The Scientific World Journal
Volume 2014 (2014), Article ID 706189, 5 pages
http://dx.doi.org/10.1155/2014/706189

Research Article

Continuous Drip Flow System to Develop Biofilm of E. faecalis under Anaerobic Conditions

Ana Maria Gonzalez,¹ Erika Corpus,¹ Amaury Pozos-Guillen,² Daniel Silva-Herzog,¹ Antonio Aragon-Piña,³ and Nestor Cohenca⁴

¹Endodontics Postgraduate Program, Faculty of Dentistry, The Autonomous University of San Luis Potosí, Manuel Nava 2, 78290 San Luis Potosí, SLP, Mexico
²Basic Sciences Laboratory, Faculty of Dentistry, The Autonomous University of San Luis Potosí, Manuel Nava 2, 78290 San Luis Potosí, SLP, Mexico
³Faculty of Engineering, The Autonomous University of San Luis Potosí, Manuel Nava 8, 78290 San Luis Potosí, SLP, Mexico
⁴Department of Endodontics and Department of Pediatric Dentistry, School of Dentistry, University of Washington, 6222 NE 74th Street, Seattle, 98115 WA, USA

Received 14 July 2014; Accepted 28 August 2014; Published 13 October 2014

Academic Editor: Louis M. Lin

Copyright © 2014 Ana Maria Gonzalez et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract.

Purpose. To evaluate a structurally mature E. faecalis biofilm developed under anaerobic/dynamic conditions in an in vitro system. Methods. An experimental device was developed using a continuous drip flow system designed to develop biofilm under anaerobic conditions. The inoculum was replaced every 24 hours with a fresh growth medium for up to 10 days to feed the system. Gram staining was done every 24 hours to control the microorganism purity. Biofilms developed under the system were evaluated under the scanning electron microscope (SEM). Results. SEM micrographs demonstrated mushroom-shaped structures, corresponding to a mature E. faecalis biofilm. In the mature biofilm bacterial cells are totally encased in a polymeric extracellular matrix. Conclusions. The proposed in vitro system model provides an additional useful tool to study the biofilm concept in endodontic microbiology, allowing for a better understanding of persistent root canal infections.

1. Introduction

Contemporary microbiology has demonstrated that microorganisms are organized under specific environmental conditions [1]. Planktonic microorganisms are single cells that may float or swim in a liquid medium. In contrast, under specific environmental conditions, phenotypic adaptations are expressed as a form of life termed biofilm. Biofilm is characterized by the immobilization on a surface, cell-cell interactions, formation of microcolonies, excretion of extracellular polymers (EPS), and development of three-dimensional structures which confer protection in order to ensure their permanence [1, 2].

Mature biofilm is a complex heterogeneous structure of dormant and actively growing bacteria colonies along with enzymes, excretory products, and small channels forming part of the overall structure [3]. It is also known that a mature biofilm is able to tolerate antimicrobial concentrations of 10 to 1000 times that required to remove planktonic bacteria [4]. The biofilm concept has changed and it is currently defined as “a bacterial community immersed in a liquid medium, characterized by one or more bacteria that are attached to each other, to a substrate or surface and embedded in an extracellular matrix produced by them, and shows an altered phenotype in the degree of cell proliferation or the expression of their genes” [5]. Thus, the study of biofilms has skyrocketed in recent years due to the increased awareness of the pervasiveness and impact of biofilms on natural and industrial systems, as well as human health (chronic bacterial prostatitis, cyst fibrosis, and periodontitis) [5].

Oral biofilms are unique mainly due to the fact that they may form under highly different factors. In the oral cavity, biofilms are found as dental plaque over the enamel surface, over the root surface in periodontal disease, and in the internal and external surface of roots suffering pulpal and periapical pathology. In the category of secondary endodontic infections, E. faecalis is the microorganism most often associated with asymptomatic chronic periradicular lesions, while not being as prominent in acute periapical periodontitis or acute periradicular abscess [6–9].

It is reported to be present in 4 to 40% of secondary endodontic infections. The frequency of E. faecalis in persistent lesions has been shown to be much higher. Previous work related to refractory periapical lesions has shown a prevalence ranging from 24 to 77%. In some cases it has been found as the only inhabitant in teeth endodontically treated [6, 10, 11]. Current literature has demonstrated that biofilms may remain viable in anatomical areas of the root canal system that remain untouched by either mechanical or chemical disinfection [12–14]. Clinical examination of root tips associated with refractory periapical periodontitis has suggested the presence of bacterial biofilm at the apical portion of the root canal [15–18]. In vitro studies have focused on the efficacy of selected irrigants and medicaments to arrest biofilms grown in different substrates [19–23], using strains of selected species and nonputative strains from the root canal. However, there is a lack of information relating to biofilm formation capabilities and characteristics of clinical isolates recovered from the root canals. Moreover, the development and validation of practical, reproducible, and clinically relevant laboratory models for the study of biofilms are still challenging. The aim of the present study was to evaluate a structurally mature E. faecalis biofilm developed under anaerobic/dynamic conditions in an in vitro system.

2. Material and Methods

An experimental device was created using a continuous drip flow system designed to develop biofilm under anaerobic conditions (Coy Laboratory Products, Grass Lake, MI, USA) (Figure 1). This experimental device was previously sterilized and built as follows. A plastic container (Figure 1(a)) with fresh growth medium was connected by tubing to a plastic chamber containing the specimen (Figure 1(b)) on which the biofilms were formed. Twenty specimens were performed as follows. The apical 3 mm of teeth recently instrumented and extracted was removed from each root. Samples were taken to an ultrasonic bath (BioSonic UC50, Coltene/Whaledent Inc., NJ, USA) soaking in EDTA (Fermont, Monterrey Chemicals, SA, Mexico) to 17% for 4 min and then samples were immersed in 5.25% sodium hypochlorite for 4 min to remove organic and inorganic tissue. The specimens were then sterilized at 121°C and 15 pounds of pressure for 20 min. A second container (Figure 1(d)) for waste collection was used. The flow was controlled to a velocity of 0.5 mL/min (Figure 1(c)).

Figure 1: Experimental device developed under a continuous drip flow system and designed for dynamic biofilm formation under anaerobic conditions. Plastic container with fresh growth medium (a) connected to a plastic chamber containing the specimen (b). The flow was controlled to a velocity of 0.5 mL/min (c) and the waste collected on a second container (d). Stepwise scale-up strategy to reach a final concentration of 1.5 × 10⁸ cells/mL (e).

The inoculum of E. faecalis previously developed and identified in root tips associated with endodontic failure was replaced every 24 hours with a fresh growth medium for up to 10 days to feed the system. The fresh medium was brain-heart infusion broth (BBL Becton Dickinson Mexico, Cuautitlan, Mexico). The inoculums were prepared using a stepwise scale-up strategy in a proportion of 1 : 10 (Figure 1(e)) to reach a final concentration of 1.5 × 10⁸ cells/mL, adjusted to 0.5 Mac Farland turbidity standard. This step is repeated from 100 to 1000 mL to obtain the volume required by the experimental device in order to be replaced in the continuous flow system every 24 hours for the time necessary for the experimental phase. Gram staining was done every 24 hours to control the microorganism purity.

All the samples were prepared for SEM analysis as follows: they were gently washed with 0.1 M phosphate buffered solution, fixed with 2% glutaraldehyde (Sigma-Aldrich, St. Louis, MO, USA) and 1% Alcian Blue Stain 8GX (Sigma-Aldrich, St. Louis, MO, USA), and stored at 4°C for 24 hours. Once fixed, samples were washed three times with 0.1 M phosphate buffer solution to remove excess material. The samples were dehydrated in a series of anhydrous ethanol (Industrial Chemical Technology, Ltd.) 20%, 40%, 60%, 80%, 90%, and 95% for 10 minutes in each series to be left finally immersed in ethanol 100% and critical point dried in a critical point dryer (CPD 030 BAL-TEC GmbH, Schalksmühle, Germany). The samples were sputter coated with 20 nm gold-palladium mixture (Fine Coat Ion Sputter JFC-1100, USA). Biofilms specimens were evaluated under the SEM (JEOL JSM-6610 LV, JAPAN) at a 5 Kv accelerating potential at different magnifications.

3. Results

SEM analysis revealed bacterial biofilm in all samples. Careful observation of these structures under higher magnification revealed clumps of coaggregated bacterial cells in a matrix of extracellular polymeric substance. E. faecalis biofilms displayed a complex three-dimensional structure which demonstrated spatial heterogeneity and a typical architecture showing microcolonies with ramifying water channels. Fibrillar structures appeared to be made up of twisted fibers. Larger structures of wrapped sheets were also present and consisted of small numbers of bacteria cells embedded in a matrix of fibers. The E. faecalis biofilm, grown under anaerobic conditions, showed a clump of bacteria cells attached to the dentine surface (Figure 2(a)). The fibers were more apparent and formed irregular, net-like structures (Figure 2(b)). Coaggregated bacterial cells in a polymeric matrix were observed forming “mushroom-shaped structures,” corresponding to a mature biofilm with cells totally encased in a polymeric extracellular matrix (Figures 2(c) and 2(d)).

(a)

(b)

(c)

(d)

Figure 2: In vitro E. faecalis biofilm formed in an anaerobic/dynamic system. ((a) and (c)) Coccoid structures attached to mature biofilm (6000x). (b) Extrapolymeric fibers (10000x). (d) “Mushroom-shaped structures” (10000x).

4. Discussion

The system discussed in this study is based on a modified Robbins device (MRD) developed by Jim Robbins to allow the reproducible and simultaneous formation of biofilms exposed to a fluid flow [24–26]. MRD is filled with a suspension of microorganisms and is inverted to improve the adhesion of the planktonic cells to the discs. Based on that model, an experimental device was developed using a disposable infusion pump in which it was possible to maintain a constant flow of culture medium infected with the microorganism on dentin surfaces for 10 days, time required to carry out the process of redistribution of cells adhered by mobility on the surface, cell division and aggregation, and finally the excretion of polymers with the consequent formation of a mature biofilm [27–29].

Based on our results and in concordance with Dunavant et al. [19] continuous flow of nutrients is the key to developing a mature and robust biofilm. As demonstrated in the present study, we successfully created an in vitro model system capable of developing E. faecalis biofilms under anaerobic conditions using a continuous flow of nutrients. The flow distribution is an important factor, allowing bacteria to make secondary colonization and contribute to developing a more robust biofilm in areas with minor nutrients. Thein et al. [30] have found that, in comparison with static conditions, dynamic conditions have a significant positive impact on microbial biofilm growth. This novel research model simulates clinical conditions in which disintegration of the sealer or undetected voids in the filling mass may create leakage channels that allowed periradicular tissue fluids to reach residual bacteria within tubules and provide nutrient for their growth [31].

The results of the present study also demonstrate that E. faecalis is capable of forming biofilms under anaerobic conditions. To our knowledge, this is one of the few studies which evaluate E. faecalis biofilm formation under anaerobic conditions. The study is based on the fact that root canal conditions and moreover the periapical area are under strict anaerobic environment. The biofilm mode of growth is a survival strategy and harsh environmental conditions existing in the root canal may favor the growth of bacteria as a biofilm. George et al. [32] found that when E. faecalis was grown under anaerobic and nutrient rich conditions, a matured biofilm with apparent water channels on the root canal was observed. On the other hand, when cultured aerobically the typical biofilm structure with mushroom-shaped microcolonies was not observed.

The current microstructural findings show that a mature biofilm formation was achieved by the end of 10 days. SEM images show an irregular layer on the dentin which consists of ovoid (similar to bacilli), elongated, and filamentous forms. Also, some coccoid forms wrapped in a kind of sheath which joined others forming an elongated structure, similar to those described for the first time by Ramachandran Nair [33]. Water channels as well as mushroom-shaped microcolonies were observed. This study suggests that hydrodynamics and environmental conditions are a significant factor in the quality of E. faecalis biofilm development. It is important to mention that the literature has described numerous systems of biofilm formation, both static and dynamic, and their applications in endodontics using E. faecalis as a microorganism of study due to its resistance to endodontic treatment [34]. However, a system in anaerobic conditions is difficult to realize. The model proposal in this study has the advantage of being easy to assemble in an anaerobic chamber; it can be to perform multiple systems at once; there is less risk of contamination because the system is not reusable; it is possible to change the flow if required; it is possible to control the time from hours to days to observe and study the formation of the biofilm thickness. It is also possible to obtain sufficient quantities of biofilm in order to realize different assays. Further studies with nondestructive chemical analytical techniques are necessary to increase our knowledge of the biofilm chemical composition in this model system.

5. Conclusion

The proposed in vitro system model provides an additional useful tool to study the biofilm concept in endodontic microbiology, allowing for a better understanding of persistent root canal infections.

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.

Acknowledgment

This study was registered and approved by the Institutional Ethics Committee of the Faculty of Dentistry at San Luis Potosi University, San Luis Potosi, Mexico (Approval Code no. CEIFE-012-010).

References

J. W. Costerton, Z. Lewandowski, D. DeBeer, D. Caldwell, D. Korber, and G. James, “Biofilms, the customized microniche,” Journal of Bacteriology, vol. 176, no. 8, pp. 2137–2142, 1994.

G. O'Toole, H. B. Kaplan, and R. Kolter, “Biofilm formation as microbial development,” Annual Review of Microbiology, vol. 54, pp. 49–79, 2000.

W. M. Dunne Jr., “Bacterial adhesion: seen any good biofilms lately?” Clinical Microbiology Reviews, vol. 15, no. 2, pp. 155–166, 2002.

S. S. Socransky, A. D. Haffajee, C. Smith, and G. W. Duff, “Microbiological parameters associated with IL-1 gene polymorphisms in periodontitis patients,” Journal of Clinical Periodontology, vol. 27, no. 11, pp. 810–818, 2000.

R. M. Donlan and J. W. Costerton, “Biofilms: survival mechanisms of clinically relevant microorganisms,” Clinical Microbiology Reviews, vol. 15, no. 2, pp. 167–193, 2002.

I. N. Rôças, J. F. Siqueira Jr., and K. R. N. Santos, “Association of Enterococcus faecalis with different forms of periradicular diseases,” Journal of Endodontics, vol. 30, no. 5, pp. 315–320, 2004.

P. N. Nair, “On the causes of persistent apical periodontitis: a review,” International Endodontic Journal, vol. 39, no. 4, pp. 249–281, 2006.

C. H. Stuart, S. A. Schwartz, T. J. Beeson, and C. B. Owatz, “Enterococcus faecalis: its role in root canal treatment failure and current concepts in retreatment,” Journal of Endodontics, vol. 32, no. 2, pp. 93–98, 2006.

G. di Filippo, S. K. Sidhu, and B. S. Chong, “The role of biofilms in endodontic treatment failure,” ENDO-Endodontic Practice Today, vol. 8, no. 2, pp. 87–103, 2014.

E. T. Pinheiro, B. P. F. A. Gomes, C. C. R. Ferraz, E. L. R. Sousa, F. B. Teixeira, and F. J. Souza-Filho, “Microorganisms from canals of root-filled teeth with periapical lesions,” International Endodontic Journal, vol. 36, no. 1, pp. 1–11, 2003.

G. Sundqvist and D. Figdor, “Life as an endodontic pathogen. Ecological differences between the untreated and root-filled root canals,” Endodontic Topics, vol. 6, no. 1, pp. 3–28, 2003.

P. N. R. Nair, S. Henry, V. Cano, and J. Vera, “Microbial status of apical root canal system of human mandibular first molars with primary apical periodontitis after “one-visit” endodontic treatment,” Oral Surgery, Oral Medicine, Oral Pathology, Oral Radiology and Endodontology, vol. 99, no. 2, pp. 231–252, 2005.

D. Ricucci and J. F. Siqueira Jr., “Biofilms and apical periodontitis: study of prevalence and association with clinical and histopathologic findings,” Journal of Endodontics, vol. 36, no. 8, pp. 1277–1288, 2010.

D. Ricucci and J. F. Siqueira Jr., “Fate of the tissue in lateral canals and apical ramifications in response to pathologic conditions and treatment procedures,” Journal of Endodontics, vol. 36, no. 1, pp. 1–15, 2010.

Y. Noiri, A. Ehara, T. Kawahara, N. Takemura, and S. Ebisu, “Participation of bacterial biofilms in refractory and chronic periapical periodontitis,” Journal of Endodontics, vol. 28, no. 10, pp. 679–683, 2002.

L. M. Lin, D. Ricucci, J. Lin, and P. A. Rosenberg, “Nonsurgical root canal therapy of large cyst-like inflammatory periapical lesions and inflammatory apical cysts,” Journal of Endodontics, vol. 35, no. 5, pp. 607–615, 2009.

D. Ricucci, L. M. Lin, and L. S. W. Spångberg, “Wound healing of apical tissues after root canal therapy: a long-term clinical, radiographic, and histopathologic observation study,” Oral Surgery, Oral Medicine, Oral Pathology, Oral Radiology and Endodontology, vol. 108, no. 4, pp. 609–621, 2009.

D. Ricucci, J. F. Siqueira Jr., A. L. Bate, and T. R. P. Ford, “Histologic investigation of root canal-treated teeth with apical periodontitis: a retrospective study from twenty-four patients,” Journal of Endodontics, vol. 35, no. 4, pp. 493–502, 2009.

T. R. Dunavant, J. D. Regan, G. N. Glickman, E. S. Solomon, and A. L. Honeyman, “Comparative evaluation of endodontic irrigants against Enterococcus faecalis biofilms,” Journal of Endodontics, vol. 32, no. 6, pp. 527–531, 2006.

M. Haapasalo and Y. Shen, “Current therapeutic options for endodontic biofilms,” Endodontic Topics, vol. 22, no. 1, pp. 79–98, 2012.

J. Chandra, D. M. Kuhn, P. K. Mukherjee, L. L. Hoyer, T. McCormick, and M. A. Ghannoum, “Biofilm formation by the fungal pathogen Candida albicans: Development, architecture, and drug resistance,” Journal of Bacteriology, vol. 183, no. 18, pp. 5385–5394, 2001.

D. W. Norrington, J. Ruby, P. Beck, and P. D. Eleazer, “Observations of biofilm growth on human dentin and potential destruction after exposure to antibiotics,” Oral Surgery, Oral Medicine, Oral Pathology, Oral Radiology and Endodontology, vol. 105, no. 4, pp. 526–529, 2008.

L. E. Chávez de Paz, G. Bergenholtz, and G. Svensäter, “The effects of antimicrobials on endodontic biofilm bacteria,” Journal of Endodontics, vol. 36, no. 1, pp. 70–77, 2010.

E. C. McCoy, H. S. Rosenkranz, and R. Mermelstein, “Evidence for the existence of a family of bacterial nitroreductases capable of activating nitrated polycyclics to mutagens,” Environmental Mutagenesis, vol. 3, no. 4, pp. 421–427, 1981.

J. C. Nickel, J. B. Wright, I. Ruseska, T. J. Marrie, C. Whitfield, and J. W. Costerton, “Antibiotic resistance of Pseudomonas aeruginosa colonizing a urinary catheter in vitro,” European Journal of Clinical Microbiology, vol. 4, no. 2, pp. 213–218, 1985.

J. Jass, J. W. Costerton, and H. M. Lappin-Scott, “The effect of electrical currents and tobramycin on Pseudomonas aeruginosa biofilms,” Journal of Industrial Microbiology, vol. 15, no. 3, pp. 234–242, 1995.

A. Heydorn, A. T. Nielsen, M. Hentzer et al., “Quantification of biofilm structures by the novel computer program COMSTAT,” Microbiology, vol. 146, no. 10, pp. 2395–2407, 2000.

Y. Shen, S. Stojicic, and M. Haapasalo, “Antimicrobial efficacy of chlorhexidine against bacteria in biofilms at different stages of development,” Journal of Endodontics, vol. 37, no. 5, pp. 657–661, 2011.

J. W. Costerton, P. S. Stewart, and E. P. Greenberg, “Bacterial biofilms: a common cause of persistent infections,” Science, vol. 284, no. 5418, pp. 1318–1322, 1999.

Z. M. Thein, Y. H. Samaranayake, and L. P. Samaranayake, “In vitro biofilm formation of Candida albicans and non-albicans Candida species under dynamic and anaerobic conditions,” Archives of Oral Biology, vol. 52, no. 8, pp. 761–767, 2007.

A. R. Vieira, J. F. Siqueira Jr., D. Ricucci, and W. S. P. Lopes, “Dentinal tubule infection as the cause of recurrent disease and late endodontic treatment failure: a case report,” Journal of Endodontics, vol. 38, no. 2, pp. 250–254, 2012.

S. George, A. Kishen, and K. P. Song, “The role of environmental changes on monospecies biofilm formation on root canal wall by Enterococcus faecalis,” Journal of Endodontics, vol. 31, no. 12, pp. 867–872, 2005.

P. N. Ramachandran Nair, “Light and electron microscopic studies of root canal flora and periapical lesions,” Journal of Endodontics, vol. 13, no. 1, pp. 29–39, 1987.

A. Kishen and M. Hapassalo, “Biofilm models and methods of biofilm assessment,” Endodontic Topics, vol. 22, no. 1, pp. 58–78, 2010.