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In situ Biofilm Formation by Multi-species Oral Bacteria Under Flowing and Anaerobic Conditions
S.K. Filoche,
M. Zhu and
C.D. Wu*
Department of Periodontics, College of Dentistry, University of Illinois at Chicago, 801 S. Paulina Street (MC 859), Chicago, IL 60612, USA;
Correspondence: * corresponding author, chriswu{at}uic.edu
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ABSTRACT
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An understanding of biofilm behavior of periodontopathic bacteria is key to the development of effective oral therapies. We hypothesized that interspecies bacterial aggregates play an important role in anaerobic biofilm establishment and proliferation, and contribute to the survivability of the biofilm against therapeutic agents. The system developed in this study assessed a multi-species (Streptococcus gordonii, Actinobacillus actinomycetemcomitans, and Fusobacterium nucleatum) biofilm formation under anaerobic and flowing conditions with the use of an in situ image analysis system. The biofilm was comprised of a base film of non-aggregated cells and complex interspecies aggregates that formed in the planktonic phase which rapidly colonized the surface, reaching 58 ± 9% and 65 ± 11.8% coverage by 5 and 24 hrs, respectively. Upon SDS (0.1%) treatment of a 24-hour biofilm, substantial detachment (55 ± 14%, P < 0.05) of the aggregates was observed, while the base film bacteria remained attached but non-viable. Rapid re-establishment of the biofilm occurred via attachment of viable planktonic aggregates.
Key Words: multispecies anaerobic biofilm • oral biofilm • in situ biofilm development • flow cell • biofilm re-establishment
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INTRODUCTION
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Dental plaque, a complex and dynamic microbial ecosystem, plays an important role in the formation of caries and periodontal disease. It is one of the most widely studied biofilm systems, yet many aspects regarding its formation, structure, and control remain to be elucidated. Many experimental models have been used to study oral biofilms, and most of these necessitate removal of the biofilm from the system and test surface, which results in its disruption and loss of structural integrity. In situ image analysis techniques offer the advantage of monitoring biofilm development in its undisturbed state (Hall-Stoodley et al., 1999). However, in situ studies examining anaerobic multi-species oral biofilms have been limited (Hansen et al., 2000).
We hypothesized that interspecies bacterial aggregates play an important role in anaerobic biofilm establishment and proliferation, and contribute to the survivability of the biofilm against therapeutic agents. In this study, we examined bacterial interactions and biofilm formation, under anaerobic and flowing conditions, of bacteria associated with colonization succession and periodontopathic plaques, namely, Streptococcus gordonii, Fusobacterium nucleatum, and Actinobacillus actinomycetemcomitans (Kolenbrander et al., 1990, 2002), using an in situ image analysis system. Planktonic and biofilm aggregation states that may influence antimicrobial treatment, specifically the detergent sodium dodecyl sulphate (SDS), present in many oral hygiene products, are also reported.
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MATERIALS & METHODS
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Growth Conditions
S. gordonii, F. nucleatum, and A. actinomycetemcomitans were chosen because of their proposed role in aggregation, colonization succession (Kolenbrander et al., 1990, 2002), pathogenicity, and distinguishable microscopic morphologies. Test bacteria were obtained from the culture collection at the College of Dentistry, University of Illinois-Chicago. S. gordonii and A. actinomycetemcomitans were grown in Brain Heart Infusion broth (Difco, Becton-Dickinson and Co., Sparks, MD, USA). F. nucleatum was grown in Schaedler Broth (Oxoid Ltd., Basingstoke, Hampshire, UK) in an anaerobic growth chamber (10% H2, 5% CO2, and 85% N2, Forma Scientific, Marietta, OH, USA).
In vitro Co-aggregation
In vitro co-aggregation between bacterial pairs F. nucleatum and S. gordonii (Fn-Sg) and F. nucleatum and A. actinomycetemcomitans (Fn-Aa) was assessed visually and assigned a score from 0 (homogeneous suspension) to 4 (clear suspension and large settled cell aggregates) (Cisar et al., 1979).
In situ Biofilm Reactor System
A flat plate reactor (FPR; BioSurface Technologies Corp., Bozeman, MT, USA) fitted with a glass coupon (1.25 cm2) pre-conditioned for 1 hr at 37°C with artificial saliva (Landa et al., 1997; Pratten et al., 1998) was connected to a glass culture vessel via sterile tubing in a recirculating batch-culture system. The culture vessel containing 700 mL of Schaedler broth (Oxoid Ltd., Basingstoke, Hampshire, UK) was inoculated with 1 mL overnight cultures of S. gordonii, F. nucleatum, and A. actinomycetemcomitans (total concentration of 1.4 x 106 CFU/mL) and maintained at 37 ± 1°C in a water bath. Flow of the culture medium to the FPR commenced 1 hr after inoculation at 30 mL/hr, approximating human resting salivary flow rate (Lamb et al., 1991; Pratten et al., 1998). We maintained anaerobic growth conditions by supplying nitrogen gas (O2 < 0.5 ppm, BOC gases, Inc., Avondale, IL, USA) to the system. An oxygen indicator strip (BBL Gas pak, Becton-Dickinson and Co., Sparks, MD, USA) was fitted inside the culture vessel in the headspace above the medium. Planktonic growth of test bacteria in this system was compared with growth in an anaerobic chamber by determination of total viable CFU/mL after 24 hrs. Cell aggregates in the culture medium were assessed microscopically at 1 and 24 hrs after inoculation.
Image Analysis of Biofilm Development
In situ biofilm growth and development was monitored by image analysis techniques. A microscope (Leica DMRE, Wetzlar, Germany) was fitted with a digital camera (Optronics, Buffalo Grove, IL, USA), and images were processed and analyzed (Image-Pro-Plus, version 4.5, Media Cybernetics, Silver Spring, MD, USA). Surface area coverage measurements were calculated from 30 images, taken with a long-distance 40x objective, which provided clear distinction of single cells and an indication of overall biofilm formation. Images were captured from different areas of the FPR. An appropriate manual threshold setting (usually a 130-gray-level cut-off resulted in clearly defined boundaries, where black is 255 and white is 0) was applied to each captured image so that the surrounding background was eliminated from the calculations, and, if necessary, incidental artifacts removed, and the areas of interest quantified. The mean area covered by either individual or cell aggregates (a cluster of cells consisting of one or more species), and the dimensions of individual and cell aggregates were calculated.
We evaluated overall biofilm development at lower magnification (10x objective) by measuring interstitial spacing (distance between micro-colonies), corroborated with surface plot profiles generated from the same image. We calculated the surface plot profiles by placing a horizontal reference line through the center of the image and calculating the pixel intensity along the reference line; the resultant plot gave a cross-sectional depiction of biofilm coverage. From these plots, we could estimate the distance between the micro-colonies (µm) as well as biomass accumulation, since the greater the accumulation, the darker the intensity. Biofilm morphology during development was assessed at various magnifications.
Viability of the biofilm and planktonic cells was estimated with the use of the fluorescent Live/Dead® BacLightTM stain (Molecular Probes, Leiden, Netherlands). The glass coupon with attached biofilm was removed from the FPR, stained according to the manufacturer’s instructions (Haughland, 2002), and examined under oil immersion (x100 objective). Bacteria with intact membranes stained green and damaged membranes stained red, indicative of depleted viability or non-viable cells, although false-positive results can occur (Haughland, 2002).
Effect of Sodium Dodecyl Sulphate (SDS) on Multi-species Biofilm
After 24 hrs of biofilm growth, flow was stopped, and 0.1% (v/v) SDS was administered to the system through a syringe via an injection port adjacent to the inlet of the FPR. When the SDS reached the outlet of the FPR, it was vigorously mixed with the biofilm for 30 sec by a repeated drawing-plunging motion of the syringe; approximately 2 mL of SDS was carried over to the reservoir. Flow was resumed, and the effect of SDS on biofilm development was evaluated for 5 hrs. Analyses included surface area coverage by individual and cell aggregates, and micro-colony distance mapping, as described above. Sterile water was administered in the same way to a replicate biofilm as the control. The amount of biofilm removed due to change in pressure during the administration of SDS was adjusted (LossSDS - Losswater control = Adjusted lossSDS).
Statistical Analysis
All experiments were independently repeated a minimum of 3 times, and the mean ± SD is calculated. Descriptive statistics were calculated (Microsoft® Excel 2000, version 9) and the results analyzed for significance by one-way analysis of variance and the post hoc Tukey’s test (ANOVA, P < 0.05; MINITABTM 2002, Version 13.32).
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RESULTS
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Growth and Development of Multi-species Biofilm under Anaerobic Conditions
The reactor system developed in this study supported the growth of a multi-species biofilm under anaerobic conditions created by nitrogen supplied to the circulating unit. Planktonic growth of the mixed culture in the reactor system (total CFU/mL 9 ± 0.1 Log10) was comparable (P > 0.05) with that obtained in the anaerobic chamber after 24 hrs (total CFU/mL 9 ± 0.05 Log10).
Examination of the planktonic cells 1 hr after inoculation showed that considerable interspecies associations, among all 3 bacteria, had occurred in the form of well-organized cell aggregates (Fig. 1a ). Biofilm formation commenced with the attachment of planktonic aggregates to the pre-conditioned glass surface minutes after flow was initiated. The aggregates attached to the glass surface at one end with a free-end that oscillated in the direction of flow (Figs. 1b, 1c). Concomitant development of a base film, which consisted of individual bacteria, was observed (Fig. 1d). No apparent hierarchy in the order of species attachment was observed.
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Figure 1. Photomicrographs illustrating the stages of biofilm development and the planktonic and biofilm interspecies aggregates. (a) Interspecies planktonic aggregate at 1 hr, showing a single F. nucleatum (Fn) cell in the center with groups of A. actinomycetemcomitans (Aa) and S. gordonii (Sg) cells on either side. (b) Representative light microscopy image of biofilm development at 1 hr and (c) at 3 hrs. Darker areas represent greater biofilm accumulation; (m) indicates micro-colonies comprised of the aggregates; (i) indicates the interstitial spacing (void fraction) between the micro-colonies of the biofilm; (fe) indicates the free-end structure which oscillated in the direction of flow and also trapped passing aggregates. (d) Biofilm aggregates (ba) at 3 hrs with the base film cells (bf). (e) Biofilm aggregate with the structures that were observed bridging adjacent aggregates together, such as the group of distinctly organized A. actinomycetemcomitans cells (Aa) and individual F. nucleatum cells (Fn). By varying the focal plane, we could visualize different layers of the aggregate. However, it was not always possible to focus the entire cluster, due to thickness variability and continuous flow of the culture medium. The unfocused area represents cells protruding into and oscillating in the culture medium. The dark deposits are medium artifacts. (f) Interspecies planktonic aggregate at 24 hrs showing all 3 bacteria surrounded by an amorphous cohesive matrix (m). The aggregates were stained with Live/Dead® BacLightTM; however, no inferences in the viability of the cells were made for these samples, since organization of the aggregates was better visualized after the sample fluorescence had faded.
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By 3 hrs, linkages among the attached aggregates were noted through groups of A. actinomycetemcomitans cells, individual F. nucleatum cells, and chains of S. gordonii (Fig. 1e ). Examination of the planktonic aggregates at 24 hrs (mean width 130 µm ± 56.2 and mean length 199 µm ± 42.5) revealed that an amorphous, cohesive matrix (Fig. 1f) surrounded the cells. Morphologically, these planktonic aggregates were analogous to a planktonic biofilm.
Considerable growth and attachment were evident between 1 and 3 hrs, with an 87 ± 10.1% increase in the mean total area covered by aggregated cells and an 71 ± 9.7% increase in that covered by individual cells (Fig. 2). No initial lag phase in attachment was observed (Fig. 2). Between 5 and 24 hrs, there was only a 9 ± 11.4% increase in the mean total area covered by aggregate cells, while there was no detectable change in the area covered by individual cells (Fig. 2). The mean total area coverage by the aggregates increased over time, reaching 65 ± 11.8% at 24 hrs (Fig. 2). The attached aggregates also trapped passing planktonic aggregates, resulting in the increase in biofilm thickness and surface area coverage. Parallel reductions in interstitial spacing were evident (Figs. 3a, 3b).
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Figure 2. Surface area coverage by aggregated and individual cells of a multi-species biofilm. SDS (0.1%) was added at the time indicated by the arrow. For each time point in each replicate experiment, 30 images were taken, and the mean area covered by aggregates and the mean area covered by individual cells were calculated. Error bars show standard deviations from 3 independent replicates and illustrate the inherent heterogeneous nature of biofilm formation.
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Figure 3. Representative surface plot profiles depicting biofilm accumulation over time and the effect of SDS. The profiles were taken from reference lines applied to the captured image. The vertical axis represents the degree of biofilm accumulation (based on pixel intensity). The horizontal axis represents distance along the horizontal profile. The width of the peaks indicates the width of the microcolonies, and the distance between the edges of the peaks indicates the distance between adjacent microcolonies (µm). The arrow indicates the distance between the micro-colonies (mc). Biofilm at (a) 3 hrs, (b) 24 hrs, (c) 1 hr after SDS application, and (d) 5 hrs after SDS application.
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Staining of a 24-hour biofilm with Live/Dead® BacLightTM demonstrated that the base film consisted primarily of viable S. gordonii. The planktonic and biofilm aggregates consisted mainly of viable cells of all 3 bacteria. Occasional non-viable cells were noted toward the center.
A. actinomycetemcomitans and S. gordonii demonstrated different in vitro co-aggregation affinity with F. nucleatum: A. actinomycetemcomitans and F. nucleatum 2 and S. gordonii and F. nucleatum 4 (on a 0–4 scale).
The Effect of SDS on Multi-species Biofilm
The addition of 0.1% SDS to a 24-hour biofilm immediately resulted in significant removal of the attached aggregates from the base film (adjusted: 55 ± 14.3%, P < 0.05, Fig. 2). The rapid clearance of aggregates gave rise to a patchy distribution, and the clearance occurred through the detachment of whole aggregates, not dissolution of the aggregates. SDS did not appear to affect the subsequent attachment of individual cells in the voids created in the base film, and a 69 ± 8.8% increase in the mean area covered was detected as a result of the removal of the aggregates (P > 0.05). Re-establishment of the biofilm was apparent 5 hrs after SDS application and was evident by the re-attachment of planktonic aggregates, reaching 67 ± 15.6% mean area coverage, representing a 95 ± 11.3% increase in surface area covered (Figs. 2, 3d). The addition of water to the system removed approximately 30 ± 9.3% of attached aggregates from the biofilm. Examination of the Live/Dead® BacLightTM-stained, SDS-treated biofilm revealed a base film consisting predominantly of non-viable S. gordonii, while the multi-species planktonic and biofilm aggregates were viable.
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DISCUSSION
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Using the in situ image analysis system reported in this study, we identified planktonic and biofilm aggregation states of S. gordonii, A. actinomycetemcomitans, and F. nucleatum, which, with destructive techniques, may have gone undetected. The constant supply of nitrogen into the FPR system maintained optimal anaerobic conditions for the growth of both planktonic and biofilm cultures. Analysis of the data obtained indicated that interspecies bacterial aggregates played an important role in anaerobic biofilm establishment and proliferation, and contributed to the survivability of the biofilm against antimicrobial agents.
The rapid attachment of the multi-species planktonic aggregates facilitated biofilm accumulation, especially in the early stages (Fig. 1), as hypothesized by Stoodley et al.(2002), indicating that this process may play an important role in plaque formation. These interspecies aggregates, surrounded by a cohesive matrix, formed a complex structure (Fig. 1f) analogous to the ‘planktonic biofilms’ described by Flemming and Wingender (2001). In the human oral cavity, optimal periods for attachment could be considered as transient, and mechanisms for surviving ambient conditions are paramount for biofilm survival (Lewandowski and Walser, 1991). Interspecies aggregates may function as a synergistic micro-consortium (Cook and Costerton, 1998; Putnins et al., 2001; Sutherland, 2001) that offers protection from shear stress and desiccation and thus contributes to biofilm accumulation (Sadasivan and Neyra, 1985; Kolenbrander et al., 1990; Saunders and Greenman, 2000; Handley et al., 2001; Stoodley et al., 2001; Rickard et al., 2003).
Co-aggregation between oral bacteria is a specific phenomenon and has been routinely evaluated in vitro with the use of non-actively-metabolizing washed cells of overnight cultures (Kolenbrander et al., 1990). However, in this study, interspecies aggregates observed in situ developed within 1 hr under nutrient-rich and anaerobic conditions among actively growing cultures of S. gordonii, A. actinomycetemcomitans, and F. nucleatum (Fig. 1a). These associations were more frequently observed among all 3 bacteria rather than pair-wise. It was not possible to ascertain whether the inter-bacterial associations formed simultaneously among the bacteria or through a hierarchical process, although no specific order in colonization was observed (Kolenbrander et al., 1990). These findings also demonstrated that a moderate in vitro co-aggregation (Fn-Aa, 2) was not indicative of the extensive interspecies associations formed in situ. This discrepancy between the extent of in vitro vs. in situ co-aggregation of oral bacteria has been previously reported and was attributed to assay limits (Guggenheim et al., 2001).
The ability of antimicrobial agents to penetrate biofilm and render the base film non-viable has been considered an important factor in antimicrobial efficacy (Stewart et al. 1998).
Analysis of our data showed that after the addition of 0.1% SDS and the subsequent detachment of the biofilm aggregates, the non-viable base film cells acted as attachment sites for a new biofilm to establish. This suggested that the viability of the base film may not be essential in biofilm re-establishment (Weiger et al., 1999), and that bacterial aggregation states appear to affect antimicrobial susceptibility.
Biofilm detachment has been implicated in the dissemination of infection and contamination of industrial systems (Stoodley et al., 2001). The present study demonstrated that bacterial aggregates readily removed from the base film remained viable after 5 hrs of exposure to 0.1% SDS (Landa et al., 1997; Eginton et al., 1998). These aggregates may re-attach at different sites and facilitate the re-establishment and proliferation of biofilms. Analysis of our data also suggests that effective antimicrobial agents need to possess the ability to dissociate and kill aggregates in a short period of exposure time, to minimize biofilm re-establishment. The effects of planktonic-biofilm aggregates on the efficacy of oral therapeutic agents (Socransky and Haffajee, 2002) and their role in the contamination of dental water units (Putnins et al., 2001) warrant further investigation.
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ACKNOWLEDGMENTS
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This research was supported by the National Institute of Dental and Craniofacial Research, NIH DE13990. The authors thank: the US Army Dental Research Detachment, Great Lakes, IL, USA, for their equipment support; Gemma Husmillo and Patrick Filoche for their technical assistance; and Dr. Paul Stoodley for his consultation regarding hydrodynamics.
Received for publication March 31, 2003.
Revision received August 5, 2004.
Accepted for publication August 25, 2004.
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Journal of Dental Research, Vol. 83, No. 10,
802-806 (2004)
DOI: 10.1177/154405910408301013
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ABSTRACT
INTRODUCTION