate analysisof bacterial substances by a new anthrone procedure.Anal. Biochem. 8:54-69.

15a. Toennies, G., and G. D. Shockman. 1958. Growth chem-istry of Streptococcus faecalis. Proc. 4th Int. Congr.Biochem. Colloq. 13:365-393.

16. van Houte, J., and H. M. Jansen. 1968. The iodophilicpolysaccharide synthesized by Streptococcussalivarius. Caries Res. 2:47-56.

604 INFECT. IMMUNITY

on May 14, 2018 by guest

http://iai.asm.org/

Dow

nloaded from