AEM
Home Help [Feedback] [For Subscribers] [Archive] [Search] [Contents]
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Yang, Y.
Right arrow Articles by Cummins, D.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Yang, Y.
Right arrow Articles by Cummins, D.
Agricola
Right arrow Articles by Yang, Y.
Right arrow Articles by Cummins, D.
Applied and Environmental Microbiology, October 2006, p. 6734-6742, Vol. 72, No. 10
0099-2240/06/$08.00+0     doi:10.1128/AEM.01013-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

Multiparameter Assessments To Determine the Effects of Sugars and Antimicrobials on a Polymicrobial Oral Biofilm

Ying Yang, Prem K. Sreenivasan,* Ravi Subramanyam, and Diane Cummins

Colgate-Palmolive Company, 909 River Road, Piscataway, New Jersey 08855

Received 1 May 2006/ Accepted 12 July 2006


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Clinical studies indicate relationships between dental plaque, a naturally formed biofilm, and oral diseases. The crucial role of nonmicrobial biofilm constituents in maintaining biofilm structure and biofilm-specific attributes, such as resistance to shear and viscoelasticity, is increasingly recognized. Concurrent analyses of the diverse nonmicrobial biofilm components for multiparameter assessments formed the focus of this investigation. Comparable numbers of Actinomyces viscosus, Streptococcus sanguinis, Streptococcus mutans, Neisseria subflava, and Actinobacillus actinomycetemcomitans cells were seeded into multiple wells of 96-well polystyrene plates for biofilm formation. Quantitative fluorescence and confocal laser scanning microscopy (CLSM) examined the influences of dietary sugars, incubation conditions, ingredients in oral hygiene formulations, and antibiotics on biofilm components. Biofilm extracellular polymeric substances (EPS) were examined with an optimized mixture of fluorescent lectins, with biofilm proteins, lipids, and nucleic acids detected with specific fluorescent stains. Anaerobic incubation of biofilms resulted in significantly more biofilm EPS and extractable carbohydrates than those formed under aerobic conditions (P < 0.05). Sucrose significantly enhanced biofilm EPS in comparison to fructose, galactose, glucose, and lactose (P < 0.05). CLSM demonstrated thicker biofilms under sucrose-replete conditions, along with significant increases in biofilm EPS, proteins, lipids, and nucleic acids, than under conditions of sucrose deficiency (P < 0.05). Agents in oral hygiene formulations (chlorhexidine, ethanol, and sodium lauryl sulfate), a mucolytic agent (N-acetyl-L-cysteine), and antibiotics with different modes of action (amoxicillin, doxycycline, erythromycin, metronidazole, and vancomycin) inhibited biofilm components (P < 0.05). Multiparameter analysis indicated a dose-dependent inhibition of biofilm EPS and protein by chlorhexidine and sodium lauryl sulfate, along with distinctive inhibitory patterns for subinhibitory concentrations of antibiotics. Collectively, these results highlight multiparameter assessments as a broad platform for simultaneous assessment of diverse biofilm components.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Biofilms representing accumulations of microorganisms in a complex matrix have now been reported for diverse environments (3, 10, 12, 13, 25, 27). Characteristics unique to biofilms include decreased susceptibilities to antimicrobial agents and biocides compared to those of planktonic organisms (10, 25). Relationships between biofilms and the etiology of microbial infections (12), including some forms of chronic and recurrent human disease (3), device-related infections, and treatment failures (11), have been the subject of recent investigations. The human mouth, with its diverse niches and environmental changes, is well known for the unrestricted formation of natural microbial biofilms (3, 12, 25). Oral biofilms are found on the tooth as dental plaque, both above and below the gum line, and on the surfaces of the tongue (25).

Clinical oral microbiology has examined the microbial diversity of oral biofilms. Investigations of oral biofilms from subjects stratified on the basis of oral health have examined the relative distributions of microorganisms in health and disease (13, 25). These efforts have been instrumental in elucidating the microorganisms in the diverse niches of the human mouth (11, 13, 25, 28), the microbiology of oral diseases, and therapeutic strategies for their control (11, 25). Analyses of the genes from oral bacteria associated with biofilms have been reported for several organisms (9, 15, 17, 30), with molecular analyses of biofilm morphogenesis and maturation as areas of future research (10, 12).

The analysis of bacteria found in biofilms (12, 13) has formed a significant focus of recent investigations. On the other hand, the nonmicrobial components of biofilms, which include the biofilm matrix, remain relatively unexplored (3, 10, 12, 14, 16, 24, 28). Initial reports indicate the complexity of the biofilm matrix and its role in maintaining biofilm structure. For instance, biofilm matrix polysaccharides comprise a major portion of the biofilm (16), serving as a three-dimensional skeleton (28) along with a number of other functions attributed to the biofilm matrix, such as viscoelastic properties and resistance to shear (3, 14). The inherent dynamic aspects of the biofilm matrix, including the lack of appropriate techniques for analysis (16), are some likely reasons for its incomplete analysis (10, 25). Analyses of the matrix for specific constituents, in addition to their changes over time as related to biofilm morphogenesis and maturation, remain to be established (16). A range of environmental variables, including solute and nutritional components, along with intrinsic factors such as the diversity of microorganisms in the biofilm and their cellular processes, reportedly influence biofilm components (3, 28).

The focus of this investigation was the development of procedures for an examination of the diverse nonmicrobial components of a polymicrobial biofilm comprising several oral bacteria. The overall recognition of the nonmicrobial components as integral elements of biofilms (28) provided the rationale for this investigation. Fluorescent lectins were utilized as probes to examine the extracellular polymeric substances (EPS) of a multispecies oral biofilm. Other nonmicrobial biofilm components were investigated with fluorescent dyes specific for lipids, proteins, and nucleic acids. These procedures facilitate rapid analysis followed by confocal laser scanning microscopy (CLSM). Optimum conditions for reproducible simultaneous assessment of each biofilm component for multiparameter analyses were established. A range of studies determined the influences of different concentrations of common dietary sugars and media and of incubation conditions. Multiparameter assessments examined the influences of ingredients found in oral hygiene formulations, including antimicrobial agents and antibiotics, on biofilm components.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Bacteria and chemicals.
Bacterial strains for biofilm studies included oral bacteria (Actinomyces viscosus ATCC 43146, Streptococcus sanguinis ATCC 10557, Streptococcus mutans ATCC 33402, Neisseria subflava 49275, and Actinobacillus actinomycetemcomitans ATCC 29522) and Pseudomonas aeruginosa 9027. All strains were obtained from American Type Culture Collection (ATCC), Manassas, Va. Bacteriological media were obtained from Becton-Dickinson, Sparks, Md., and prepared in accordance with the manufacturer's recommendations. Trypticase soy broth supplemented with 0.6% yeast extract (TSB-YE) was prepared for routine bacterial growth. Buffers and chemicals, including antibiotics for tests, were reagent grade or better and routinely obtained from Sigma Chemical Company, St. Louis, Mo., unless indicated otherwise. Fluorescent lectins concanavalin A (ConA) and wheat germ agglutinin (WGA) labeled with Alexa fluor 488 were obtained from Molecular Probes, Eugene, OR. Other fluorescent dyes obtained from Molecular Probes included Syto-60 and Syto-84 to stain nucleic acids and Sypro red and Nile red to stain proteins and lipids, respectively. The fluorescent dyes along with excitation and emission filters chosen for analyses are shown in Table 1. Stock solutions of antibiotics (amoxicillin, doxycycline, erythromycin, metronidazole, tetracycline, and vancomycin) were prepared as described previously (6, 19, 21, 22). Other agents obtained from Sigma Chemical Co. included the mucolytic agent N-acteyl-L-cysteine, a 20% aqueous solution of chlorhexidine (CHX), and sodium lauryl sulfate (SLS). A stock solution of 5% SLS was prepared in sterile phosphate-buffered saline (PBS) and filter sterilized (0.2-µm syringe filters; Corning, Acton, MA) prior to use. Stock solutions of aqueous solutions (5 to 20%) of sugars (fructose, galactose, glucose, lactose, and sucrose [Sigma Chemical Co.]) were prepared when required and filtered through a 0.2-µm filter prior to use. TSB-YE was reconstituted aseptically with the appropriate volume of each sugar stock solution as required for each study.


View this table:
[in this window]
[in a new window]
  		TABLE 1. Fluorescent dyes for biofilm assessments

 
Growth of microbial biofilms. (i) Preparation of saliva-coated 96-well plates.
Volunteers for saliva collection were healthy adults (two males and three females; age range, 28 to 48 years) who had no history of prescription medications and antibiotics in the preceding 3 months. Subjects selected for saliva collection were instructed on the study design, with all aspects of the study conducted in accordance with procedures well accepted for human subjects. Selected volunteers were provided a commercially available fluoride dentifrice and a soft-bristled toothbrush for their routine oral hygiene and discontinued the use of all other oral hygiene formulations for 1 week prior to the start of their scheduled saliva collection. Procedures for collection of whole saliva from subjects were done in accordance with previously described procedures (7, 20). In brief, subjects chewed on paraffin wax, and the saliva (20 to 50 ml) collected from each subject was pooled and centrifuged (8,000 x g for 10 min) to obtain clarified saliva and then stored at –20°C prior to use (7, 20, 24). Clarified saliva (0.1 ml) was added to each well of a 96-well plate (Becton Dickinson, Sparks, MD) and incubated overnight at 37°C to coat the wells and form salivary pellicles. Salivary pellicle-coated plates were rinsed twice with PBS (0.1 ml per well) prior to the addition of the microbial mixture described below for biofilm formation.

(ii) Procedures for microbial biofilms.
Oral bacteria (Actinomyces viscosus, Streptococcus sanguinis, Streptococcus mutans, Neisseria subflava, and Actinobacillus actinomycetemcomitans) were routinely cultivated in TSB-YE supplemented with 0.2% sucrose under anaerobic conditions in a BBL GasPak Plus (BBL, Sparks, MD) at 37°C as described previously (6, 9, 17). Overnight cultures were diluted to an optical density at 610 nm of 0.1 ± 0.02 (between 107 and 107.4 CFU/ml for each strain) in fresh TSB-YE supplemented with 0.2% sucrose. Equal volumes of the strains were mixed to obtain a stock mixture. A 0.1-ml volume of this microbial mixture was added to multiple wells of a salivary pellicle-coated 96-well plate. This provided replicate samples of biofilms for each treatment. The bacteria were allowed to adhere to the plates for 3 h at 37°C under anaerobic conditions. Next, the medium in these plates was replaced with 0.2 ml fresh medium (TSB-YE plus 0.2% sucrose). Plates were incubated for 24 h at 37°C under anaerobic conditions for biofilm formation. The medium in these plates was replaced and allowed to incubate for a further 24 h under anaerobic conditions. After 48 h of incubation, each well was rinsed three times with 0.2 ml PBS per wash, and the plates were analyzed as described below.

Procedures for biofilm formation were modified for studies that examined the effects of aerobic incubation, the effects of medium strength, and the effects of different sugars or inhibitory agents on biofilm formation. To determine the effects of different media, biofilms were allowed to form in TSB-YE plus 0.2% sucrose or in medium diluted to concentrations of 50%, 25%, and 12.5% by the addition of sterile water. The effects of sugars on biofilm formation were examined by supplementing TSB-YE to a final concentration of 0.2% with aqueous solutions of sugars filtered through a syringe filter as described above. Medium with the required concentration of each sugar was prepared as needed prior to use. The activities of antimicrobial agents were elucidated following a 30-min exposure of 48-h-old biofilms or following the incorporation of antimicrobials into the medium during the 48-h period of biofilm formation (4, 6).

MICs.
MICs were determined as described by CLSI (formerly NCCLS) (19), with minor modifications. Stock solutions of amoxicillin, doxycycline, erythromycin, metronidazole, vancomycin, and N-acteyl-L-cysteine had concentrations of 1.2, 3.125, 3.125, 2.5, 6.25, and 25 mg/ml, respectively, and were prepared as described previously (19, 22). The agents were serially diluted in 96-well microtiter plates in TSB-YE prior to incubation with bacterial cultures diluted to an optical density at 610 nm of 0.1. MIC results were recorded after 48 h of anaerobic incubation. The MICs for oral bacteria are shown in Table 2. This procedure was also utilized to examine the effects of chlorhexidine and SLS.


View this table:
[in this window]
[in a new window]
  		TABLE 2. MICs of agents

 
Fluorescence assessments of biofilms. (i) Fluorescence quantitation by microplate analysis.
A stock solution (2.5 mg/ml) of each fluorescent lectin (ConA and WGA) prepared in PBS was diluted to a working solution comprising 250 µg/ml of each lectin for tests. Solutions of each lectin ranging from 0 to 250 µg/ml were utilized in tests to identify the optimal concentrations of lectins for examining biofilms. The optimal concentrations of lectins were determined by a response surface analysis that utilized a central composite design and Minitab statistical software (Minitab Inc., State College, PA). The program selected 10 concentrations (µg/ml) of the lectins (ConA/WGA ratios of 8.57:150, 50:50, 50:250, 150:291.42, 150:8.57, 250:50, and 291.42:150) and duplicated the central concentration of the lectin mixture (150 µg/ml [each] of ConA and WGA) for the studies. Each test concentration of lectins was used to measure eight replicate samples of biofilms. The results of the response surface analysis indicated that a quadratic model fit the data well (R2 = 83.9%) and was highly significant (P < 0.001). An optimal concentration of 125 µg/ml of each lectin was chosen as the concentration for further studies.

Fluorescent lectins (50 µl) were added to each well of a 96-well plate and incubated with biofilms or salivary pellicles at room temperature for 30 min to stain. Plates were washed three times with 0.2 ml sterile water per wash. Fluorescence was quantified in a Cytofluor series 4000 (PerSeptive Biosystems, Framingham, MA) plate reader. Excitation and emission settings for experiments are shown in Table 1.

The fluorescent stains used for the assessment of proteins, lipids, and nucleic acids were Sypro red, Nile red, and Syto-84, respectively. Their excitation and emission settings are shown in Table 1. Sypro red was diluted 1:500 in 50 mM Tris-1 mM EDTA, pH 7.5, for staining. Nile red and Syto-84 were diluted in 10 mM Tris-1 mM EDTA, pH 8.0, to obtain final concentrations of 20 µM of Nile red and 10 µM of Syto-84. Diluted stains (50 µl) were added to each well of a 96-well plate and allowed to incubate with biofilms for 30 min at room temperature. Stained biofilms were washed three times with sterile water for fluorescence quantitation. In some instances, biofilms were stained with dual dyes. For these studies, biofilms were initially stained with the first stain (lectins or Syto-84 for 30 min) and then washed three times with sterile water prior to staining with the second stain (Syto-84 or lectins for 30 min).

(ii) Confocal microscopy.
A Carl Zeiss model 410 CLSM (Thornwood, NY) with lasers at 488, 568, and 647 nm was utilized to examine biofilms grown on 96-well plates. Procedures and concentrations of fluorescent dyes for staining biofilms for CLSM with lectins, Sypro red, and Nile red were as indicated previously for studies with the fluorescence microplate reader. Syto-60, a nucleic acid stain, was chosen to comply with filter combinations available for CLSM and diluted in 50 mM Tris-1 mM EDTA, pH 7.5, to a working concentration of 10 µM. For staining, 50 µl of the working solution of Syto-60 was added to each well and incubated for 30 min at room temperature. Syto-60 enabled simultaneous assessments of biofilms stained with two dyes. Lasers for excitation and filter combinations for CLSM are shown in Table 1. Hydrated biofilms in 96-well plates stained with the appropriate dyes (as described for fluorescence microplate analysis) were mounted on a CLSM microplate adapter for xy analysis at a magnification of x20. The xy analysis provided surface coverage of the biofilm. Additional z section analyses, performed by sectioning the biofilm, determined the depth of the biofilm and its topography.

Estimation of biofilm carbohydrates.
Biofilm carbohydrates were determined by the phenol-sulfuric acid method as described previously (5, 27). In brief, deionized water (40 µl) and a 5% phenol solution (40 µl), followed immediately by 95 to 97% sulfuric acid (200 µl), were added to each well of a 96-well plate. Plates were incubated at room temperature for 30 min, and the amount of carbohydrate was estimated by measuring the absorbance at 490 nm in a microplate reader (Bio-Tek ELX800 absorbance microplate reader; Bio-Tek Instruments, Inc., Winooski, VT).

Statistical analyses.
Statistical analyses were conducted with replicate samples by using JMP software (SAS Institute, Cary, NC). The binding of fluorescent lectins to microbial biofilms and salivary pellicles was compared by Student's t test. The effects of different sugars, incubation conditions, and inhibitory agents on biofilms were examined by one-way analysis of variance with post hoc analysis by the Dunnett multiple comparison test, with statistical significance reported for P values of <0.05.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Development of a method for examining biofilm EPS.
Human saliva comprises a range of components, including those derived from the host, that coat oral surfaces and form the salivary pellicle (12, 13, 25). The salivary pellicle provides sites for subsequent growth of the oral microbial biofilm (25). A response surface optimization experimental design determined the optimal concentrations of lectins (ConA and WGA) and mixtures of these lectins for biofilm binding. The design tested lectin concentrations that ranged from 0 to 250 µg/ml for each lectin. Average results from 48 wells (Fig. 1) indicated more lectin binding on biofilms than on the salivary pellicle by t test analysis (P < 0.00001). Decreasing concentrations of lectins resulted in less fluorescence. An optimal response was observed with a lectin mixture comprising 125 µg/ml of each lectin (data not shown), and this mixture was selected for further tests. In control studies, incubation of the selected concentration of lectins (125 µg/ml) in 96-well plates without salivary pellicles and on plates coated with bacteriological medium resulted in low background fluorescence responses (ranging from 20 to 30 fluorescence units). These background fluorescence results were significantly lower than those noted for the salivary pellicle or microbial biofilm (P < 0.001).


Figure 1
View larger version (10K):
[in this window]
[in a new window]
  		FIG. 1. Analysis of salivary pellicles and microbial biofilms of oral bacteria by fluorescent lectins ConA and WGA (125 µg/ml). Results from 48 replicate microtiter wells are shown in each box plot.

 
Effects of medium concentration and microbial growth conditions on biofilm components. (i) Influence of medium concentration and incubation conditions.
A variety of factors, such as sugars, dietary constituents, and environmental conditions, influence the growth of oral microorganisms (13, 25). The influences of these factors were determined in concurrently conducted experiments that examined the effects of incubation and growth conditions on biofilm EPS by fluorescence lectin analysis and biofilm carbohydrate determinations. An additional rationale for these studies was to relate the EPS results with those for biofilm carbohydrates. For these studies, bacteriological medium (TSB-YE plus 0.2% sucrose) was diluted with sterile water to test the effects of four medium concentrations (100%, 50%, 25%, and 12.5%) on biofilms. Biofilms (12 replicates each) were allowed to form at each medium concentration under either aerobic or anaerobic conditions and were analyzed for EPS and biofilm carbohydrates by the phenol-sulfuric acid method (Fig. 2A and B). Anaerobic conditions and undiluted medium resulted in the highest levels of EPS and biofilm carbohydrates in comparison to all other medium concentrations tested (P < 0.05). Biofilms formed at decreasing concentrations of medium resulted in corresponding reductions in EPS and biofilm carbohydrates (P < 0.05). Differences between anaerobic and aerobic incubation conditions were evident in full-strength medium for both EPS and biofilm carbohydrates (P < 0.05). At all medium concentrations tested, more biofilm EPS were observed with biofilms formed under anaerobic conditions than with those formed with aerobic incubation (P < 0.05). Although significant differences in biofilm carbohydrates were observed between the two incubation conditions for undiluted medium (P < 0.05), these differences were insignificant at all other medium concentrations tested.


Figure 2
View larger version (19K):
[in this window]
[in a new window]
  		FIG. 2. Effects of medium concentration and incubation conditions on (A) oral biofilm EPS (determined with fluorescent lectins WGA and ConA) and (B) oral biofilm carbohydrates. The results shown are averages ± standard deviations for 12 replicate microtiter wells of biofilms grown under anaerobic (heavily dotted bars) and aerobic (lightly dotted bars) conditions.

 
Pseudomonas aeruginosa represents the paradigm organism for biofilm studies (3, 12). Therefore, we analyzed lectins and carbohydrates of P. aeruginosa biofilms (quadruplicate samples) grown under aerobic and anaerobic conditions (27) at full strength and decreasing concentrations of medium. Aerobic incubation resulted in more biofilm EPS and carbohydrate than did anaerobic incubation (P < 0.05) in full-strength medium. At lower medium concentrations, corresponding reductions in biofilm EPS and carbohydrate were observed under aerobic conditions (data not shown).

(ii) Effects of sugars.
Common sugars are prevalent in the daily diet, and clinical studies have indicated their influences on the dental plaque biofilm, their cariogenic potential, and their effects on oral health (2, 13, 25). Therefore, the effects of sugars (fructose, galactose, glucose, lactose, and sucrose) at a 0.2% final concentration on biofilm EPS were studied. Results from quadruplicate samples demonstrated a significant increase in lectin binding among biofilms grown in the presence of 0.2% sucrose compared to that in the presence of all other sugars tested (data not shown) (P < 0.05). The other sugars significantly enhanced biofilm EPS in comparison to the control without any sugar (data not shown) (P < 0.05).

Multiparameter analysis of biofilms. (i) Fluorescence analysis of biofilm components.
Multiparameter assessments examined biofilms grown in the presence or absence of 0.2% sucrose. The studies examined several biofilm parameters simultaneously. Results for 12 replicates (Fig. 3A) indicated significantly higher fluorescence responses (>3-fold) for EPS, lipids, nucleic acids, and proteins from biofilms grown in the presence of sucrose than for those from corresponding biofilms grown in the absence of sucrose (P < 0.05). The usefulness of these assays for the simultaneous examination of two biofilm components was investigated further. Figure 3B shows the results for biofilms stained for two components, i.e., EPS and nucleic acids. In this case, biofilms (12 replicates) were initially stained for EPS or nucleic acids and subsequently stained for nucleic acids or EPS, respectively. The results indicate fluorescence from the first stain and demonstrate significantly larger amounts of EPS and nucleic acids in the presence of sucrose (P < 0.05). The ability of sucrose to enhance biofilm components in the dual-stain studies was comparable to that obtained in the single-stain studies. Interestingly, dually stained biofilms grown in the presence of sucrose produced more nucleic acid fluorescence. Other combinations of dyes were not feasible due to the spectral properties of the selected probes. In multiple studies, minimal background fluorescence staining (20 to 30 units) was observed for each of the fluorescent dyes.


Figure 3
View larger version (18K):
[in this window]
[in a new window]
  		FIG. 3. Multiparameter analysis of biofilms grown in the presence or absence of 0.2% sucrose (Suc). (A) Averages ± standard deviations of fluorescence of biofilm EPS, carbohydrates, proteins, and nucleic acids from 12 replicate microtiter wells. (B) Biofilms stained for both EPS and nucleic acids and assessed for either EPS or nucleic acids. Results are averages ± standard deviations for 12 replicates. (C) Assessment of biofilms by the crystal violet procedure. Results are averages ± standard deviations for 24 replicates.

 
A staining procedure with crystal violet finds wide application in the examination of biofilms produced by a variety of bacteria (3). Therefore, studies utilizing the crystal violet procedure (9, 17) evaluated 24 replicates grown in the presence or absence of sucrose (Fig. 3C) in comparison to the fluorescence-based multiparameter approaches described above. As described above, sucrose-replete conditions resulted in significantly more (2.5-fold) crystal violet staining than did sucrose deficiency (P < 0.05). Therefore, the results indicate that the multiparameter approaches corroborate results obtained by the widely used crystal violet procedure. However, in the presence of sucrose, the fluorescence-based multiparameter analyses demonstrated a >3-fold increase in biofilm components, in contrast to the 2.5-fold increase noted by the crystal violet procedure.

(ii) Analysis of biofilm components by CLSM.
Biofilms grown in 96-well plates in the presence or absence of sucrose were stained for each biofilm component for CLSM analysis. CLSM xy analyses provided biofilm surface coverage and indicated more uniform biofilm EPS with sucrose-supplemented medium (Fig. 4) than those in biofilms grown in the absence of sucrose (Fig. 4B). These results corroborate the increase in biofilm EPS by sucrose shown in Fig. 3. CLSM z-section analyses (Fig. 4C and D) demonstrated thicker biofilms ({approx}200 µm) in the presence of sucrose (panel C) than in the absence of sucrose ({approx}100 µm) (panel D), as indicated by the depth markers in Fig. 4A and B. CLSM z sections also provided surface coverage of the biofilms. As shown in Fig. 4B, sucrose deficiency resulted in patchy biofilms. In the presence of sucrose, CLSM assessments also indicated an enhancement in the thickness and uniformity of other biofilm components, i.e., lipids, proteins, and nucleic acids (data not shown). The spectral properties of the selected dyes facilitate CLSM assessments to colocalize biofilm lipids and nucleic acids or biofilm proteins and nucleic acids (data not shown). In all of these analyses, the selected dyes demonstrated uniform staining throughout the entire thickness of the biofilm and allowed for three-dimensional biofilm assessments.


Figure 4
View larger version (82K):
[in this window]
[in a new window]
  		FIG. 4. CLSM of biofilms grown in the presence or absence of 0.2% sucrose. Biofilms were stained with fluorescent probes to examine EPS. (A and B) CLSM images of biofilms grown in the presence and absence of sucrose, respectively (depth markings for CLSM images of biofilms are provided). Bar, 100 µm. (C and D) Corresponding biofilm topographies for panels A and B, respectively, derived by z-section analyses. The surface areas of the biofilms shown in panels C and D were obtained following analysis of 1,200- by 1,200-µm and 640- by 640-µm sections, respectively. The vertical aspects of the biofilms in panels C and D are 50 and 100 µm, respectively.

 
(iii) Multiparameter effects of antimicrobial agents on oral biofilm components.
The attributes unique to biofilms include relative resistance to inhibitory agents in comparison to that of planktonic cells (10, 12, 22, 25). Therefore, the influence of antimicrobial agents on the components of oral biofilms was studied. The effects of different concentrations of CHX and SLS on biofilm EPS and proteins (with these agents added to the medium during the 48-h period of biofilm formation) are shown in Fig. 5A and B, respectively. The results from triplicate samples indicated that increasing concentrations of either CHX or SLS resulted in corresponding increases in inhibition of biofilm EPS and proteins (P < 0.05). The least inhibition of biofilm EPS was noted with CHX at a concentration of 0.001% and was not significantly different from that of the control (P > 0.05). The inhibitory effects of these agents on biofilm proteins were considerably greater than those on biofilm EPS. As seen with biofilm EPS, dilution of CHX also resulted in a corresponding decrease in the inhibition of biofilm proteins. The inhibitory effects of different concentrations of SLS on biofilm components were more prominent than those noted with CHX. Similar procedures were used to examine the effects of different concentrations of ethanol (0.15%, 0.31%, 0.62%, 1.25%, 2.5%, 10%, and 20%) on biofilm components. Significant effects were observed with 20% ethanol compared to the untreated control (P < 0.05) (data not shown). Concentrations of ethanol below 20% did not demonstrate significant inhibition (P > 0.05) (data not shown).


Figure 5
View larger version (16K):
[in this window]
[in a new window]
  		FIG. 5. Effects of inhibitory agents on biofilm EPS and proteins. The effects of different concentrations of chlorhexidine (A) and sodium lauryl sulfate (B) are shown. Graphs indicate the residual biofilm EPS (lightly dotted bars) and proteins (heavily dotted bars) after treatments (averages for triplicate wells ± standard deviations) as percentages of those in the untreated control.

 
Another set of studies examined the effects of antibiotics on biofilm components. The MICs for oral strains are presented in Table 2. Whereas the MIC is used to define therapeutic strategies, concentrations of antibiotics below the MIC are prevalent during the therapeutic regimen. Therefore, studies have examined the effects of sub-MIC antibiotic doses on bacteria, including effects on bacterial surface properties, such as inhibition of bacterial adhesion and biofilm formation (1, 4, 8). Multiparameter effects of antibiotics with different modes of action (amoxicillin, doxycycline, erythromycin, and vancomycin) at one-fourth the MIC were elucidated. The results (Fig. 6) from quadruplicate samples indicated that the addition of one-fourth the MIC during the 48-h period of biofilm formation led to significant reductions of the residual biofilm components (presented as percentages of the amounts in the untreated controls) versus those in untreated biofilms (P < 0.05). Amoxicillin demonstrated little inhibition of biofilm EPS, proteins, and lipids, in contrast to the other antibiotics. Vancomycin demonstrated similar inhibition of three biofilm components. Doxycycline, on the other hand, demonstrated more inhibition of nucleic acids than of proteins, lipids, or EPS. Erythromycin demonstrated greater inhibition of proteins than did the other antibiotics. Similar effects were observed at one-half and one-eighth the MIC with these antibiotics (data not shown). Metronidazole (at all concentrations serially diluted from 2.5 mg/ml) was ineffective in MIC tests and did not result in biofilm inhibition (P > 0.05) (data not shown). The mucolytic agent N-acetyl-L-cysteine, which was tested at 6.25 mg/ml, demonstrated a significant inhibition of biofilm EPS (P < 0.05) (data not shown). With the exception of metronidazole, all agents inhibited biofilm components at the MIC (data not shown).


Figure 6
View larger version (32K):
[in this window]
[in a new window]
  		FIG. 6. Multiparameter assessments of the effects of antibiotics on oral biofilm components. Graphs indicate the residual biofilm components (averages for triplicate samples ± standard deviations) as percentages of those in the untreated control after treatment with 0.097 mg/ml amoxicillin, 0.0061 mg/ml erythromycin, 0.0061 mg/ml doxycycline, and 0.39 mg/ml vancomycin (see the legend in the figure).

 
A separate series of studies assessed the effects of a 30-min treatment with a range of agents on biofilm EPS. Neither treatments with amoxicillin, erythromycin, doxycycline, metronidazole, and vancomycin at their MICs nor treatments with different concentrations of ethanol (0.15% to 20%), chlorhexidine (0.001% to 0.15%), or sodium lauryl sulfate (0.125% to 1%) resulted in any significant reduction of biofilm components compared to untreated controls (data not shown).


   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The primary goal of this investigation with biofilms formed by several oral bacteria included the development of procedures for simultaneous assessment of the nonmicrobial constituents of biofilms for multiparameter analysis. An additional objective was to utilize these procedures to study the effects of common sugars, growth conditions, and antimicrobials on biofilm components. For these studies, analyses of biofilm components were accomplished by fluorescence-based quantitation and by CLSM. CLSM has found wide application for the investigation of hydrated biofilms (3, 12) and the examination of the overall structures of unstained oral biofilms (23) and the extracellular matrix (14, 18, 29).

Lectins derived from diverse sources demonstrate specificities for carbohydrate targets with affinities that resemble those of antigen-antibody interactions (16, 27). This unique property of lectins has been used previously to analyze microbial biofilms. For instance, previous reports demonstrated the utility of fluorescent lectins comprising ConA and/or WGA to examine biofilms of P. aeruginosa (27), Escherichia coli (18), and microbial isolates from food manufacturing facilities (16), as well as biofilms grown in river water (14). The binding specificities of ConA and WGA are well described in the literature and include affinities for mannopyranosyl and glucopyranosyl residues for ConA and sialic acid and N-acetylglucosamine for WGA. An optimal concentration of fluorescent ConA and WGA was selected on the basis of a surface response experimental design that tested several concentrations of each lectin. Lectins bound microbial biofilms at significantly higher levels than those for the salivary pellicle. Fluorescent dyes for the localization of glycoconjugates, proteins, lipids, and nucleic acids in oral biofilms were based on previous reports (14). Optimal concentrations of lectins and fluorescent dyes demonstrated significant staining of biofilms with low background fluorescence.

Environmental changes prevalent in the human mouth include alterations in osmolarity, pH, ionic strength, and specific nutrients (10, 12, 13, 15, 26) that serve as cues for the initiation and subsequent steps of biofilm formation (13, 25). The influences of different environmental and nutritional conditions on biofilm components were examined. Anaerobic conditions led to more biofilm EPS than did aerobic conditions, which was comparable to previous observations in Streptococcus gordonii and Streptococcus parasanguinis (9, 17). As a control, the effects of environmental and nutritional alterations on biofilm EPS were elucidated in parallel with quantitation of biofilm carbohydrate by the phenol-sulfuric acid procedure. Although similarities were observed between biofilm EPS estimations and biofilm carbohydrate analysis, biofilm EPS determinations provided significant discrimination under the conditions tested. The influences of ionic strength on biofilm components were not evident under the conditions tested.

Among dietary sugars, sucrose enhances biofilm EPS the most, with lesser effects noted for fructose, galactose, glucose, and lactose. Laboratory studies demonstrated the cariogenic properties of sucrose and its influences on the enhancement of insoluble glucans and biofilm formation among oral streptococci (13, 25). Correspondingly, in clinical studies, dental plaques revealed alterations in matrix proteins along with increases in insoluble glucans among human subjects provided sucrose (2). All biofilm components (EPS, proteins, lipids, and nucleic acids) were enhanced under sucrose-replete conditions. CLSM studies corroborated these observations, with thicker biofilms observed in the presence of sucrose, with larger amounts of EPS, nucleic acids, lipids, and proteins than those observed for sucrose deficiency. Quantitative fluorescence-based assessment of biofilm components demonstrated a >3-fold difference between biofilms grown in the presence and absence of sucrose. In contrast, an approximately 2.5-fold difference was observed by the widely used crystal violet analysis method (9, 17). Therefore, the reported fluorescence assessments provided larger differences than did the crystal violet procedure. Future studies will include investigations of the effects of simulated daily intake of food, dietary supplements, or specific nutritional factors on biofilm components. Another feature of these procedures is dual staining to examine EPS and nucleic acids for microplate fluorescence analysis and to colocalize lipids and nucleic acids or proteins and nucleic acids by CLSM due to the differences in fluorescence spectra of the selected dyes.

Pseudomonas aeruginosa, a nonoral organism, represents the paradigm bacterium for biofilm studies (12, 27, 28). Studies that assess biofilms of P. aeruginosa, including the commonly used crystal violet procedure, to estimate biofilm mass and biofilm carbohydrate are available. We included a few studies with P. aeruginosa to assess our procedures in relation to published observations. Our results with this bacterium corroborate with previous observations with P. aeruginosa.

Differences in the susceptibilities of planktonic and biofilm bacteria to antimicrobials remain an area of significant importance (10) in developing therapies for biofilm-mediated diseases. Data from bacterial viability analyses are commonly presented as the MICs or subinhibitory concentrations of antibiotics and antimicrobials (1, 4, 6, 8, 21). This investigation is the first to examine the simultaneous effects of antimicrobials on multiple biofilm components. Two types of studies were conducted. In the first series of studies, mature biofilms were treated for 30 min with the MICs of antibiotics. This resulted in negligible effects on biofilm components and corroborated earlier reports indicating modest effects of antibiotics on the bacteria of biofilms (4, 6, 10, 21, 25). In the second series of studies, significant reductions among biofilm components were observed when biofilms were allowed to form over a 48-h period in the presence of either the MICs or sub-MIC doses of agents. A relationship between the MIC and the inhibition of biofilm components was apparent for the antibiotics (amoxicillin, doxycycline, erythromycin, and vancomycin) and other agents (ethanol and N-acteyl-L-cysteine) tested. With SLS and CHX, a dose-dependent relationship between the concentration of agent and the inhibition of biofilm EPS and proteins was noted. Erythromycin demonstrated more inhibition of biofilm proteins than did doxycycline. At sub-MIC levels, amoxicillin, with its effects on cell wall synthesis, demonstrated lesser inhibitory effects on nucleic acids than did vancomycin and doxycycline. The effects of doxycycline were approximately the same for biofilm lipids and proteins as those for nucleic acids. Vancomycin, with its effect of increasing cell wall permeability and its inhibitory effects on cell wall and RNA synthesis, resulted in a similar inhibition of three biofilm components. Together, these results demonstrate the contrasting effects of the tested antibiotics on biofilm components for multiparameter assessments. These approaches will facilitate further characterization of the modes of action of agents (22) and the development of new therapeutic strategies for biofilm mitigation.

In conclusion, a broad platform utilizing fluorescence-based approaches to quantify selected biofilm components from mixed-species oral biofilms is presented. The procedures are amenable for routine use to examine the effects of several exogenous agents on biofilm components and future automation. Additional efforts indicate that the procedures are appropriate for ex vivo analysis of biofilms formed in the human mouth and on hydroxyapatite disks (data not shown). Together, the approaches form a broad platform useful for both laboratory efforts and clinical investigations to examine the kinetics of biofilm growth, effects of specific therapies for mode-of-action studies, influences of dietary factors, and differences in oral biofilms from subjects stratified on the basis of oral health.


   ACKNOWLEDGMENTS
 
We acknowledge R. Sharma and P. Moghe from The Confocal Microscopy and Cell Culture Facility for Biomaterials, Department of Chemical and Biochemical Engineering, Rutgers University, Piscataway, N.J., for confocal microscopy. Statistical analysis by V. Galicia and C. Kloos of Colgate Technology Statistics is gratefully acknowledged.


   FOOTNOTES
 
* Corresponding author. Mailing address: Colgate-Palmolive Company, 909 River Road, Piscataway, NJ 08855. Phone: (732) 878-6375. Fax: (732) 878-7084. E-mail: Prem_Sreenivasan{at}colpal.com. Back


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Cerca, N., S. Martins, S. Sillankorva, K. K. Jefferson, G. B. Pier, R. Oliveira, and J. Azeredo. 2005. Effects of growth in the presence of subinhibitory concentrations of dicloxacillin on Staphylococcus epidermidis and Staphylococcus haemolyticus biofilms. Appl. Environ. Microbiol. 71:8677-8682.[Abstract/Free Full Text]
  2. Cury, J. A., M. A. Rebelo, A. A. Del Bel Cury, M. T. Derbyshire, and C. P. Tabchoury. 2000. Biochemical composition and cariogenicity of dental plaque formed in the presence of sucrose or glucose and fructose. Caries Res. 34:491-497.[CrossRef][Medline]
  3. Davey, M. E., and G. A. O'Toole. 2000. Microbial biofilms: from ecology to molecular genetics. Microbiol. Mol. Biol. Rev. 64:847-867.[Abstract/Free Full Text]
  4. Di Bonaventura, G., I. Spedicato, D. D'Antonio, I. Robuffo, and R. Piccolomini. 2004. Biofilm formation by Stenotrophomonas maltophilia: modulation by quinolones, trimethoprim-sulfamethoxazole, and ceftazidime. Antimicrob. Agents Chemother. 48:151-160.[Abstract/Free Full Text]
  5. Dubois, M., K. A. Gilles, J. K. Hamilton, P. A. Rebers, and F. Smith. 1956. Colorimetric method for determination of sugars and related substances. Anal. Chem. 28:350-356.[CrossRef]
  6. Eick, S., T. Seltmann, and W. Pfister. 2004. Efficacy of antibiotics to strains of periodontopathogenic bacteria within a single species biofilm—an in vitro study. J. Clin. Periodontol. 31:376-383.[CrossRef][Medline]
  7. Fine, D. H., D. Furgang, H. C. Schreiner, P. Goncharoff, J. Charlesworth, G. Ghazwan, P. Fitzgerald-Bocarsly, and D. H. Figurski. 1999. Phenotypic variation in Actinobacillus actinomycetemcomitans during laboratory growth: implications for virulence. Microbiology 145:1335-1347.[Abstract]
  8. Fonseca, A. P., C. Extremina, A. F. Fonseca, and J. C. Sousa. 2004. Effect of subinhibitory concentration of piperacillin/tazobactam on Pseudomonas aeruginosa. J. Med. Microbiol. 53:903-910.[Abstract/Free Full Text]
  9. Froeliger, E. H., and P. Fives-Taylor. 2001. Streptococcus parasanguis fimbria-associated adhesin Fap1 is required for biofilm formation. Infect. Immun. 69:2512-2519.[Abstract/Free Full Text]
  10. Gilbert, P., T. Maira-Litran, A. J. McBain, A. H. Rickard, and F. W. Whyte. 2002. The physiology and collective recalcitrance of microbial biofilm communities. Adv. Microb. Physiol. 46:202-256.[Medline]
  11. Haffajee, A. D., S. S. Socransky, and J. C. Gunsolley. 2003. Systemic anti-infective periodontal therapy. A systematic review. Ann. Periodontol. 8:115-181.[CrossRef][Medline]
  12. Hall-Stoodley, L., J. W. Costerton, and P. Stoodley. 2004. Bacterial biofilms: from the natural environment to infectious diseases. Nat. Rev. Microbiol. 2:95-108.[CrossRef][Medline]
  13. Jenkinson, H. F., and R. J. Lamont. 2005. Oral microbial communities in sickness and in health. Trends Microbiol. 13:589-595.[CrossRef][Medline]
  14. Lawrence, J. R., G. D. Swerhone, G. G. Leppard, T. Araki, X. Zhang, M. M. West, and A. P. Hitchcock. 2003. Scanning transmission X-ray, laser scanning, and transmission electron microscopy mapping of the exopolymeric matrix of microbial biofilms. Appl. Environ. Microbiol. 69:5543-5554.[Abstract/Free Full Text]
  15. Lemos, J. A., T. A. Brown, Jr., and R. A. Burne. 2004. Effects of RelA on key virulence properties of planktonic and biofilm populations of Streptococcus mutans. Infect. Immun. 72:1431-1440.[Abstract/Free Full Text]
  16. Leriche, V., P. Sibille, and B. Carpentier. 2000. Use of an enzyme-linked lectinsorbent assay to monitor the shift in polysaccharide composition in bacterial biofilms. Appl. Environ. Microbiol. 66:1851-1856.[Abstract/Free Full Text]
  17. Loo, C. Y., K. Mitrakul, I. B. Voss, C. V. Hughes, and N. Ganeshkumar. 2003. Involvement of an inducible fructose phosphotransferase operon in Streptococcus gordonii biofilm formation. J. Bacteriol. 185:6241-6254.[Abstract/Free Full Text]
  18. Maeyama, R., Y. Mizunoe, J. M. Anderson, M. Tanaka, and T. Matsuda. 2004. Confocal imaging of biofilm formation process using fluoroprobed Escherichia coli and fluoro-stained exopolysaccharide. J. Biomed. Mater. Res. 70A:274-282.[CrossRef]
  19. National Committee for Clinical Laboratory Standards. 2002. Performance standards for antimicrobial susceptibility testing, 12th informational supplement. M100-S12. National Committee for Clinical Laboratory Standards, Wayne, Pa.
  20. Nikawa, H., T. Hamada, H. Yamashiro, H. Murata, and A. Subiwahjudi. 1998. The effect of saliva or serum on Streptococcus mutans and Candida albicans colonization of hydroxylapatite beads. J. Dent. 26:31-37.[CrossRef][Medline]
  21. Noiri, Y., Y. Okami, M. Narimatsu, Y. Takahashi, T. Kawahara, and S. Ebisu. 2003. Effects of chlorhexidine, minocycline, and metronidazole on Porphyromonas gingivalis strain 381 in biofilms. J. Periodontol. 74:1647-1651.[CrossRef][Medline]
  22. O'Neill, A. J., and I. Chopra. 2004. Preclinical evaluation of novel antibacterial agents by microbiological and molecular techniques. Expert Opin. Investig. Drugs 13:1045-1063.[CrossRef][Medline]
  23. Pratten, J., C. S. Andrews, D. Q. Craig, and M. Wilson. 2000. Structural studies of microcosm dental plaques grown under different nutritional conditions. FEMS Microbiol. Lett. 189:215-218.[CrossRef][Medline]
  24. Rozen, R., G. Bachrach, M. Bronshteyn, I. Gedalia, and D. Steinberg. 2001. The role of fructans on dental biofilm formation by Streptococcus sobrinus, Streptococcus mutans, Streptococcus gordonii and Actinomyces viscosus. FEMS Microbiol. Lett. 195:205-210.[CrossRef][Medline]
  25. Socransky, S. S., and A. D. Haffajee. 2002. Dental biofilms: difficult therapeutic targets. Periodontol. 2000 28:12-55.
  26. Steinberg, D., M. Feldman, I. Ofek, and E. I. Weiss. 2004. Effect of a high-molecular-weight component of cranberry on constituents of dental biofilm. J. Antimicrob. Chemother. 54:86-89.[Abstract/Free Full Text]
  27. Strathmann, M., J. Wingender, and H. C. Flemming. 2002. Application of fluorescently labeled lectins for the visualization and biochemical characterization of polysaccharides in biofilms of Pseudomonas aeruginosa. J. Microbiol. Methods 50:237-248.[CrossRef][Medline]
  28. Sutherland, I. W. 2001. The biofilm matrix—an immobilized but dynamic microbial environment. Trends Microbiol. 9:222-227.[CrossRef][Medline]
  29. Thurnheer, T., R. Gmur, S. Shapiro, and B. Guggenheim. 2003. Mass transport of macromolecules within an in vitro model of supragingival plaque. Appl. Environ. Microbiol. 69:1702-1709.[Abstract/Free Full Text]
  30. Vesey, P. M., and H. K. Kuramitsu. 2004. Genetic analysis of Treponema denticola ATCC 35405 biofilm formation. Microbiology 150:2401-2407.[Abstract/Free Full Text]

Applied and Environmental Microbiology, October 2006, p. 6734-6742, Vol. 72, No. 10
0099-2240/06/$08.00+0     doi:10.1128/AEM.01013-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.



This article has been cited by other articles:


This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Yang, Y.
Right arrow Articles by Cummins, D.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Yang, Y.
Right arrow Articles by Cummins, D.
Agricola
Right arrow Articles by Yang, Y.
Right arrow Articles by Cummins, D.

Home Help [Feedback] [For Subscribers] [Archive] [Search] [Contents]
J. Bacteriol. Microbiol. Mol. Biol. Rev. Eukaryot. Cell All ASM Journals