Biofilm Journal Pontificia Universidad Católica de Valparaíso ISSN: 1360-3655 Vol. 2, Num. 1, 1997

Susceptibility of biofilms of Streptococccus sanguis and Actinomyces naeslundii to chlorhexidine

Tove Larsen*, Kaj Stoltze and Nils-Erik Fiehn

Department of Oral Microbiology and Department of Periodontology School of Dentistry, Faculty of Health Sciences, University of Copenhagen, Norre Alle 20, DK - 2200, Copenhagen N, Denmark.
e-mail: tl@odont.ku.dk

* Corresponding author

Date received: May 14th 1997
Date accepted: July 16th 1997
Date Published: July 17th 1997

Code Number: BF97001
Sizes of Files:
    Text: 27.7K
    Graphics files: line drawings (gif) - 66.8K

ABSTRACT

Single species and two species biofilms comprising Streptococcus sanquis and Actinomyces naeslundii were established in order to determine the susceptibility to chlorhexidine of the bacteria growing in biofilms compared to planktonic growth, and when growing in single species biofilms compared to mixed biofilms. The MIC of the bacteria to chlorhexidine was determined by a broth dilution method. Single and two species biofilms were established in a flow model in a modified Robbins device and subsequently exposed to chlorhexidine for 48 h at concentrations from 10 to 2000 mg/ml. Further, two species biofilms were exposed to 1000 mg/ml of chlorhexidine in 10 min pulses two or six times a day. The biofilm cell number was determined by viable counting at different time intervals. Two species biofilms were established faster than single species biofilms, but after establishment A. naeslundii was suppressed by S. sanguis. When lower concentrations of chlorhexidine were added, however, A. naeslundii also remained in the biofilm. Resistance of S. sanguis and A. naeslundii to chlorhexidine increased considerably when growing in biofilms and >100 mg/ml was needed to eliminate growth of the bacteria. Somewhat different patterns of susceptibility were observed in single species compared to two species biofilms. Thus, the results of this study indicate that the susceptibility of bacteria to antimicrobial agents is influenced by biofilm growth as well as by bacterial interactions in mixed bacterial communities.

Key words: biofilm, antimicrobial susceptibility, chlorhexidine, Streptococcus sanquis, Actinomyces naeslundii

INTRODUCTION

In the oral cavity dental plaque bacteria grow in biofilms on the tooth surfaces. Biofilm growth is known from environmental and medical microbiology, where the majority of bacteria in aqueous environments are found attached to surfaces (Costerton et al., 1995). The biofilm mode of growth where bacteria lie embedded in an extracellular matrix has been shown to offer the bacteria protection against different kinds of antimicrobial agents such as antibiotics, antiseptics and factors of the immune system (Costerton et al., 1987). The susceptibility of oral bacteria growing in biofilms has also been shown to decrease compared to their planktonic counterparts, though information on the subject is still not extensive (Wilson, 1996). In a previous study we established a flow model for formation of oral biofilms in vitro and found that the susceptibility of Streptococcus sanquis to different antimicrobial agents decreased considerably when it was grown in biofilms compared to in aqueous suspensions (Larsen and Fiehn, 1995, Larsen and Fiehn, 1996). In the present study a mixed biofilm comprising two major supragingival plaque bacteria, S. sanguis and Actinomyces naeslundii, was established in order to determine the susceptibility of mixed biofilms to chlorhexidine, the most commonly used antiseptic to control dental plaque. Further, the study compared the susceptibility of single species and two species biofilms to chlorhexidine in order to examine possible influences of ecological interactions taking place in a mixed bacterial community that might affect the susceptibility of the bacteria.

MATERIALS AND METHODS

Establishment of one and two species biofilms

Single species biofilms: Biofilms of either Streptococcus sanquis ATCC 10556 or Actinomyces naeslundii (formerly A. viscosus) ATCC 15987 were established on 0.5 cm^2 silicone disks in a modified Robbins device (MRD) as previously described (Larsen and Fiehn, 1995). In brief, a log-phase culture in brain heart infusion (BHI, Difco) with 1% sucrose was pumped through a MRD at 37 C at a rate of 40 ml/h for 48 h. The number of colony forming units in the biofilm was determined after rinsing the disks with 5 ml sterile saline to remove loosely adherent cells and after dispersion of the biofilm cells by vortex mixing for 60 s and low-output ultrasonication for 10 min. 10-fold serial dilutions in sterile saline were plated on blood agar plates (Difco) with 1% sucrose. The number of colony forming units (cfu)/cm^2 was counted after aerobic incubation at 37 C for 48 h. Biofilms of S. sanguis and A. naeslundii were established 13 and 10 times, respectively. Every time the number of colony forming units on 10 or 20 disks was determined. Further, biofilms with each of the species were established three times in the above culture medium supplemented with 0.1% mucin (Sigma) to evaluate any changes induced by the growth medium subsequently used for formation of mixed biofilms.

Bacterial mixture in batch culture: A 2% inoculum, corresponding to the concentration used for biofilm formation, of 24 h cultures of S. sanguis and A. naeslundii was inoculated either alone or in mixture into flasks with BHI or BHI with 1% sucrose and 0.1% mucin (the same medium was used for single as well as mixed cultures throughout each experiment). The cultures were incubated aerobically at 37 C for 72 h, and the cell number of each species was determined after 24, 48 and 72 h. The total cell number was counted in a Petroff Hauser counting chamber, and the viable count was determined by plating on blood agar plates and counting after aerobic incubation at 37 C for 48 h. Further, the cultures were Gram stained for microscopical examination. To enhance growth of A. naeslundii this species was reinoculated in the mixture after 24 h in three of the 8 determinations. Further, pH measurements were performed every 24 h in order to determine metabolic processes possibly affecting the co-existence of the bacteria.

Mixed biofilms: Two species biofilms were established as described above in BHI with 1% sucrose and 0.1 % mucin. S. sanguis and A. naeslundii were grown in separate flasks, and their outlets were connected just before entering the MRD, supplying equal volumes of each species at a total rate of 40 ml/h. The number of cfu/cm^2 of each species was determined after 24, 48 and 72 h of biofilm formation, respectively. Two species biofilms were established 7 times, determining the number of colony forming units on 7 or 10 disks every time.

Effect of chlorhexidine gluconate on S. sanguis and A. naeslundii biofilms

MIC determination: The minimal inhibitory concentration for S. sanguis and A. naeslundii to chlorhexidine gluconate (Nomeco) was determined using the broth dilution method described by Ericsson and Sherris (1971).

Exposure of biofilms to chlorhexidine: Biofilms of S. sanguis and A. naeslundii were established as described above. Single species biofilms were grown in BHI with 1% sucrose for 48 h, while two species biofilms were grown in BHI with 1% sucrose and 0.1% mucin for 24 h, as these growth conditions yielded the highest cell number in the two kinds of biofilms (Larsen and Fiehn, 1995). Subsequently all biofilms were exposed to chlorhexidine gluconate in the growth substrate for 48 h, single species biofilms at concentrations of 0, 16, 50 and 100 mg/ml and two species biofilms at concentrations of 0, 10, 16, 50, 100, 500, 1000 and 2000 mg/ml. Further, two species biofilms were exposed to 1000 mg/ml chlorhexidine in 10 min pulses (7 ml) either 2 or 6 times a day for 48 h. The cfu/cm^2 of each species in the biofilms was determined before introduction of chlorhexidine (time 0) and after 4, 6, 24, 30 and 48 h. Biofilms were exposed to each concentration of chlorhexidine 3-6 times. Every time the number of colony forming units on 3 disks was determined at each time interval.

RESULTS

Establishment of one and two species biofilms

The median cell number in single species biofilms grown in BHI with 1% sucrose for 48 h was 5.74 log10 cfu/cm^2 for S. sanguis and 5.50 log10 cfu/cm^2 for A. naeslundii, respectively. When 0.1% mucin was added to the growth medium the corresponding numbers were 5.83 log10 for S. sanguis and 5.89 log10 for A. naeslundii. Extending biofilm formation from 48 to 72 h did not increase these numbers (data not shown).

The total number of S. sanguis and A. naeslundii grown in separate flasks increased approximately 2 log10 steps to 10^8-10^9 cells/ml, corresponding to the cell number in the 24 h cultures used for inoculation. When inoculated in mixture, however, only S. sanguis attained this number, while the number of A. naeslundii remained at about 10^6-10^7 cells/ml, and the cells showed microscopical signs of degeneration after 48 h and 72 h. Viable counting confirmed that A. naeslundii did not survive in mixture with S. sanguis. Reinoculation of A. naeslundii in the mixture after 24 h did not enhance growth of A. naeslundii, nor did growth in BHI without sucrose, causing an increase in the average pH in the culture medium from 5.0 to 5.8 after 24 h and from 5.3 to 6.4 after 48 h. Thus, it was not possible to grow the two species in the same culture. Accordingly the MRDs for mixed biofilms were supplied with the two species from separate flasks. (Table 1) The highest number of both species was obtained after 24 h.

Table 1. Median (and range) log10 cfu/cm^2 of S. sanguis and A. naeslundii in mixed biofilms

-------------------------------------------------------
                     24 h        48 h          72 h 

S. sanguis          6.43          5.02         4.70     
           
                 (5.21-7.42)   (4.03-6.91)   (4.02-5.81) 

A. naeslundii       5.90           5.95        4.64

                 (4.90-6.94)   (3.92-6.28)   (3.91-5.90)
-------------------------------------------------------

Effect of chlorhexidine gluconate on S. sanguis and A. naeslundii biofilms

The MIC of chlorhexidine gluconate for S. sanguis and A. naeslundii was 16 mg/ml and 4 mg/ml, respectively.

The effect of chlorhexidine on S. sanguis and A. naeslundii biofilms is shown in Figs. 1-7. While the number of bacteria in A. naeslundii biofilms and two species biofilms had reached their maximal level at time 0, the number of S. sanguis in single species biofilms increased about 4 log10 steps during the first 24 h after addition of bacteria-free substrate and then stabilized. When chlorhexidine was added the growth of S. sanguis was arrested, although a marked reduction reaching 0 cfu/cm^2 after 48 h was only obtained at 100 mg/ml (Fig. 1)

Figure 1: Reduction in median log10 cfu/cm2 for established S. sanguis single species biofilms exposed to 0, 16, 50 and 100 mg/ml chlorhexidine for 48 h).

Figure 2: Reduction in median log10 cfu/cm^2 for established A. naeslundii single species biofilms exposed to 0, 16, 50 and 100 mg/ml chlorhexidine for 48 h).

In two species biofilms perfused with BHI without chlorhexidine the number of S. sanguis remained at the initial level throughout the experimental period, while A. naeslundii only maintained its original cell number for 24 h and then rapidly decreased to 0 after 48 h. When exposed to chlorhexidine at 10 or 16 mg/ml both species remained at their initial level. At 50 mg/ml and concentrations above this, a gradual reduction of both species was seen with a progressive decline at higher concentrations and elimination of viable bacteria in <30 h at 500 mg/ml and higher concentrations (Figs. 3 - 6).

Figures 3-6. Reduction in median log10 cfu/cm^2 for established S. sanguis and A. naeslundii mixed biofilms exposed to 0 (Figure 3), 10 and 16 (Figure 4), 50 and 100 (Figure 5) and 500, 1000 and 2000 (Figure 6) mg/ml chlorhexidine for 48 h.

Figure 7: Reduction in median log10 cfu/cm^2 for established S. sanguis and A. naeslundii mixed biofilms exposed to 1000 mg/ml chlorhexidine in pulses 2 or 6 times a day for 48 h.

DISCUSSION

In the present study S. sanguis and A. naeslundii could not co-exist in batch culture as A. naeslundii was suppressed, even when reinoculated after 24 h. When a MRD was supplied with the two species from separate culture flasks, however, mixed biofilms of almost equal cell numbers of both species were established. Interestingly maximal cell numbers of both S. sanguis and A. naeslundii in the mixed biofilm were attained after 24 h while each of the bacteria required 48 h for formation of single species biofilms of comparable cell numbers (Larsen and Fiehn, 1995), indicating that the two species are of mutual benefit to each other during colonization. One possible factor accounting for this may be coadhesion of the cells forming the biofilm due to the specific co-aggregation between strains of S. sanguis and A. naeslundii/A. viscosus described by Kolenbrander (1988). (For many years A. naeslundii and A. viscosus, though very similar, have been regarded as two species. Though they are now considered as belonging to the same species the designation used by the authors in articles referred to below will be followed.) Thus, a recent study showed that deposition of S. sanguis on hydroxyapatite surfaces in a parallel plate flow chamber increased in areas coated with Actinomyces species, but only when co-aggregating bacterial pairs were investigated (Bos et al., 1996).

When bacteria were continuously supplied to the established two species biofilm the number of S. sanguis and A. naeslundii remained at the same level though a minor reduction of both species was observed (Table 1). When, on the other hand, the 24 h biofilm was only supplied with the growth medium S. sanguis maintained at its initial cell number, while A. naeslundii was suppressed to below detectable levels after 48 h (Fig. 3). This occurred even though the growth medium included mucin which has been reported to promote the growth of Actinomyces species in complex communities (Glenister et al., 1988). The same displacement of A. viscosus by S. sanguis in a two species biofilm in an artificial mouth model has previously been described by Ahmed and Russell (1978). They found that while A. viscosus alone formed a "heavy plaque" this organism was gradually replaced by S. sanguis when grown in mixture, and eventually eliminated after 96 to 102 h. Similarly, in another "model mouth" A. viscosus obtained a much smaller cell number in a 3- or 4-species biofilm, than when grown on its own (Donoghue and Perrons, 1988). In more diverse biofilms (5-9 species) established in different flow models A. viscosus only accounted for a very small proportion of the total flora (<<1%), and decreasing cell numbers of A. viscosus were also observed after a maximum had been reached (Herles et al., 1994, Kinniment et al., 1996).

Several factors may be involved in the suppression of A. naeslundii by streptococci, including production of inhibitory substances such as bacteriocins, H2O2 and organic acids lowering the environmental pH (Marsh, 1989). Studies on mixed microbial communities including biofilms have shown that addition of glucose to the growth medium causes changes in the microbial composition due to the low pH generated from carbohydrate metabolism (Bradshaw et al., 1989, Bradshaw et al., 1996). This may explain why A. naeslundii/A. viscosus was eventually eliminated from two species biofilms in the present study as well as in the study by Ahmed and Russell (1978), as the biofilms were grown in sucrose-supplemented medium (1%) and no pH control was performed. In the present study pH-measurements in the mixed batch cultures revealed that pH decreased to averagely 5.0 and 5.3 after 24 and 48 h, respectively. Some of the mixed batch culture studies were, however, performed in a non-sucrose containing medium yielding a higher average pH (5.8 and 6.4 after 24 and 48 h, respectively) and still A. naeslundii could not be recovered, indicating that factors other than pH may also be involved in the suppression of A. naeslundii. Thus, several strains of S. sanguis including ATCC 10556 used in the present study have been shown to produce H2O2 capable of inhibiting the growth of other bacterial species in vitro (Holmberg and Hallander, 1973, Donoghue and Tyler, 1975). Also a strain of S. sanguis, though rather inactive as producer of antagonistic substances, was shown specifically to inhibit the growth of A. viscosus, possibly due to a bacteriocin (Weerkamp et al., 1977).

Interestingly, when the two species biofilms were exposed to chlorhexidine at concentrations too low to arrest growth of the bacteria (10 and 16 mg/ml), A. naeslundii was not eliminated from the biofilm, but maintained its initial cell number at the same level as S. sanguis. Whether this observation could be related to some effect of chlorhexidine on the production of antagonistic substances or the carbohydrate metabolism of S. sanguis, possibly affecting the co-existence of the two bacteria as mentioned above, or whether it was due to some protective effect more directly related to A. naeslundii is not known. However, in some manner the effect seemed to be dependent on a continuous or at least prolonged presence of chlorhexidine in the environment, as it was only observed after exposure of biofilms to chlorhexidine continuously (at low concentrations) or in pulses six times a day (at high concentrations), while pulses two times a day did not protect A. naeslundii but rather accelerated its elimination. Though this rapid elimination of A. naeslundii is in accordance with the results of 0.125% (~1250 mg/ml) chlorhexidine pulsing twice a day in a 9 species biofilm (Kinniment et al., 1996), the maintenance of the cell number of S. sanguis in the present study and the only gradual reduction of both species after pulses 6 times a day may indicate that A. naeslundii rather than being suppressed by chlorhexidine was suppressed by S. sanguis by some bacterial interaction.

The MIC of chlorhexidine to S. sanguis and A. naeslundii found in the present study was in accordance with previous reported MIC-values (Emilson, 1977, Baker et al., 1987). When growing in biofilms resistance of the bacteria increased considerably which is also consistent with previous studies. Both S. sanguis and A. viscosus grown on membrane filters were found only to be killed by very high concentrations of chlorhexidine (Millward and Wilson, 1989; Caufield et al., 1987). Further, the viable count of S. sanguis biofilms established in a constant depth film fermenter under flow conditions was only reduced 2-3 log10 steps after exposure to 0.2% chlorhexidine for up to 4 h (Wilson et al., 1996). This decrease is almost identical to the reduction in cell numbers found in the present study when two species biofilms were exposed to 0.2% (~2000 mg/ml) chlorhexidine for 4 h (Fig. 6). At the lower concentrations of chlorhexidine, however, somewhat different patterns of susceptibility were observed in single species compared to two species biofilms in the present study, indicating the importance of ecological interactions occurring in mixed bacterial communities. In mixed biofilms the cell number of the two species decreased almost similarly, while bacteria in single species biofilms sometimes followed a different and not always the same pattern. Thus, A. naeslundii was eliminated from mixed biofilms by 100 mg/ml of chlorhexidine but was only partly reduced by the same concentration when growing in single species biofilms. Further, in single species biofilms the growth of S. sanguis was arrested by 16 mg/ml of chlorhexidine corresponding to its MIC-value, while the growth of A. naeslundii was not affected by the same concentration though it was equivalent to four times its MIC. These results further emphasize the unreliability of planktonic MIC-determinations for predicting the effect of antimicrobials on bacteria growing in biofilms (Wilson, 1996). Further, they indicate that susceptibility testing of bacteria in single species biofilms should also be interpreted with some caution as bacteria in mixed communities like dental plaque may react differently due to bacterial interactions.

ACKNOWLEDGEMENTS

The excellent technical assistance of laboratory technicians Trine Lemvigh, Tina Friis Mikkelssen and Marie Therp is greatly appreciated. The study was supported financially by the Danish Medical Research Council and by the Danish Dental Association.

REFERENCES

Ahmed, F.I.K. and Russell, C. 1978. Plaque formation in vitro by Actinomyces naeslundii in the presence of Streptococcus sanquis or Streptococcus mutans. Microbios 23: 93-98.

Baker, P.J., Coburn, R.A., Genco, R.J. and Evans, R.T. 1987. Structural determinants of activity of chlorhexidine. J. Dent. Res. 66: 1099-1106.

Bos, R., van der Mei, H.C. and Busscher, H.J. 1996. Co-adhesion of oral microbial pairs under flow in the presence of saliva and lactose. J. Dent. Res. 75: 809-815.

Bradshaw, D.J., McKee, A.S. and Marsh, P.D. 1989. Effects of carbohydrate pulses and pH on population shifts within oral microbial communities in vitro. J. Dent. Res. 68:1298-1302.

Bradshaw, D.J., Marsh, P.D., Schilling, K.M. and Cummins, D. 1996. A modified chemostat system to study the ecology of oral biofilms. J. Appl. Bacteriol. 80: 124-130.

Caufield, P.W., Allen, D.N. and Childers, N.K. 1987. In vitro susceptibilities of suspected periodontopathic anaerobes as determined by membrane transfer assay. Antimicrob. Agents Chemother. 31: 1989-1993.

Costerton, J.W., Cheng, K.-J., Geesey, G.G., Ladd, T.I., Nickel, J.C., Dasgupta, M. and Marrie, T.J. 1987. Ann. Rev. Microbiol. 41: 435-464.

Costerton, J.W., Lewandowski, Z., Caldwell, D.E., Korber, D.R. and Lappin-Scott, H.M. 1995. Microbial biofilms. Ann. Rev. Microbiol. 49: 711-745.

Donoghue, H.D. and Perrons, C.J. 1988. Establishment of defined mixed bacterial plaques on teeth in a laboratory microcosm (model mouth). Microb. Ecol. Health Dis. 1: 193-200.

Donoghue, H.D. and Tyler, J.E. 1975. Antagonisms amongst streptococci isolated from the human oral cavity. Archs. Oral Biol. 20: 381-387.

Emilson, C.G. 1977. Susceptibility of various microorganisms to chlorhexidine. Scand. J. Dent. Res. 85: 255-265.

Ericsson, H.M. and Sherris, J.C. 1971. Antibiotic sensitivity testing. Report of an international collaborative study. Acta path. microbiol. scand. Sect. B (Supppl.) 217:65-67.

Glenister, D.A., Salamon, K.E., Smith, K., Beighton, D. and Keevil, C.W. 1988. Enhanced growth of complex communities of dental plaque bacteria in mucin limited continuous culture. Microb. Ecol. Health Dis. 1: 31-38.

Herles, S., Olsen, S., Afflitto, J. and Gaffar, A. 1994. Chemostat flow cell system: An in vitro model for the evaluation of antiplaque agents. J. Dent. Res. 73:1748-1755.

Holmberg, K. and Hallander, H.O. 1973. Production of bactericidal concentrations of hydrogen peroxide by Streptococcus sanquis. Arch. oral Biol. 18: 423-434.

Kinniment, S.L., Wimpenny, J.W.T., Adams, D. and Marsh. P.D. 1996. Development of a steady-state oral microbial biofilm community using the constant-depth film fermenter. Microbiology 142: 631-638.

Kinniment, S.L., Wimpenny, J.W.T., Adams, D. and Marsh. P.D. 1996. The effect of chlorhexidine on defined, mixed culture oral biofilms grown in a novel model system. J. Appl. Bacteriol. 81: 120-125.

Kolenbrander, P.E. 1988. Intergeneric coaggregation among human oral bacteria and ecology of dental plaque. Ann. Rev. Microbiol. 42: 627-656.

Larsen, T. and Fiehn, N.-E. 1995 Development of a flow method for susceptibility testing of oral biofilms in vitro. APMIS 103: 339-344.

Larsen, T. and Fiehn, N.-E. 1996. Resistance of Streptococcus sanquis biofilms to antimicrobial agents. APMIS 104: 280-284.

Marsh, P.D. 1989. Host defenses and microbial homeostasis: Role of microbial interactions. J. Dent. Res. 68: 1567-1575.

Millward, T.A. and Wilson, M. 1989. The effect of chlorhexidine on Streptococcus sanquis biofilms. Microbios 58: 155-164.

Weerkamp, A., Vogels, G.D. and Skotnicki, M. 1977. Antagonistic substances produced by streptococci from human dental plaque and their significance in plaque ecology. Caries Res. 11: 245-256.

Wilson, M. 1996. Susceptibility of oral bacterial biofilms to antimicrobial agents. J. Med. Microbiol. 44: 79-87.

Wilson, M., Patel, H. and Fletcher, J. 1996. Susceptibility of biofilms of Streptococcus sanquis to chlorhexidine gluconate and cetylpyridinium chloride. Oral Microbiol. Immunol. 11: 188-192.

Copyright remains with the author.

Published by Bioline Publications.

The following images related to this document are available:

Line drawing images