Antimicrobial Effect of Chitosan Nanoparticles on Streptococcus ...

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Dec 16, 2010 - Duncan S. Sutherland,2 and Peter L. Wejse3. Department of Oral .... Chávez de Paz, L. E., I. R. Hamilton, and G. Svensäter. 2008. Oral bacteria.

APPLIED AND ENVIRONMENTAL MICROBIOLOGY, June 2011, p. 3892–3895 0099-2240/11/$12.00 doi:10.1128/AEM.02941-10 Copyright © 2011, American Society for Microbiology. All Rights Reserved.

Vol. 77, No. 11

Antimicrobial Effect of Chitosan Nanoparticles on Streptococcus mutans Biofilms䌤 Luis E. Cha´vez de Paz,1* Anton Resin,2 Kenneth A. Howard,2 Duncan S. Sutherland,2 and Peter L. Wejse3 Department of Oral Biology, Faculty of Odontology, Malmo ¨ University, S-20506 Malmo ¨, Sweden1; Interdisciplinary Nanoscience Center (iNANO), Faculty of Science, Aarhus University, DK-8000 Aarhus C, Denmark2; and Global Ingredients R&D, Arla Foods amba, Viby J, Denmark3 Received 16 December 2010/Accepted 1 April 2011

Nanoparticle complexes were prepared from chitosans of various molecular weights (MW) and degrees of deacetylation (DD). The antimicrobial effect was assessed by the Live/Dead BacLight technique in conjunction with confocal scanning laser microscopy (CSLM) and image analysis. Nanocomplexes prepared from chitosans with high MW showed a low antimicrobial effect (20 to 25% of cells damaged), whereas those prepared from low-MW chitosans showed high antimicrobial effect (>95% of cells damaged). These chitosans were categorized into three groups: group A (high DD and low MW), group B (high DD and high MW), and group C (low DD and low MW). Chitosan nanoparticles were created by ion gelation with polyanionic sodium triphosphate (TPP) (Fig. 2a) (13). Since one of the reported problems with chitosan preparations has been its low solubility under neutral pH conditions, we further developed the system to produce stable nanoparticle at neutral pH. Nanoparticle assemblies were generally formulated by dissolving chitosan in acetic acid buffer (1 mg/ml) and by adding 100 ␮l of this mixture drop by drop while stirring vigorously to a premixed solution composed of 50 ␮l of TPP solution (0.1%) in water and 500 ␮l of phosphate-buffered saline (PBS) buffer (pH 7.4). The mass ratio of chitosan to TPP was kept constant (2:1). The final pH of the solution was around 5 and was adjusted to 7 by the addition of 2 M NaOH. The nanoparticle solution was equilibrated at room temperature for an hour by mixing. Particle size distribution and zeta potential were further characterized in a Malvern Zetasizer Nano ZS instrument to measure particle size distributions and zeta potential in different media. Scanning electron microscopy (SEM) (Fig. 2b) was also used to visualize particles in deionized water before mixing with media. The particle size distribution in water was relatively narrow with some batch-to-batch variation. After mixing the particles with media, the size distribution broadened and was between 20 and 1,000 nm showing a similar profile for the different formulations. Exposure of S. mutans biofilms to chitosan nanoparticles. Biofilms were formed in the flow chamber system ␮-Slide VI (Integrated BioDiagnostics) as previously described (2). In brief, 120 ␮l of a washed suspension of Streptococcus mutans UA159 in mid-exponential growth (optical density at 600 nm [OD600] ⫽ 0.4 ⫾ 0.1) were inoculated in the mini-flow chamber slides and incubated in an atmosphere of 5% CO2 at 37°C for 24 h under static conditions. Flow chambers were rinsed with PBS to remove nonadherent cells. S. mutans biofilms were then exposed to the different chitosan/TPP nanoparticle formulations diluted in Todd-Hewitt medium (BD Biosciences, Sweden) at a ratio of 1:1. Ratios of 1:4 and 1:10 were also tested.

Oral biofilm communities are naturally formed on tooth surfaces and are associated with diseases such as caries, gum inflammation (gingivitis), and degradation of periodontal tissues (periodontitis) (5, 6). These diseases are mainly caused by those biofilm organisms that exhibit phenotypic traits capable of surviving adverse environmental conditions. For example, Streptococcus mutans undergoes an acid tolerance response to adapt and survive the acidic conditions provoked by excess sugar intake (3, 11). This important phenotypic trait of S. mutans is directly linked with caries development (5, 12). Therefore, novel approaches for developing oral care products, such as dentifrices and mouthwashes, rely on targeting these highly adaptable oral organisms and blocking their key mechanisms of phenotypic variation. One major step forward in achieving this goal has been the development of antimicrobial systems that could effortlessly diffuse across all biofilm structures (4). For this purpose, there has been increasing interest in developing nanoscale systems to be used as biological carriers within biofilms. Of special interest are those nanoscale systems developed from natural polymers, e.g., chitosan (10). Chitosan is obtained by deacetylation of chitin and is used in biomedical applications due to its high biocompatibility and antimicrobial properties (8). In the present study, we formulated nanoparticles from different commercially available chitosans (with different molecular weights [MW] and degrees of deacetylation [DD]) by ion gelation with polyanionic sodium triphosphate (TPP) and studied their penetrative antimicrobial effect on 24-h-old biofilms of S. mutans. Formulation of chitosan nanoparticles. The first step was to formulate nanoparticles using various chitosans with different degrees of deacetylation and molecular weights (Bioneer A/S and NovaMatrix, Norway). Figure 1 shows 9 different chitosans used in this study distributed according to their DD and MW.

* Corresponding author. Mailing address: Department of Oral Biology, Faculty of Odontology, Malmo ¨ University, SE-20506 Malmo ¨, Sweden. Phone: 46 40 6658659. Fax: 46 40 929359. E-mail: luis.chavez [email protected] 䌤 Published ahead of print on 15 April 2011. 3892

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FIG. 1. Categorization of the different chitosan subtypes based on their molecular size (kDa) and degree of deacetylation.

A solution of TPP (0.1%) in PBS was used as control. Flow cells were incubated for 2 h in an atmosphere of 5% CO2 at 37°C. Antimicrobial activity was assessed using the Live/Dead BacLight bacterial viability kit for microscopy (Invitrogen Ltd., Paisley, United Kingdom). The fluorescence from stained cells was viewed using an inverted confocal scanning laser microscope, Eclipse TE2000 (Nikon, Tokyo, Japan), where 10 randomly selected image stack sections were imaged in each biofilm sample. Image stack sections were composed of 10 images, each taken with a variation of 2 ␮m along the z position. Images were acquired with a 60⫻ oil immersion objective and digitalized by the software EZ-C1 v.3.40 build 691 (Nikon, Tokyo, Japan) at a resolution of 512 by 512 pixels and with a zoom factor of 1.0, giving a final pixel resolution of 0.42 ␮m/ pixel. Individual biofilm images covered an area of 0.05 mm2 per field of view. Experiments were done in three triplicate sets of biofilms with each set having its own unique total cell count. Confocal scanning laser microscopy (CSLM) images were analyzed to produce information of the total biofilm population as well as the independent subpopulations represented by red and green fluorescent colors by using the bioImage_L software program (1). The structure and spatial differences in green (viable) and red (damaged) biofilm subpopulations were characterized by three-dimensional analysis where parameters such as biovolume (␮m3), substratum coverage (reported as a percentage), and spatial thickness (␮m) were calculated. Large effect of low-molecular-weight chitosans. When the biofilms were 24 h old, they were observed to have average substratum coverage of 44.7% ⫾ 3.1%, mean total biovolume of 6.2 ⫻ 104 ⫾ 0.4 ⫻ 104 ␮m3, mean thickness of 16.4 ⫾ 1.1 ␮m, and 96.4% ⫾ 2.3% of its total population with intact cell membranes (cells that were stained green when the Live/Dead BacLight bacterial viability kit was used). After the application of the chitosan nanoparticles, no significant difference in the structure of biofilms was found. However, the effect of the tested nanoparticle formulations gave strong differences in the level of cell membrane damage (cells that stained red when the Live/Dead BacLight bacterial viability kit was used) compared with the negative control (TPP) (see representative three-dimensional [3D] biofilm reconstructions in Fig. 3). The

FIG. 2. Formation of the chitosan-tripolyphosphate complex by ionotropic gelation. (a) Schematic illustration of the chitosan-TPP complex and (b) SEM image. Bar, 200 nm.

ratio that the chitosan nanoparticles were mixed in the medium did not have an effect on their antimicrobial activity (data not shown). The representative nanoparticle preparations comprising chitosans with low molecular weights, groups A and C, showed the highest antimicrobial activity at the various depths of the biofilms formed by S. mutans (⬎95% of total cells damaged). These results indicated that the differences in the levels of deacetylation in group A (high DD) and group C (low DD) did not seem to affect the antimicrobial activity of the chitosan nanoparticles. This lack of influence of the deacetylation level may be due to free amino groups being neutral at pH 7. This may explain why our results differ from results in other studies that found a positive correlation between the levels of deacetylation and the antimicrobial effect of chitosan at a lower pH (7). In contrast, we observed a clear tendency between the molecular weight of the chitosan and the effect on the membrane integrity of S. mutans with the lower-molecular-weight chitosans showing the highest effect (groups A and C) and with progressively decreased effect on membrane integrity for higher molecular weights (group B) (Table 1). Chitosan nanoparticles from group B showed a scattered effect in damaging the cell membrane of cells (20 to 25% of cells damaged) mainly at the upper levels of the biofilms (⬎15 ␮m). At the levels closer to the substrate (⬍4 ␮m), chitosan nanoparticles showed a slight effect in cell membrane damage (⬍5%). We do not have a clear picture of the mechanism of the molecular weight effect on chitosan antimicrobial activity in nanoparticles. The heterogeneous distribution of cell membrane damage at the upper levels of the biofilms and the lack of effect at the substratum levels when using high-MW formulations, how-






FIG. 3. Antimicrobial effect of different chitosan-tripolyphosphate complexes in S. mutans biofilms. 3D biofilm reconstructions show results with the Live/Dead stain (green [viable cells] and red [damaged cells]) at different depths of the biofilms. The units on the axes are micrometers.

ever, indicate a lower diffusive potential than low-MW formulations which affected the membrane integrity of cells all across the biofilm. It is also possible that the lower-molecular-weight nanoparticles have a systematically reduced number of TPP molecules available per molecule and may thus be more susceptible to disaggregation in the biofilm. Once homogenously distributed across the biofilms, chitosan particles can directly interact with bacterial cells. Different mechanisms of interaction with bacteria have been proposed for chitosans (9). One proposed mechanism is based on the interaction between positively charged chitosan molecules and negatively charged microbial cell membranes leading to the leakage of proteinaceous and other intracellular constituents; however, at pH 7, free chitosan and the outer chitosan in the nanoparticle complexes are expected to be neutral, but chitosan within the particle may retain its charge. A second proposed mechanism is based on

TABLE 1. Antimicrobial effect of different groups of chitosan nanoparticlesa

binding of chitosan with microbial DNA, in turn interfering with mRNA and protein synthesis, which clearly requires entry of the chitosan into the cell. Concluding remarks. This study showed antimicrobial activity of chitosan nanoparticles at neutral pH to have a strong trend toward higher activity of particles formed from lower-MW chitosans. Furthermore, the effect of low-molecular-weight formulations affected the cell membrane integrity of S. mutans in a homogenous manner across the entire biofilm. It is expected that this system will greatly improve the uniform delivery of chitosan formulated as nanoparticles through biofilm structures at neutral pH, and combined with other compounds, it could aid targeting of strongly adaptable organisms in complex biofilm systems. This work was carried out within the ProSURF platform (ProteinBased Functionalisation of Surfaces), which is funded by the Danish National Advanced Technology Foundation. We are grateful to Jørgen Kjems (Interdisciplinary Nanoscience Center, Aarhus University) for assistance in formulating the chitosan nanoparticles.

% of damaged biofilm cells (mean ⫾ SEM) Level (distance from substratum)

Upper (20 ␮m) Middle (15 ␮m) Low (2 ␮m) a b

Group A (low MW, high DD)b

Group B (high MW, high DD)

Group C (low MW, low DD)

95.5 ⫾ 0.8 94.6 ⫾ 1.1 96.1 ⫾ 3

21.4 ⫾ 1.2 7.5 ⫾ 1 1.2 ⫾ 0.6

94.9 ⫾ 1 93.6 ⫾ 1.7 96.7 ⫾ 2.3

The values for the control (no chitosan nanoparticles) were ⬍1 for all levels. MW, molecular weight; DD, degree of deacetylation.

REFERENCES 1. Cha ´vez de Paz, L. E. 2009. Image analysis software based on color segmentation for characterization of viability and physiological activity of biofilms. Appl. Environ. Microbiol. 75:1734–1739. 2. Cha ´vez de Paz, L. E., I. R. Hamilton, and G. Svensa ¨ter. 2008. Oral bacteria in biofilms exhibit slow reactivation from nutrient deprivation. Microbiology 154:1927–1938. 3. Hamilton, I. R., and N. D. Buckley. 1991. Adaptation by Streptococcus mutans to acid tolerance. Oral Microbiol. Immunol. 6:65–71.

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4. Hetrick, E. M., J. H. Shin, H. S. Paul, and M. H. Schoenfisch. 2009. Antibiofilm efficacy of nitric oxide-releasing silica nanoparticles. Biomaterials 30:2782–2789. 5. Marsh, P. D. 2004. Dental plaque as a microbial biofilm. Caries Res. 38: 204–211. 6. Marsh, P. D. 2005. Dental plaque: biological significance of a biofilm and community life-style. J. Clin. Periodontol. 32(Suppl. 6):7–15. 7. Nasti, A., et al. 2009. Chitosan/TPP and chitosan/TPP-hyaluronic acid nanoparticles: systematic optimisation of the preparative process and preliminary biological evaluation. Pharm. Res. 26:1918–1930. 8. Rabea, E. I., M. E. Badawy, C. V. Stevens, G. Smagghe, and W. Steurbaut. 2003. Chitosan as antimicrobial agent: applications and mode of action. Biomacromolecules 4:1457–1465.


9. Senel, S., et al. 2000. Chitosan films and hydrogels of chlorhexidine gluconate for oral mucosal delivery. Int. J. Pharm. 193:197–203. 10. Sinha, V. R., et al. 2004. Chitosan microspheres as a potential carrier for drugs. Int. J. Pharm. 274:1–33. 11. Svensa ¨ter, G., U. B. Larsson, E. C. Greif, D. G. Cvitkovitch, and I. R. Hamilton. 1997. Acid tolerance response and survival by oral bacteria. Oral Microbiol. Immunol. 12:266–273. 12. Van Houte, J., C. Sansone, K. Joshipura, and R. Kent. 1991. Mutans streptococci and non-mutans streptococci acidogenic at low pH, and in vitro acidogenic potential of dental plaque in two different areas of the human dentition. J. Dent. Res. 70:1503–1507. 13. Zhang, H., M. Oh, C. Allen, and E. Kumacheva. 2004. Monodisperse chitosan nanoparticles for mucosal drug delivery. Biomacromolecules 5:2461–2468.

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