Optimized protocol for collecting root canal biofilms for in vitro studies

Optimized protocol for collecting root canal biofilms for in vitro studies | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Method Article Optimized protocol for collecting root canal biofilms for in vitro studies Rafael da Silva Goulart, Mariana Oliveira-Silva, Yara Teresinha Correa Silva-Sousa, and 2 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-4006763/v1 This work is licensed under a CC BY 4.0 License Status: Posted Version 1 posted You are reading this latest preprint version Abstract Aim The goal of this study was to standardize a new protocol for collecting biofilm from the interior of the root canal system (RCS) for in vivo testing. Methodology: In this study, 44 bovine incisors were used. The samples were divided into three experimental groups: 14 teeth, 12 for counting colony-forming units (CFU), and two samples for scanning electron microscopy (SEM). The first group was used for the biofilm collection protocol proposed here, the second group for the 2nd Biofilm Collection Protocol collection, and the third group for biofilm collection with an absorbent paper tip. Two additional teeth were used as sterilization controls to ensure that the experiments were free of contamination. The coronal region was removed and standardized at 15 mm. They were fitted with a Protaper up to the F5 insert, and the apical foramen was sealed with composite resin. The roots were stabilized with acrylic resin in a 1.5 mL Eppendorf tube. The specimens were sterilized and then inoculated with Enterococcus faecalis NTCT 775 every 24 h for 21 days. Following the period, each group underwent biofilm collection protocols, and CFU and scanning electron microscopy (SEM) data were analyzed. Shapiro–Wilk and one-way ANOVA tests were used to determine statistically significant differences between groups. Results The biofilm collection protocol group had the most CFUs, with extremely high values when compared with the other groups when converted to Log10. The results of the One-Way ANOVA test revealed that the 2nd collection protocol and absorbent paper tip collection groups were statistically similar (p > 0.05), whereas the biofilm collection protocol group was not. Conclusion The biofilm collection protocol proposed in this study was effective at collecting microorganisms from within the RCS. Compared to the biofilm collection protocol with paper cones, the in vivo collection protocol from bovine teeth yielded significantly more CFUs. Thus, the proposed protocol significantly increases the bacterial load of biofilms collected from the RCS sample, bringing the experiments closer to the reality of endodontic infections. Dentistry General Microbiology bacterial biofilms root canal system endodontic infection colony-forming units biofilm collection Figures Figure 1 Figure 2 Figure 3 Figure 4 1 INTRODUCTION The root canal system (RCS) has a complex structure that is characterized by the presence of various anatomical features, including accessory, lateral, apical delta, and collateral canals (Pagonis et al., 2010; Vera et al., 2012 ; Haapasalo et al., 2014 ; Wong et al., 2021 ). This structure is also distinguished by anatomical variations such as canal flattening, isthmuses, and indentations, which collectively prevent the effective removal of residual tissue materials and bacteria (Leoni et al., 2014 ; Versiani et al., 2016 ). Therefore, anatomical complexity promotes the formation of bacterial biofilms (Nair et al., 2005 ; Siqueira, Roças, 2008; Vera et al., 2012 ; Wong et al., 2021 ). Biofilms are microbial communities in which cells adhere to a substrate via an extracellular polymer matrix, altering growth phenotypes (Sadiq et al., 2022 ; Pandey et al., 2022 ). Within biofilm communities, microorganisms benefit from several favorable conditions, the most notable of which is protection from antimicrobial agents. The mechanisms underlying antimicrobial tolerance within biofilm-structured cells have been thoroughly investigated, revealing four distinct pathways. The first mechanism involves the matrix, which is primarily composed of exopolysaccharides (EPS) and serves as a natural antibiotic barrier. This matrix concentrates extracellular enzymes, including β-lactamase, which effectively inhibit the activity of β-lactam antibiotics (Anderl et al., 2000 ; Singh et al., 2021 ). The second is due to biofilm growth dynamics, whose microorganisms show slower growth rates than planktonic cells (He et al., 2024 ). Consequently, if nutrient availability is reduced, bacteria enter a latent or stationary growth phase, making them less vulnerable to nutritional cutback (Portenier et al., 2006 ; Singh et al., 2021 ). The third mechanism is related to oxygen metabolism, which is influenced by biofilm surface cells that can deplete available oxygen, resulting in anaerobic niches for the bacterial community (De Beer et al., 1994 ). This phenomenon has a differential effect on biofilm cells’ responses to certain antibiotics, such as aminoglycosides, which are more effective against aerobic bacteria (Tresse et al., 1995 ; Duggan, Sedgley, 2007 ). Finally, a persistent subpopulation of microorganisms, despite accounting for a minor fraction of the original population, can evade antimicrobial agents (Spoering, Lewis, 2001 ). The human oral cavity contains a large and diverse microbiome, with approximately 700 microbial species present in both forms of biofilm and planktonic cells (Paster et al., 2001 ; Valera et al., 2009 ; Zhao et al., 2017 ; Deo, Deshmukh., 2019). These microorganisms have varying penetration capacities into dentinal tubules, typically around 500 µm (Love, Jenkinson., 2002; Parmar et al., 2011 ; Brittan et al., 2016 ; Teoh et al., 2023 ). Located within the root canal, a conical-shaped structure with distinct diameters in its thirds, cervical (3.20 µm), middle (2.80 µm), and apical (2.11 µm), the dentinal tubules can suffer bacterial infiltration, which allows microorganisms to reach deeper dentin regions (Camargo et al., 2007 ; Ribeiro et al., 2010 ). It is worth highlighting that Enterococcus faecalis has a remarkable penetration ability, reaching depths of 800–1000 µm after three weeks of incubation (Haapasalo, Orstavik., 1987; Ran et al., 2014 ; Vatkar et al., 2016 ). This bacterium is significant as a nosocomial pathogen capable of causing various infections in humans (Jett, Huycke., 1994; Richards et al., 2000 ; Ali et al., 2017 ). Despite its small population in the oral microbiota, E. faecalis is important in the etiology of persistent periapical lesions (Evans et al., 2002 ; Sunde et al., 2002 ; Roças et al., 2004 ; Johnson et al., 2006 ; Arias-Moliz et al., 2009 ) and persistent apical periodontitis (Siqueira, Roças., 2022). Biofilm formation on the dentin walls of root canals is a common occurrence that can be prevented with biomechanical interventions (Stojicic et al., 2013 ; Neelakantan et al., 2017 ). However, E. faecalis is extremely resistant to biomechanical preparation and various intracanal medications, including calcium hydroxide (Vivacqua-Gomes et al., 2005 ; Dunavant et al., 2006 ; Abu Hasna et al., 2020 ). This microorganism has an exceptional ability to survive in alkaline pH conditions within dentinal tubules for extended periods (Kaufman et al., 2005 ; Berber et al., 2006 ; Gomes et al., 2006 ). Accordingly, to assess the efficacy of antimicrobial medications, the number of colonies forming units (CFUs) must be determined. To accomplish this, biofilm must be removed from the root canal system while microbial cells are still viable. This procedure usually involves the use of sterilized absorbent paper points, which is the same method used in the in vivo collection stage. Nonetheless, this approach has limitations, particularly in terms of achieving efficient biofilm dislodgement during the removal step. Thus, in the current study, a novel methodology was proposed to improve the displacement of mature E. faecalis biofilms from the root canal system, ensuring concomitant bacterial viability while minimizing potential lysis or cell death. 2 MATERIALS AND METHODS 2.1 Root canal preparation Bovine incisor teeth were preserved in a 0.1% thymol solution at 9°C and washed thoroughly with continuous exposure to running water for 24 h to remove any residual thymol solution. Macroscopic examination and mesiodistal radiography were then used to standardize the samples. Initially, a set of 44 teeth free of cracks, fractures, calcifications, and pronounced curvatures, each with a single canal and cervical diameter of 2–3 mm, was chosen (Brittan et al., 2016 ; Camargo et al., 2007 ; Ribeiro et al., 2010 ; Valera et al., 2009 ). The biomechanical preparation protocol began with perpendicular sectioning of the teeth, 15 mm above the root apex, using a carborundum disk attached to a straight handpiece and a low-speed motor (Beltec, Araraquara, SP, Brazil). The root canal was systematically explored with a K #30 file (Dentsply-Maillefer, Ballaigües, Switzerland) until the tip reached the apical foramen, which was then retracted by 1 mm to determine the working length (CT). The root apexes were sealed with Z100 composite resin (3M, Maplewood, Minnesota, USA) (Fig. 1 A), then 37% phosphoric acid etchant (Dentsply, York, Pennsylvania, USA) was applied to a height of 3 mm height above the apex for 20 s. The next steps were to rinse with water and dry the absorbent paper. Next, the Adper Single Bond adhesive system (3M, Maplewood, Minnesota, USA) was applied, followed by a 10 s air dry jet and a 40s light cure. Repeated adhesive system application was followed by an additional 40s light-curing cycle. This strategy aimed to keep adhesive material from entering the root canal via the apical foramen. The biomechanical preparation was done using the ProTaper Universal rotary system (Dentsply-Sirona, Petrópolis, Rio de Janeiro, Brazil) in the following order: SX, S1, S2, F1, F2, F3, F4, and F5. Nickel–titanium instruments were integrated into an electric motor contra-angle (VDW Silver, GmbH, Munich, Bavaria, Germany), which introduced the instruments into the canal via an insertion/withdrawal movement with an approximately 3 mm-controlled amplitude and imposed gentle canal wall pressure. Throughout the procedure, an alternated sequence of irrigation-aspiration steps were adopted at each instrument exchange, employing 2 mL of 2.5% NaOCl (BioFlora, Ribeirão Preto, São Paulo, Brazil) using a disposable 10 mL plastic syringe (BD, New Jersey, USA) and a Navy tip needle (Ultradent, South Jordan, Utah, USA). An aspiration cannula was also used, along with instrument cleaning and gauze. The final irrigation step involved a 5 min irrigation with 2 mL of 17% EDTA (Da Terra, Ribeirão Preto, São Paulo, Brazil), followed by 2 mL of 2.5% NaOCl and 10 mL of distilled water (Kaufman et al., 2005 ; Berber et al., 2006 ; Gomes et al., 2006 ). 2.2 Experimental groups The specimens were randomly divided into distinct experimental collection groups, each with 12 individual specimens for the quantification of colony-forming units. In each experimental group, a subgroup of two specimens was designated for subsequent scanning electron microscopy (SEM) analysis. PBS, a group consisting of two laterally cleaved specimens, was set aside for SEM analysis. It should be noted that the PBS group was not subjected to the biofilm collection protocol. This subgroup was chosen as a control group for comparison and validation purposes, serving as a positive control reference. Table 1 Experimental groups. Groups CFU (n) SEM (n) Biofilm Collection Protocol 12 2 2nd Biofilm Collection Protocol 12 2 Absorbent Paper Points 12 2 PBS 0 2 2.3 Preparation of specimens A set of 44 1.5 mL Eppendorf tubes was used to premold the roots. Thus, 1 mL of colorless acrylic resin (powder/liquid) (Artigos Odontológicos Clássico, LTDA, Campo Limpo Paulista, SP, Brazil) was injected into the tubes to immobilize the roots, resulting in the proof bodies (Fig. 1 B). The tubes were then sterilized at 121°C for 30 min before being immediately closed to prevent contamination. Before the immobilization step, a 2 mm longitudinal incision was made on both sides of the two roots in each experimental group to allow for subsequent cleavage for SEM analysis (Fig. 1 C). Finally, the 44 prepared specimens were loaded onto an Eppendorf tube rack (Fig. 1 D) and prepared for new sterilization in an autoclave (Phoenix Luferco, Araraquara, SP, Brazil) at 121°C for 30 min. The sterilization procedure was validated using chemical temperature indicator tape and microbiological tests. 2.4 Bacterial species and inoculum preparation The strain of E. faecalis NCTC 775 was chosen for this study due to its biofilm formation and survival in RCS. The strain was cryopreserved at − 80°C in cryotubes containing BHI broth and 15% glycerol. To reactivate the strain, it was aerobically grown in 5 mL of BHI at 37°C for 24 h. The strain was then grown in a Petri dish containing Müeller Hinton Agar (MH) (Sigma Aldrich, San Luis, MO, USA) under a laminar flow hood at 37°C for 24 h. To prepare the inoculum, bacterial colonies were collected from the MH Petri dish using a sterilized swab and transferred to a test tube with 2.5 mL of sterilized PBS until reaching a bacterial suspension equivalent to 2.0 McFarland scale turbidity (approximately 6.0 × 10 8 CFU/mL). Next, 2.5 mL of BHI medium (Sigma Aldrich, San Luis, MO, USA) was added to the suspension, resulting in a final concentration of approximately 3.0 × 10 8 CFU/mL, corresponding to 1.0 MacFarland scale. This step was necessary because the color of the culture medium prevented the McFarland scale from being prepared directly in it. To achieve the desired scale, the inoculum’s initial concentration was double the appropriate concentration. The initial inoculum was transferred to an equal volume of culture medium, achieving 1.0 on the MacFarland scale. Fresh bacterial cultures were prepared daily over 21 days to ensure the viability of the bacterial culture and to maintain the necessary inoculum volume for successful biofilm development (Jett et al., 1994 ; Ran et al., 2014 ). 2.5 Biofilm formation in the root canal Among the 44 bovine incisor tooth roots chosen, one was purposefully left uninoculated to serve as a control for the sterilization process during the trial period, filled with PBS only. The specimens’ root canals were then inoculated with 10 µL of bacterial suspension (3.0 × 10 8 CFU/mL) using aseptic techniques. The canals were carefully filled with micropipettes with sterilized tips to avoid inadvertent inoculum leakage and keep the external root surface clean. Following the inoculation step, the prepared roots were incubated for 24 h at 37°C. Following this period, the inoculum was extracted from the root canals. Simultaneously, the canals were gently washed with PBS, which was then aspirated and discarded to effectively remove nonviable bacterial cells. To ensure optimal biofilm growth, daily inoculation renewal was implemented, with a fresh inoculum introduced into the root canal every 24 h. This procedure allowed for viable bacterial inoculums, which aided in the development of a continuous biofilm throughout the RCS. Throughout the study, all samples were incubated in an aerobic environment at 37°C for 21 days. This time interval allows for the formation of a dense and fully mature biofilm within the root canal, simulating clinical conditions in vivo and making it suitable for further analysis and evaluation (Stuart et al., 2006 ; Vivacqua-Gomes et al., 2005 ). 2.6 Biofilm collection protocol The biofilm removal procedures (Fig. 2 ) were performed in a strict sequence of steps to ensure process accuracy. First, the specimens’ root canals were filled with sterilized PBS. Subsequently, PBS was carefully aspirated from the canals using a micropipette with sterilized tips. Following this step, renewed PBS was introduced into the canals to aid in the removal of any remaining planktonic cells. The specimens were then transferred to 5 mL Eppendorf tubes with conical bottoms and manipulated using sterilized tweezers. Root canals were carefully placed facing the bottom of the tubes, which were then tightly sealed. The specimens were then vigorously agitated in a vortex (Gehaka, São Paulo, SP, Brazil) at maximum speed for 1 min. This mechanical agitation detaches the biofilm that has formed on the root canal walls. The next step of the biofilm collection process involved placing 5 mL Eppendorf tubes containing the specimens on a rack and immersing them in an ultrasonic bath (Digital Ultrasonic Cleaner, Kondortech, São Carlos, SP, Brazil) at room temperature (approximately 25° C). This set was immersed in an ultrasonic bath for 480 s to ensure that the biofilm was completely removed from the root canal walls. To remove the detached biofilm from the interior of the root canal, centrifugation was performed at 700 g for 1 min. Finally, the pellet formed at the bottom of the 5 mL Eppendorf was collected for further counting of CFU. The protocol was standardized to maintain bacterial viability, as demonstrated by Robertson et al., 2019 , who demonstrated bacterial viability while using a vortex at maximum power and centrifugation at 7000 g for 7 min. 2.7 Second Biofilm Collection Following the collection of RCS biofilms, the same specimens were subjected to a new collection protocol using the same protocol. This strategy was used to determine whether there would be a significant amount of biofilm remaining from the first collection, as well as whether the proposed protocol would require only one or two collections. The biofilms collected in both protocols were quantified by counting CFU. The difference between the first and second biofilm collections was statistically determined, as described below. 2.8 Absorbent paper point collection For comparison with the new proposed method, biofilms were collected from the RCS using the absorbent paper point method described by Alfirdous et al. ( 2022 ), with adjustments for the number of paper points. The number of paper tips used for collection increased in proportion to the diameter of the root canal, ranging from one and three units. Before inserting the paper points into the root canals, each one was saturated with PBS. After the paper points were extracted from the root canals, PBS was drained from them using a pipette with sterilized tips. This procedure eliminated any planktonic bacteria that were present inside the canals. Following this preliminary step, the absorbent paper points were resoaked with fresh PBS. Five absorbent paper points with appropriate working diameters (taper 50) were inserted sequentially into the root canals. This strategy ensured that the points were consistently and uniformly placed within the root canal structure. All biofilm-soaked points were placed in 1.5 mL Eppendorf tubes containing 1 mL of PBS and mixed thoroughly in a vortex mixer for 1 min. These tubes, designed to facilitate subsequent specimen manipulation and analysis, served as storage vessels for subsequent steps, such as serial dilution followed by counting CFUs. 2.9 Microbiological analysis 2.9.1 Colony-forming units (CFU) Biofilms extracted from all experimental groups’ root canals were serially diluted at a ratio of 1:9. According to this protocol, aliquots of 0.1 mL of each suspension were aseptically inoculated into Petri dishes containing MH Agar medium. This inoculation was performed using a Drigalski spatula and the spread plate methodology (Tortora et al., 2018 ). The Petri dishes were then incubated under controlled conditions at 37°C for 24 h. After incubation, the colony-forming units (CFU) per square millimeter (CFU/ mm²) were carried out (Tortora et al., 2018 ). To count CFU, the image analysis software ImageJ (Schneider et al., 2012 ) was employed. To aid analysis, the photograph corresponding to the last dilution was taken. Then, the ImageJ software imported this photograph and converted it to an 8-bit resolution image. Calibration was performed using a 180/250 correlation and a black background was selected. The next step involved isolating a specific region using the circumference option. The analysis focused on particles ranging from 10 to 130 pixels². Consequently, the number of colonies was expressed in pixels2. CFU counting can also be done visually with trained and calibrated readers; however, using software greatly reduces the possibility of errors. 2.9.2 Scanning electron microscopy analysis of bacterial adhesion Two specimens from each experimental group were designated for SEM analysis. Before the SEM analysis, slits were precisely cut into distal and mesial regions of the specimens using a chisel. In the next step, the specimens were cleaved into two halves with a chisel for further analysis. The prepared specimen halves were immersed in a series of ethanol solutions at increasing concentrations to facilitate dehydration. This set featured ethanol concentrations of 30%, 50%, 70%, 80%, 85%, and 100%. The specimens were immersed in each solution for 1 h, for a total of two hours in absolute ethanol. After dehydration, the specimens were ready for SEM analysis (Huang et al., 2017; Li et al., 2018). The specimens were appropriately labeled and mounted on cylindrical structures known as stubs. Double-sided carbon adhesive tape was used to hold the specimens in place on the stubs. A fine coating of electrically conductive metal, typically gold or gold–palladium alloy, was then applied to the specimens using vacuum metallization equipment (SDC 050, Bal-Tec AG, Liechtenstein). This final step was executed to ensure optimal conductivity and structural integrity during the SEM analysis, with specific parameters properly adjusted to ensure precise data acquisition and interpretation. These parameters include 0.01 mbar of pressure, 40 mA of current to facilitate electron emission and interaction with the sample, 50 mm of working distance, and a coverage time of 110 s, which determines the duration of the interaction between the sample and the electron beam. A subsequent metal deposition step with an average thickness of 20–30 nm ensured a controlled and uniform metal coating. After sample preparation, the specimens were transferred to a scanning electron microscope (JSM 5410, JEOL Ltd, Tokyo, Japan) for examination. The specimens’ apical, middle, and cervical thirds were selected as areas of interest. To capture varying levels of detail, magnifications of 500, 2000, and 5000x were used. The acquired images were analyzed with a focus on two distinct aspects. First, the assessment confirmed the extension of bacterial adhesion to dentinal tubules, which provided information about interactions at the biofilm–tubule interface. Second, the study assessed the effectiveness of biofilm removal techniques in the root canal. 2.10 Statistical Analysis For statistical analysis, the Shapiro–Wilk test was used to verify data distribution. It was discovered that measurements were not normally distributed and thus were transformed into log 10 values before the one-way parametric method was used to compare the groups. The significance level was set to 5%. 3 RESULTS 3.1 Evaluation of Collection Efficiency Considering CFU Counting The evaluation of collection efficiency produced comparable results in terms of CFU counting for all experimental groups (Table 2 ). Surprisingly, the collection conducted under the group biofilm collection protocol showed higher CFU values. Conversely, the absorbent paper point collection and 2° biofilm collection protocols produced significantly lower CFU values (Fig. 3 ). It is worth noting that the biofilm collection protocol effectively removed the biofilm from the root canal, with only a few bacteria found inside the dentinal tubules. Therefore, we believe that the second collection is unnecessary to ensure the experiments’ effectiveness. Table 2 Number of colonies forming units (CFU) in different biofilm collection protocols. Distinct letters in the same column indicate statistically significant differences between the groups. Experimental groups The number of colonies forming units (CFU) log 10 Biofilm Collection Protocol 7.38 ± 6.40 A 2nd Biofilm Collection Protocol 6.20 ± 5.67 B Absorbent Paper Point Collection 6.05 ± 5.68 B 3.2 Analysis of Collected Biofilms by Scanning Electron Microtomography (SEM) Qualitative evaluation of photomicrographs obtained through SEM analysis revealed significant differences among all experimental groups, each distinguished by the specific collection techniques used. The PBS control and absorbent paper point collection groups revealed dense biofilm at the periphery of the dentinal tubules. In stark contrast, the proposed biofilm collection protocol effectively removed bacterial biofilm content from within the dentinal tubules. Notably, the absorbent paper point group’s collection showed a significant accumulation of biofilm adhering across all assessed regions, which was similar to the results observed in the control group (Fig. 4 ). 4 DISCUSSION Biofilms are closely associated with RCS infections, and their complexity, combined with the nature of biofilms, makes disinfection of this system extremely difficult. Much of the research in the field of endodontics has focused on the characterization of microbial biofilms formed in RCS, as well as clinical methods for killing and eliminating them. Biofilms must be studied in endodontics microbiology to understand the pathogenic potential of the microorganisms involved in the infection and to evaluate new disinfection strategies for RCS (Jhajharia et al., 2015 ; Neelakantan et al., 2017 ). Recent research has shown that biofilm development is complex, requiring a delicate balance of bacterial proliferation and the production of extracellular polymeric substances (EPS). Therefore, the treatment of biofilm infections is challenging and requires strategies tailored to the stage of biofilm development (He et al., 2024 ). Traditionally, bacterial biofilms are removed from root canals using sterilized absorbent paper points, allowing bacteria absorption. The proposed method aims to simulate the in vivo collection procedure (Alfirdous et al., 2022 ). It is important to note that in vivo investigations gather a distinct set of RCS decontamination protocols involving physical process instrumentation (Alfirdous et al., 2022 ) and chemical methods using irrigating solutions with or without activation (Căpută et al., 2019 ; Susila, Minu, 2019 ). Nonetheless, certain areas of the RCS present challenges to effective paper point absorption. Therefore, to bring the experimental approach closer to the clinical reality of endodontic infections, a methodology for optimal collection of microbial biofilms in RCS was proposed. To propose the biofilm collection protocol in the RCS, several parameters were evaluated and consulted in the literature, including the preservation of cell viability when the biofilms were subjected to various experimental steps. The cellular viability of the collected biofilms was assessed individually in three stages: after vortexing and ultrasonic bath protocols, and then after collection centrifugation. Finally, the cell viability of the biofilms was assessed following their sequential exposure to the three processes. The study found bacterial viability in all phases tested (data not shown). Robertson et al. ( 2019 ) reported similar results, using vortexing and centrifugation at 7,000 g for 7 min to maintain bacterial viability. The authors used the Live/Dead kit to determine the number of dead and living cells. Chen et al. ( 2021 ) achieved similar results by centrifugation at 5,000 g for 15 min to form pellets for analysis while preserving bacterial viability. The results from the collection using absorbent paper points and the second collection after the initial collection step were statistically similar. On the other hand, the initial biofilm collection protocol had significantly higher CFU values than the other two protocols. The CFU values discovered in the second collection are statistically insignificant for analysis, indicating that the second collection is irrelevant in terms of CFU values when considering the cost, time, and labor required for its execution. These findings highlight the effectiveness of the proposed collection method when compared to collection using absorbent paper points. The traditional method of collecting biofilm using sterilized absorbent paper points has limitations in accessing the entire root canal structure, resulting in flaws in the bacterial sampling step (Swimberghe, Coenye, et al., 2019; Hoedke et al., 2021 ; Alfirdous et al., 2022 ). The inherent limitation of this method is its tendency to predominantly capture planktonic bacteria while failing to effectively retrieve biofilm-associated bacteria (Neelakantan et al., 2015 ; Neuhaus et al., 2016 ). To address this limitation, a new collection protocol that includes vortex mixing, ultrasonic bath immersion, and centrifugation is proposed. The primary goal of this research is to develop a collection methodology that not only ensures the effective removal of biofilm material from the radicular canal but also protects the viability and integrity of bacterial cells by preventing lysis and death. In this regard, it is a pioneering study because the proposal addresses both the efficiency of material removal from the radicular canal using a three-step process and longterm bacterial viability. To demonstrate the efficacy and reliability of the proposed collection protocol, a comparison with a consolidated method was conducted. It is also important to note that the variability and often subjective nature of operator skills can have a significant impact on the outcomes of various collection methods. Thereby, the operator’s skills make an expressive contribution to potential discrepancies in results. To achieve reliable and unbiased results, the ideal collection method must be highly reproducible, separating the obtained results from operator-dependent factors. Conversely, techniques that use instruments to mechanically dislodge biofilms run the risk of lysing bacterial cells, compromising the accuracy of the results. In contrast, the centrifuge-based collection protocol used in this study proved advantageous because it avoids bacterial lysis (Chen et al., 2021 ). The methodology used allows for the preservation of cell viability during the detachment of biofilm material from the root walls, as demonstrated in the findings of this study. It is important to note that there are several classic and contemporary techniques for removing biofilms from the RCS to eliminate pathogenic microorganisms and, as a result, disinfecting the canal in the practice of endodontic therapy (Josic et al., 2022 ). However, the goal of this article is not to evaluate RCS disinfection; rather, it is to provide researchers with an experimental model so that such disinfection strategies can be evaluated in vivo with greater representativeness of biofilm collection, reproducibility, and without interfering with biofilm viability, thereby avoiding undesirable results. Comparative analysis of SEM images provided valuable qualitative information about the efficacy of the various collection methods tested. There is a similarity in terms of CFU when comparing the absorbent paper point group to the second collect protocol group, which collects proc planktonic bacteria preferentially. Qualitative SEM analyses revealed the presence of bacterial cells in the dentinal tubules following the paper tip collection process, whereas the proposed protocol revealed a significant lack of bacteria in these regions. These findings are significant because many protocols for disinfecting endodontic canals have varying effectiveness in penetrating dental tubules (Josic et al., 2022 ). Therefore, the effective in vivo evaluation of these protocols via CFU counting is dependent on the efficient collection of biofilms in these deeper regions. 5 CONCLUSION In conclusion, this study found that a new protocol for collecting biofilm from RCS in vivo was effective. The current study supports the efficacy of the devised collection protocol over the conventional paper point method, establishing it as the best approach for thoroughly retrieving bacterial biofilm from within the RCS. The protocol’s robust performance is highlighted by its inherent reproducibility and avoidance of operator-dependent variations. Visual analysis with scanning electron microscopy provides compelling evidence, revealing almost no bacterial presence following collection. It should be noted that the proposed protocol has the advantage of being able to collect biofilm from deeper areas of RCS, which allows it to be used in different experimental protocols. However, in this study, teeth with anatomical variations in the RCS were not evaluated during biofilm collection, which may limit future studies of teeth with such characteristics. DECLARATIONS Author Contributions Rafael da Silva Goulart: methodology (lead); conceptualization (lead); writing–original draft (lead); formal analysis (lead); writing–review and editing (equal). Mariana Oliveira-Silva: methodology (supporting) and review and editing (supporting). Yara Teresinha Correa Silva Sousa: conceptualization (supporting)and review and editing (equal). Carlos Eduardo Saraiva Miranda: conceptualization (lead); writing–original draft (lead); writing–review and editing (equal). André Pitondo-Silva: conceptualization (lead); writing–original draft (lead); writing–review and editing (equal). Data Availability No data were used to support this study. Conflicts of Interest The authors declare no conflicts of interest. Ethical approval Ethical approval for the use of bovine teeth in this study was obtained from the Animal Use Ethics Committee of the University of Ribeirão Preto (Ribeirão Preto, SP, Brazil) [approval number 01/2017]. There was no patient involvement in the study. Funding Rafael da Silva Goulart is a Ph.D. student fellow of Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES); therefore, this study was financed in part by the CAPES (grant no 88887.493929/2020-00). Acknowledgments This work was reviewed by a native English speaker with expertise in Endodontics from the ENAGO (https://www.enago.com.br/). REFERENCES Abu Hasna, A., Khoury, R. D., Toia, C. C., Gonçalves, G. B., de Andrade, F. B., Talge Carvalho, C. A., Ribeiro Camargo, C. H., & Carneiro Valera, M. (2020). In vitro Evaluation of the Antimicrobial Effect of N-acetylcysteine and Photodynamic Therapy on Root Canals Infected with Enterococcus faecalis. Iranian endodontic journal, 15(4), 236–245. Alfirdous, R. A., Alquiria, T. A., Jacinto, R. C., & Martinho, F. C. (2022). 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Method for concentrating viable microorganisms for microbial load determination and eliminating uncertainty from matrix effects from urine and whole blood. MethodsX, 8, 101451. De Beer, D., Stoodley, P., Roe, F., & Lewandowski, Z. (1994). Effects of biofilm structures on oxygen distribution and mass transport. Biotechnology and Bioengineering, 43(11), 1131–1138. Deo, P. N., & Deshmukh, R. (2019). Oral microbiome: Unveiling the fundamentals. Journal of oral and maxillofacial pathology, 23(1), 122–128. Duggan, J. M., & Sedgley, C. M. (2007). Biofilm formation of oral and endodontic Enterococcus faecalis . Journal of endodontics, 33(7), 815–818. Dunavant, T. R., Regan, J. D., Glickman, G. N., Solomon, E. S., & Honeyman, A. L. (2006). Comparative evaluation of endodontic irrigants against Enterococcus faecalis biofilms. Journal of endodontics, 32(6), 527–531. Evans, M., Davies, J. K., Sundqvist, G., & Figdor, D. (2002). 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Kaufman, B., Spångberg, L., Barry, J., & Fouad, A. F. (2005). Enterococcus spp. in endodontically treated teeth with and without periradicular lesions. Journal of endodontics, 31(12), 851–856. Leoni, G. B., Versiani, M. A., Pécora, J. D., & Damião de Sousa-Neto, M. (2014). Micro–Computed Tomographic Analysis of the Root Canal Morphology of Mandibular Incisors. Journal of Endodontics, 40(5), 710–716. Love, R. M., & Jenkinson, H. F. (2002). Invasion of dentinal tubules by oral bacteria. Critical reviews in oral biology and medicine: an official publication of the American Association of Oral Biologists, 13(2), 171–183. Nair, P. N. R., Henry, S., Cano, V., & Vera, J. (2005). 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, 99(2), 231–252. Neelakantan, P., Cheng, C. Q., Mohanraj, R., Sriraman, P., Subbarao, C., & Sharma, S. (2015). Antibiofilm activity of three irrigation protocols activated by ultrasonic, diode laser or Er:YAG laser in vitro . International endodontic journal, 48(6), 602–610. Neelakantan, P., Romero, M., Vera, J., Daood, U., Khan, A. U., Yan, A., & Cheung, G. S. P. (2017). Biofilms in Endodontics-Current Status and Future Directions. International journal of molecular sciences, 18(8), 1748. Neuhaus, K. W., Liebi, M., Stauffacher, S., Eick, S., & Lussi, A. (2016). Antibacterial Efficacy of a New Sonic Irrigation Device for Root Canal Disinfection. Journal of endodontics, 42(12), 1799–1803.. Pagonis, T. C., Chen, J., Fontana, C. R., Devalapally, H., Ruggiero, K., Song, X., … Yamazaki, H. (2010). Nanoparticle-based Endodontic Antimicrobial Photodynamic Therapy. Journal of Endodontics, 36(2), 322–328. Pandey, R. P., Mukherjee, R., & Chang, C. M. (2022). Emerging Concern with Imminent Therapeutic Strategies for Treating Resistance in Biofilm. Antibiotics (Basel, Switzerland), 11(4), 476. Parmar, D., Hauman, C. H. J., Leichter, J. W., McNaughton, A., & Tompkins, G. R. (2011). Bacterial localization and viability assessment in human ex vivo dentinal tubules by fluorescence confocal laser scanning microscopy. International Endodontic Journal, 44(7), 644–651. Paster, B. J., Boches, S. K., Galvin, J. L., Ericson, R. E., Lau, C. N., Levanos, V. A., Dewhirst, F. E. (2001). Bacterial Diversity in Human Subgingival Plaque. Journal of Bacteriology, 183(12), 3770–3783. Portenier, I., Waltimo, T., Ørstavik, D., & Haapasalo, M. (2006). Killing of Enterococcus faecalis by MTAD and chlorhexidine digluconate with or without cetrimide in the presence or absence of dentine powder or BSA. Journal of endodontics, 32(2), 138–141. Ran, S., Wang, J., Jiang, W., Zhu, C., & Liang, J. (2014). Assessment of dentinal tubule invasion capacity of Enterococcus faecalis under stress conditions ex vivo. International Endodontic Journal, 48(4), 362–372. Ribeiro, R. G., Marchesan, M. A., Silva, R. G., Sousa-Neto, M. D., & Pécora, J. D. (2010). Dentin permeability of the apical third in different groups of teeth. Brazilian dental journal, 21(3), 216–219.. Richards, M. J., Edwards, J. R., Culver, D. H., & Gaynes, R. P. (2000). Nosocomial infections in combined medical-surgical intensive care units in the United States. Infection control and hospital epidemiology, 21(8), 510–515. Roças, I., Siqueira Jr, J., & Santos, K. (2004). Association of Enterococcus faecalis With Different Forms of Periradicular Diseases. Journal of Endodontics, 30(5), 315–320. Sadiq, F. A., Hansen, M. F., Burmølle, M., Heyndrickx, M., Flint, S., Lu, W., Chen, W., & Zhang, H. (2022). Trans-kingdom interactions in mixed biofilm communities. FEMS microbiology reviews, 46(5), fuac024. Schneider, C. A., Rasband, W. S., & Eliceiri, K. W. (2012). NIH Image to ImageJ: 25 years of image analysis. Nature methods, 9(7), 671-675. Singh, S., Datta, S., Narayanan, K. B., & Rajnish, K. N. (2021). Bacterial exo-polysaccharides in biofilms: role in antimicrobial resistance and treatments. Journal, genetic engineering & biotechnology, 19(1), 140. Siqueira, J. F., Jr, & Rôças, I. N. (2008). Clinical implications and microbiology of bacterial persistence after treatment procedures. Journal of endodontics, 34(11), 1291–1301. Siqueira, J. F., Jr, & Rôças, I. N. (2022). Present status and future directions: Microbiology of endodontic infections. International endodontic journal, 55 Suppl 3, 512–530. Spoering, A. L. & Lewis, K. (2001). Biofilms and Planktonic Cells of Pseudomonas aeruginosa Have Similar Resistance to Killing by Antimicrobials. Journal of Bacteriology, 183(23), 6746–6751. Stojicic, S., Amorim, H., Shen, Y., & Haapasalo, M. (2013). Ex vivo killing of Enterococcus faecalis and mixed plaque bacteria in planktonic and biofilm culture by modified photoactivated disinfection. International endodontic journal, 46(7), 649–659. Stuart, C. H., Schwartz, S. A., Beeson, T. J., & Owatz, C. B. (2006). Enterococcus faecalis : its role in root canal treatment failure and current concepts in retreatment. Journal of endodontics, 32(2), 93–98. Sunde, P. T., Olsen, I., Debelian, G. J., & Tronstad, L. (2002). Microbiota of periapical lesions refractory to endodontic therapy. Journal of endodontics, 28(4), 304–310. Susila, A. & Minu, J. (2019). Activated Irrigation vs. Conventional non-activated Irrigation in Endodontics - A Systematic Review. European endodontic journal, 4(3), 96–110. Teoh, Y. Y., Liew, K. Y., Siao, J., Wong, S., Chandler, N., & Bogen, G. (2023). The effects of chelation on the intratubular penetration depth of mineral trioxide aggregate. Australian endodontic journal : the journal of the Australian Society of Endodontology Inc, 10.1111/aej.12766. Tortora, G. J., Funke, B. R., & Case, C. L. (2018). Microbiology an introduction 13th edition. Tresse, O., Jouenne, T., & Junter, G.-A. (1995). The role of oxygen limitation in the resistance of agar-entrapped, sessile-like Escherichia coli to aminoglycoside and β-lactam antibiotics. Journal of Antimicrobial Chemotherapy, 36(3), 521–526. Valera, M. C., Silva, K. C. G. da, Maekawa, L. E., Carvalho, C. A. T., Koga-Ito, C. Y., Camargo, C. H. R., & Lima, R. S. e. (2009). Antimicrobial activity of sodium hypochlorite associated with intracanal medication for Candida albicans and Enterococcus faecalis inoculated in root canals. Journal of Applied Oral Science, 17(6), 555–559. Vatkar, N. A., Hegde, V., & Sathe, S. (2016). Vitality of Enterococcus faecalis inside dentinal tubules after five root canal disinfection methods. Journal of conservative dentistry : JCD , 19 (5), 445–449. Vera, J., Siqueira, J. F., Jr, Ricucci, D., Loghin, S., Fernández, N., Flores, B., & Cruz, A. G. (2012). One- versus two-visit endodontic treatment of teeth with apical periodontitis: a histobacteriologic study. Journal of endodontics, 38(8), 1040–1052. Versiani, M. A., Ordinola-Zapata, R., Keleş, A., Alcin, H., Bramante, C. M., Pécora, J. D., & Sousa-Neto, M. D. (2016). Middle mesial canals in mandibular first molars: A micro-CT study in different populations. Archives of Oral Biology, 61, 130–137. Vivacqua-Gomes, N., Gurgel-Filho, E. D., Gomes, B. P., Ferraz, C. C., Zaia, A. A., & Souza-Filho, F. J. (2005). Recovery of Enterococcus faecalis after single- or multiple-visit root canal treatments carried out in infected teeth ex vivo. International endodontic journal, 38(10), 697–704. Wong J, Manoil D, Näsman P, Belibasakis GN, Neelakantan P. (2021). Microbiological Aspects of Root Canal Infections and Disinfection Strategies: An Update Review on the Current Knowledge and Challenges. Front Oral Health.;2:672887. Zhao, H., Chu, M., Huang, Z., Yang, X., Ran, S., Hu, B., Zhang, C., & Liang, J. (2017). Variations in oral microbiota associated with oral cancer. Scientific reports, 7(1), 11773. Chen J, Tomasek M, Nuñez E, Gau V. Method for concentrating viable microorganisms for microbial load determination and eliminating uncertainty from matrix effects from urine and whole blood. MethodsX. 2021;8:101451. Robertson, J., McGoverin, C., Vanholsbeeck, F., Swift, S.. Optimisation of the protocol for the LIVE/DEAD® BacLightTM bacterial viability kit for rapid determination of bacterial load. Frontiers in microbiology. 10, 801. 2019 Additional Declarations The authors declare no competing interests. Cite Share Download PDF Status: Posted Version 1 posted You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. 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Also discoverable on Platform About Our Team In Review Editorial Policies Advisory Board Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {"props":{"pageProps":{"initialData":{"identity":"rs-4006763","acceptedTermsAndConditions":true,"allowDirectSubmit":true,"archivedVersions":[],"articleType":"Method Article","associatedPublications":[],"authors":[{"id":275984069,"identity":"3b15ab57-0cc5-4528-b937-9d25c4390e76","order_by":0,"name":"Rafael da Silva Goulart","email":"","orcid":"https://orcid.org/0000-0003-1822-4865","institution":"Programa de Pós-Graduação em Odontologia, Universidade de Ribeirão Preto, Brazil.","correspondingAuthor":false,"prefix":"","firstName":"Rafael","middleName":"da Silva","lastName":"Goulart","suffix":""},{"id":275984070,"identity":"10f609f0-34a8-47f7-927d-a6fcb73e727d","order_by":1,"name":"Mariana Oliveira-Silva","email":"","orcid":"https://orcid.org/0000-0003-0459-8284","institution":"Programa de Pós-Graduação em Tecnologia Ambiental, Universidade de Ribeirão Preto, Brazil.","correspondingAuthor":false,"prefix":"","firstName":"Mariana","middleName":"","lastName":"Oliveira-Silva","suffix":""},{"id":275984071,"identity":"b6fa56e2-ee25-4c3f-96ad-aa7c78006153","order_by":2,"name":"Yara Teresinha Correa Silva-Sousa","email":"","orcid":"https://orcid.org/0000-0002-7671-1656","institution":"Programa de Pós-Graduação em Odontologia, Universidade de Ribeirão Preto, Brazil.","correspondingAuthor":false,"prefix":"","firstName":"Yara","middleName":"Teresinha Correa","lastName":"Silva-Sousa","suffix":""},{"id":275984072,"identity":"a6a60bf6-3e4f-4b1a-9c28-c2c4ccb2ec8f","order_by":3,"name":"Carlos Eduardo Saraiva Miranda","email":"","orcid":"https://orcid.org/0000-0003-0409-2523","institution":"Programa de Pós-Graduação em Odontologia, Universidade de Ribeirão Preto, Brazil.","correspondingAuthor":false,"prefix":"","firstName":"Carlos","middleName":"Eduardo Saraiva","lastName":"Miranda","suffix":""},{"id":275984073,"identity":"610f27ac-84aa-4755-9da1-1a9a0f26b21a","order_by":4,"name":"André Pitondo-Silva","email":"data:image/png;base64,iVBORw0KGgoAAAANSUhEUgAAAZAAAAAyAQMAAABI0h/eAAAABlBMVEX///8AAABVwtN+AAAACXBIWXMAAA7EAAAOxAGVKw4bAAAAzUlEQVRIiWNgGAWjYHACAwaGCgYeEEsCIsBDjJYzDDw8pGlhbIOoI04L/+zmbR8+zrsjY8/ee/DGBwY7Od0G3mMf8GmRuHOseObMbc94eHjOJVvOYEg2NjvAlzwDrzU3coyZebcd5uGRyDGT5mE4kLjtAI8xXh3yIC1/55CixQCkhbGBFC2GN9KKGXuOAf1y5oyx5QwDoF8O8yXj1SJ3I3kzw4+aO/bs7T2GNz5U2MmZHe89jFcLFByAuROImYnRgNAyCkbBKBgFowALAADk9UMBDdrOzAAAAABJRU5ErkJggg==","orcid":"https://orcid.org/0000-0003-0098-9667","institution":"Programa de Pós-Graduação em Odontologia, Universidade de Ribeirão Preto, Brazil.","correspondingAuthor":true,"prefix":"","firstName":"André","middleName":"","lastName":"Pitondo-Silva","suffix":""}],"badges":[],"createdAt":"2024-03-02 15:20:43","currentVersionCode":1,"declarations":{"humanSubjects":false,"vertebrateSubjects":false,"conflictsOfInterestStatement":false,"humanSubjectEthicalGuidelines":false,"humanSubjectConsent":false,"humanSubjectClinicalTrial":false,"humanSubjectCaseReport":false,"vertebrateSubjectEthicalGuidelines":false},"doi":"10.21203/rs.3.rs-4006763/v1","doiUrl":"https://doi.org/10.21203/rs.3.rs-4006763/v1","draftVersion":[],"editorialEvents":[],"editorialNote":"","failedWorkflow":false,"files":[{"id":51939248,"identity":"0f5d8e07-bfa7-4357-9885-087cb4284d20","added_by":"auto","created_at":"2024-03-04 08:25:06","extension":"png","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":407721,"visible":true,"origin":"","legend":"\u003cp\u003eRoot of bovine dental element apically sealed (A); specimen mounted with acrylic resin for immobilization (B); specimen with slit for cleavage highlighted (C); Specimens immobilized and arranged in an Eppendorf tube rack (D).\u003c/p\u003e","description":"","filename":"image1.png","url":"https://assets-eu.researchsquare.com/files/rs-4006763/v1/49f2a967061830b9a2795c30.png"},{"id":51939515,"identity":"d8459412-eb1b-4723-9bb7-523a7778b908","added_by":"auto","created_at":"2024-03-04 08:33:06","extension":"png","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":143131,"visible":true,"origin":"","legend":"\u003cp\u003eBiofilm protocol collects: A- Sample; B- Removed root from the sample and the root filled with PBS; C- Transferred the root for a 5 mL Eppendorf with the canal entrance facing downward; D- Vortex at maximum power for 1 min; E- After the vortex, the samples are transferred to ultrasonic cub for 480 s; F- Centrifuged 700 g for 1 min.\u003c/p\u003e","description":"","filename":"image2.png","url":"https://assets-eu.researchsquare.com/files/rs-4006763/v1/8fe9e6f6649ec802fcfcbdcb.png"},{"id":51939249,"identity":"fdb74742-9ea0-4168-96e7-1a7b34a8fcbd","added_by":"auto","created_at":"2024-03-04 08:25:06","extension":"png","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":653916,"visible":true,"origin":"","legend":"\u003cp\u003eColony-Forming Unit in Muller Hinton plates related to the first dilution: A, Biofilm Collection Protocol; B,2° Biofilm Collection Protocol; C, Absorbent Paper Point Collection.\u003c/p\u003e","description":"","filename":"image3.png","url":"https://assets-eu.researchsquare.com/files/rs-4006763/v1/e8d09ece0f6fc2447633f296.png"},{"id":51939247,"identity":"396b69d7-1ef3-4b37-9f88-a72b8a7c6934","added_by":"auto","created_at":"2024-03-04 08:25:06","extension":"png","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":453107,"visible":true,"origin":"","legend":"\u003cp\u003ePhotomicrographs (5000x) of groups and the presence of biofilm:(A), PBS control; (B) Absorbent paper point collection; (C) Biofilm collection protocol; (D) PBS control long-axis dentinal tubule; (F) Absorbent paper point collection protocol long-axis dentinal tubule. (F) Biofilm collection protocol for the long-axis dentinal tubule.\u003c/p\u003e","description":"","filename":"image4.png","url":"https://assets-eu.researchsquare.com/files/rs-4006763/v1/05a1da046696f39907174190.png"},{"id":51939644,"identity":"9d55e68a-f0e4-4beb-828d-0e3acfc07640","added_by":"auto","created_at":"2024-03-04 08:41:07","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":2115187,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-4006763/v1/40883f57-f53a-47ff-8c5b-eb587a1782eb.pdf"}],"financialInterests":"The authors declare no competing interests.","formattedTitle":"\u003cp\u003e\u003cstrong\u003eOptimized protocol for collecting root canal biofilms for in vitro studies\u003c/strong\u003e\u003c/p\u003e","fulltext":[{"header":"1 INTRODUCTION","content":"\u003cp\u003eThe root canal system (RCS) has a complex structure that is characterized by the presence of various anatomical features, including accessory, lateral, apical delta, and collateral canals (Pagonis et al., 2010; Vera et al., \u003cspan citationid=\"CR56\" class=\"CitationRef\"\u003e2012\u003c/span\u003e; Haapasalo et al., \u003cspan citationid=\"CR18\" class=\"CitationRef\"\u003e2014\u003c/span\u003e; Wong et al., \u003cspan citationid=\"CR59\" class=\"CitationRef\"\u003e2021\u003c/span\u003e). This structure is also distinguished by anatomical variations such as canal flattening, isthmuses, and indentations, which collectively prevent the effective removal of residual tissue materials and bacteria (Leoni et al., \u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e2014\u003c/span\u003e; Versiani et al., \u003cspan citationid=\"CR57\" class=\"CitationRef\"\u003e2016\u003c/span\u003e). Therefore, anatomical complexity promotes the formation of bacterial biofilms (Nair et al., \u003cspan citationid=\"CR28\" class=\"CitationRef\"\u003e2005\u003c/span\u003e; Siqueira, Ro\u0026ccedil;as, 2008; Vera et al., \u003cspan citationid=\"CR56\" class=\"CitationRef\"\u003e2012\u003c/span\u003e; Wong et al., \u003cspan citationid=\"CR59\" class=\"CitationRef\"\u003e2021\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eBiofilms are microbial communities in which cells adhere to a substrate via an extracellular polymer matrix, altering growth phenotypes (Sadiq et al., \u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e2022\u003c/span\u003e; Pandey et al., \u003cspan citationid=\"CR33\" class=\"CitationRef\"\u003e2022\u003c/span\u003e). Within biofilm communities, microorganisms benefit from several favorable conditions, the most notable of which is protection from antimicrobial agents. The mechanisms underlying antimicrobial tolerance within biofilm-structured cells have been thoroughly investigated, revealing four distinct pathways. The first mechanism involves the matrix, which is primarily composed of exopolysaccharides (EPS) and serves as a natural antibiotic barrier. This matrix concentrates extracellular enzymes, including β-lactamase, which effectively inhibit the activity of β-lactam antibiotics (Anderl et al., \u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e2000\u003c/span\u003e; Singh et al., \u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e2021\u003c/span\u003e). The second is due to biofilm growth dynamics, whose microorganisms show slower growth rates than planktonic cells (He et al., \u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e2024\u003c/span\u003e). Consequently, if nutrient availability is reduced, bacteria enter a latent or stationary growth phase, making them less vulnerable to nutritional cutback (Portenier et al., \u003cspan citationid=\"CR36\" class=\"CitationRef\"\u003e2006\u003c/span\u003e; Singh et al., \u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e2021\u003c/span\u003e). The third mechanism is related to oxygen metabolism, which is influenced by biofilm surface cells that can deplete available oxygen, resulting in anaerobic niches for the bacterial community (De Beer et al., \u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e1994\u003c/span\u003e). This phenomenon has a differential effect on biofilm cells\u0026rsquo; responses to certain antibiotics, such as aminoglycosides, which are more effective against aerobic bacteria (Tresse et al., \u003cspan citationid=\"CR53\" class=\"CitationRef\"\u003e1995\u003c/span\u003e; Duggan, Sedgley, \u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e2007\u003c/span\u003e). Finally, a persistent subpopulation of microorganisms, despite accounting for a minor fraction of the original population, can evade antimicrobial agents (Spoering, Lewis, \u003cspan citationid=\"CR46\" class=\"CitationRef\"\u003e2001\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eThe human oral cavity contains a large and diverse microbiome, with approximately 700 microbial species present in both forms of biofilm and planktonic cells (Paster et al., \u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e2001\u003c/span\u003e; Valera et al., \u003cspan citationid=\"CR54\" class=\"CitationRef\"\u003e2009\u003c/span\u003e; Zhao et al., \u003cspan citationid=\"CR60\" class=\"CitationRef\"\u003e2017\u003c/span\u003e; Deo, Deshmukh., 2019). These microorganisms have varying penetration capacities into dentinal tubules, typically around 500 \u0026micro;m (Love, Jenkinson., 2002; Parmar et al., \u003cspan citationid=\"CR34\" class=\"CitationRef\"\u003e2011\u003c/span\u003e; Brittan et al., \u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e2016\u003c/span\u003e; Teoh et al., \u003cspan citationid=\"CR51\" class=\"CitationRef\"\u003e2023\u003c/span\u003e). Located within the root canal, a conical-shaped structure with distinct diameters in its thirds, cervical (3.20 \u0026micro;m), middle (2.80 \u0026micro;m), and apical (2.11 \u0026micro;m), the dentinal tubules can suffer bacterial infiltration, which allows microorganisms to reach deeper dentin regions (Camargo et al., \u003cspan citationid=\"CR8\" class=\"CitationRef\"\u003e2007\u003c/span\u003e; Ribeiro et al., \u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e2010\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eIt is worth highlighting that \u003cem\u003eEnterococcus faecalis\u003c/em\u003e has a remarkable penetration ability, reaching depths of 800\u0026ndash;1000 \u0026micro;m after three weeks of incubation (Haapasalo, Orstavik., 1987; Ran et al., \u003cspan citationid=\"CR37\" class=\"CitationRef\"\u003e2014\u003c/span\u003e; Vatkar et al., \u003cspan citationid=\"CR55\" class=\"CitationRef\"\u003e2016\u003c/span\u003e). This bacterium is significant as a nosocomial pathogen capable of causing various infections in humans (Jett, Huycke., 1994; Richards et al., \u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e2000\u003c/span\u003e; Ali et al., \u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e2017\u003c/span\u003e). Despite its small population in the oral microbiota, \u003cem\u003eE. faecalis\u003c/em\u003e is important in the etiology of persistent periapical lesions (Evans et al., \u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e2002\u003c/span\u003e; Sunde et al., \u003cspan citationid=\"CR49\" class=\"CitationRef\"\u003e2002\u003c/span\u003e; Ro\u0026ccedil;as et al., \u003cspan citationid=\"CR40\" class=\"CitationRef\"\u003e2004\u003c/span\u003e; Johnson et al., \u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e2006\u003c/span\u003e; Arias-Moliz et al., \u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e2009\u003c/span\u003e) and persistent apical periodontitis (Siqueira, Ro\u0026ccedil;as., 2022).\u003c/p\u003e \u003cp\u003eBiofilm formation on the dentin walls of root canals is a common occurrence that can be prevented with biomechanical interventions (Stojicic et al., \u003cspan citationid=\"CR47\" class=\"CitationRef\"\u003e2013\u003c/span\u003e; Neelakantan et al., \u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e2017\u003c/span\u003e). However, \u003cem\u003eE. faecalis\u003c/em\u003e is extremely resistant to biomechanical preparation and various intracanal medications, including calcium hydroxide (Vivacqua-Gomes et al., \u003cspan citationid=\"CR58\" class=\"CitationRef\"\u003e2005\u003c/span\u003e; Dunavant et al., \u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e2006\u003c/span\u003e; Abu Hasna et al., \u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e2020\u003c/span\u003e). This microorganism has an exceptional ability to survive in alkaline pH conditions within dentinal tubules for extended periods (Kaufman et al., \u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e2005\u003c/span\u003e; Berber et al., \u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e2006\u003c/span\u003e; Gomes et al., \u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e2006\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eAccordingly, to assess the efficacy of antimicrobial medications, the number of colonies forming units (CFUs) must be determined. To accomplish this, biofilm must be removed from the root canal system while microbial cells are still viable. This procedure usually involves the use of sterilized absorbent paper points, which is the same method used in the \u003cem\u003ein vivo\u003c/em\u003e collection stage. Nonetheless, this approach has limitations, particularly in terms of achieving efficient biofilm dislodgement during the removal step. Thus, in the current study, a novel methodology was proposed to improve the displacement of mature \u003cem\u003eE. faecalis\u003c/em\u003e biofilms from the root canal system, ensuring concomitant bacterial viability while minimizing potential lysis or cell death.\u003c/p\u003e"},{"header":"2 MATERIALS AND METHODS","content":"\u003cdiv id=\"Sec3\" class=\"Section2\"\u003e\n\u003ch2\u003e2.1 Root canal preparation\u003c/h2\u003e\n\u003cp\u003eBovine incisor teeth were preserved in a 0.1% thymol solution at 9\u0026deg;C and washed thoroughly with continuous exposure to running water for 24 h to remove any residual thymol solution. Macroscopic examination and mesiodistal radiography were then used to standardize the samples. Initially, a set of 44 teeth free of cracks, fractures, calcifications, and pronounced curvatures, each with a single canal and cervical diameter of 2\u0026ndash;3 mm, was chosen (Brittan et al., \u003cspan class=\"CitationRef\"\u003e2016\u003c/span\u003e; Camargo et al., \u003cspan class=\"CitationRef\"\u003e2007\u003c/span\u003e; Ribeiro et al., \u003cspan class=\"CitationRef\"\u003e2010\u003c/span\u003e; Valera et al., \u003cspan class=\"CitationRef\"\u003e2009\u003c/span\u003e).\u003c/p\u003e\n\u003cp\u003eThe biomechanical preparation protocol began with perpendicular sectioning of the teeth, 15 mm above the root apex, using a carborundum disk attached to a straight handpiece and a low-speed motor (Beltec, Araraquara, SP, Brazil). The root canal was systematically explored with a K #30 file (Dentsply-Maillefer, Ballaig\u0026uuml;es, Switzerland) until the tip reached the apical foramen, which was then retracted by 1 mm to determine the working length (CT).\u003c/p\u003e\n\u003cp\u003eThe root apexes were sealed with Z100 composite resin (3M, Maplewood, Minnesota, USA) (Fig.\u0026nbsp;\u003cspan class=\"InternalRef\"\u003e1\u003c/span\u003eA), then 37% phosphoric acid etchant (Dentsply, York, Pennsylvania, USA) was applied to a height of 3 mm height above the apex for 20 s. The next steps were to rinse with water and dry the absorbent paper. Next, the Adper Single Bond adhesive system (3M, Maplewood, Minnesota, USA) was applied, followed by a 10 s air dry jet and a 40s light cure. Repeated adhesive system application was followed by an additional 40s light-curing cycle. This strategy aimed to keep adhesive material from entering the root canal via the apical foramen. The biomechanical preparation was done using the ProTaper Universal rotary system (Dentsply-Sirona, Petr\u0026oacute;polis, Rio de Janeiro, Brazil) in the following order: SX, S1, S2, F1, F2, F3, F4, and F5. Nickel\u0026ndash;titanium instruments were integrated into an electric motor contra-angle (VDW Silver, GmbH, Munich, Bavaria, Germany), which introduced the instruments into the canal via an insertion/withdrawal movement with an approximately 3 mm-controlled amplitude and imposed gentle canal wall pressure.\u003c/p\u003e\n\u003cp\u003eThroughout the procedure, an alternated sequence of irrigation-aspiration steps were adopted at each instrument exchange, employing 2 mL of 2.5% NaOCl (BioFlora, Ribeir\u0026atilde;o Preto, S\u0026atilde;o Paulo, Brazil) using a disposable 10 mL plastic syringe (BD, New Jersey, USA) and a Navy tip needle (Ultradent, South Jordan, Utah, USA). An aspiration cannula was also used, along with instrument cleaning and gauze. The final irrigation step involved a 5 min irrigation with 2 mL of 17% EDTA (Da Terra, Ribeir\u0026atilde;o Preto, S\u0026atilde;o Paulo, Brazil), followed by 2 mL of 2.5% NaOCl and 10 mL of distilled water (Kaufman et al., \u003cspan class=\"CitationRef\"\u003e2005\u003c/span\u003e; Berber et al., \u003cspan class=\"CitationRef\"\u003e2006\u003c/span\u003e; Gomes et al., \u003cspan class=\"CitationRef\"\u003e2006\u003c/span\u003e).\u003c/p\u003e\n\u003c/div\u003e\n\u003cdiv id=\"Sec4\" class=\"Section2\"\u003e\n\u003ch2\u003e2.2 Experimental groups\u003c/h2\u003e\n\u003cp\u003eThe specimens were randomly divided into distinct experimental collection groups, each with 12 individual specimens for the quantification of colony-forming units. In each experimental group, a subgroup of two specimens was designated for subsequent scanning electron microscopy (SEM) analysis.\u003c/p\u003e\n\u003cp\u003ePBS, a group consisting of two laterally cleaved specimens, was set aside for SEM analysis. It should be noted that the PBS group was not subjected to the biofilm collection protocol. This subgroup was chosen as a control group for comparison and validation purposes, serving as a positive control reference.\u003c/p\u003e\n\u003cdiv class=\"gridtable\"\u003e\n\u003ctable id=\"Tab1\" border=\"1\"\u003e\u003ccaption\u003e\n\u003cdiv class=\"CaptionNumber\"\u003eTable 1\u003c/div\u003e\n\u003cdiv class=\"CaptionContent\"\u003e\n\u003cp\u003eExperimental groups.\u003c/p\u003e\n\u003c/div\u003e\n\u003c/caption\u003e\n\u003cthead\u003e\n\u003ctr\u003e\n\u003cth align=\"left\"\u003e\n\u003cp\u003eGroups\u003c/p\u003e\n\u003c/th\u003e\n\u003cth align=\"left\"\u003e\n\u003cp\u003eCFU (n)\u003c/p\u003e\n\u003c/th\u003e\n\u003cth align=\"left\"\u003e\n\u003cp\u003eSEM (n)\u003c/p\u003e\n\u003c/th\u003e\n\u003c/tr\u003e\n\u003c/thead\u003e\n\u003ctbody\u003e\n\u003ctr\u003e\n\u003ctd align=\"left\"\u003e\n\u003cp\u003eBiofilm Collection Protocol\u003c/p\u003e\n\u003c/td\u003e\n\u003ctd align=\"char\" char=\".\"\u003e\n\u003cp\u003e12\u003c/p\u003e\n\u003c/td\u003e\n\u003ctd align=\"char\" char=\".\"\u003e\n\u003cp\u003e2\u003c/p\u003e\n\u003c/td\u003e\n\u003c/tr\u003e\n\u003ctr\u003e\n\u003ctd align=\"left\"\u003e\n\u003cp\u003e2nd Biofilm Collection Protocol\u003c/p\u003e\n\u003c/td\u003e\n\u003ctd align=\"char\" char=\".\"\u003e\n\u003cp\u003e12\u003c/p\u003e\n\u003c/td\u003e\n\u003ctd align=\"char\" char=\".\"\u003e\n\u003cp\u003e2\u003c/p\u003e\n\u003c/td\u003e\n\u003c/tr\u003e\n\u003ctr\u003e\n\u003ctd align=\"left\"\u003e\n\u003cp\u003eAbsorbent Paper Points\u003c/p\u003e\n\u003c/td\u003e\n\u003ctd align=\"char\" char=\".\"\u003e\n\u003cp\u003e12\u003c/p\u003e\n\u003c/td\u003e\n\u003ctd align=\"char\" char=\".\"\u003e\n\u003cp\u003e2\u003c/p\u003e\n\u003c/td\u003e\n\u003c/tr\u003e\n\u003ctr\u003e\n\u003ctd align=\"left\"\u003e\n\u003cp\u003ePBS\u003c/p\u003e\n\u003c/td\u003e\n\u003ctd align=\"char\" char=\".\"\u003e\n\u003cp\u003e0\u003c/p\u003e\n\u003c/td\u003e\n\u003ctd align=\"char\" char=\".\"\u003e\n\u003cp\u003e2\u003c/p\u003e\n\u003c/td\u003e\n\u003c/tr\u003e\n\u003c/tbody\u003e\n\u003c/table\u003e\n\u003c/div\u003e\n\u003c/div\u003e\n\u003cdiv id=\"Sec5\" class=\"Section2\"\u003e\n\u003ch2\u003e2.3 Preparation of specimens\u003c/h2\u003e\n\u003cp\u003eA set of 44 1.5 mL Eppendorf tubes was used to premold the roots. Thus, 1 mL of colorless acrylic resin (powder/liquid) (Artigos Odontol\u0026oacute;gicos Cl\u0026aacute;ssico, LTDA, Campo Limpo Paulista, SP, Brazil) was injected into the tubes to immobilize the roots, resulting in the proof bodies (Fig.\u0026nbsp;\u003cspan class=\"InternalRef\"\u003e1\u003c/span\u003eB). The tubes were then sterilized at 121\u0026deg;C for 30 min before being immediately closed to prevent contamination. Before the immobilization step, a 2 mm longitudinal incision was made on both sides of the two roots in each experimental group to allow for subsequent cleavage for SEM analysis (Fig.\u0026nbsp;\u003cspan class=\"InternalRef\"\u003e1\u003c/span\u003eC). Finally, the 44 prepared specimens were loaded onto an Eppendorf tube rack (Fig.\u0026nbsp;\u003cspan class=\"InternalRef\"\u003e1\u003c/span\u003eD) and prepared for new sterilization in an autoclave (Phoenix Luferco, Araraquara, SP, Brazil) at 121\u0026deg;C for 30 min. The sterilization procedure was validated using chemical temperature indicator tape and microbiological tests.\u003c/p\u003e\n\u003c/div\u003e\n\u003cdiv id=\"Sec6\" class=\"Section2\"\u003e\n\u003ch2\u003e2.4 Bacterial species and inoculum preparation\u003c/h2\u003e\n\u003cp\u003eThe strain of \u003cem\u003eE. faecalis\u003c/em\u003e NCTC 775 was chosen for this study due to its biofilm formation and survival in RCS. The strain was cryopreserved at \u0026minus;\u0026thinsp;80\u0026deg;C in cryotubes containing BHI broth and 15% glycerol. To reactivate the strain, it was aerobically grown in 5 mL of BHI at 37\u0026deg;C for 24 h. The strain was then grown in a Petri dish containing M\u0026uuml;eller Hinton Agar (MH) (Sigma Aldrich, San Luis, MO, USA) under a laminar flow hood at 37\u0026deg;C for 24 h.\u003c/p\u003e\n\u003cp\u003eTo prepare the inoculum, bacterial colonies were collected from the MH Petri dish using a sterilized swab and transferred to a test tube with 2.5 mL of sterilized PBS until reaching a bacterial suspension equivalent to 2.0 McFarland scale turbidity (approximately 6.0 \u0026times; 10\u003csup\u003e8\u003c/sup\u003e CFU/mL). Next, 2.5 mL of BHI medium (Sigma Aldrich, San Luis, MO, USA) was added to the suspension, resulting in a final concentration of approximately 3.0 \u0026times; 10\u003csup\u003e8\u003c/sup\u003e CFU/mL, corresponding to 1.0 MacFarland scale. This step was necessary because the color of the culture medium prevented the McFarland scale from being prepared directly in it. To achieve the desired scale, the inoculum\u0026rsquo;s initial concentration was double the appropriate concentration. The initial inoculum was transferred to an equal volume of culture medium, achieving 1.0 on the MacFarland scale.\u003c/p\u003e\n\u003cp\u003eFresh bacterial cultures were prepared daily over 21 days to ensure the viability of the bacterial culture and to maintain the necessary inoculum volume for successful biofilm development (Jett et al., \u003cspan class=\"CitationRef\"\u003e1994\u003c/span\u003e; Ran et al., \u003cspan class=\"CitationRef\"\u003e2014\u003c/span\u003e).\u003c/p\u003e\n\u003c/div\u003e\n\u003cdiv id=\"Sec7\" class=\"Section2\"\u003e\n\u003ch2\u003e2.5 Biofilm formation in the root canal\u003c/h2\u003e\n\u003cp\u003eAmong the 44 bovine incisor tooth roots chosen, one was purposefully left uninoculated to serve as a control for the sterilization process during the trial period, filled with PBS only. The specimens\u0026rsquo; root canals were then inoculated with 10 \u0026micro;L of bacterial suspension (3.0 \u0026times; 10\u003csup\u003e8\u003c/sup\u003e CFU/mL) using aseptic techniques. The canals were carefully filled with micropipettes with sterilized tips to avoid inadvertent inoculum leakage and keep the external root surface clean.\u003c/p\u003e\n\u003cp\u003eFollowing the inoculation step, the prepared roots were incubated for 24 h at 37\u0026deg;C. Following this period, the inoculum was extracted from the root canals. Simultaneously, the canals were gently washed with PBS, which was then aspirated and discarded to effectively remove nonviable bacterial cells.\u003c/p\u003e\n\u003cp\u003eTo ensure optimal biofilm growth, daily inoculation renewal was implemented, with a fresh inoculum introduced into the root canal every 24 h. This procedure allowed for viable bacterial inoculums, which aided in the development of a continuous biofilm throughout the RCS. Throughout the study, all samples were incubated in an aerobic environment at 37\u0026deg;C for 21 days. This time interval allows for the formation of a dense and fully mature biofilm within the root canal, simulating clinical conditions \u003cem\u003ein vivo\u003c/em\u003e and making it suitable for further analysis and evaluation (Stuart et al., \u003cspan class=\"CitationRef\"\u003e2006\u003c/span\u003e; Vivacqua-Gomes et al., \u003cspan class=\"CitationRef\"\u003e2005\u003c/span\u003e).\u003c/p\u003e\n\u003c/div\u003e\n\u003cdiv id=\"Sec8\" class=\"Section2\"\u003e\n\u003ch2\u003e2.6 Biofilm collection protocol\u003c/h2\u003e\n\u003cp\u003eThe biofilm removal procedures (Fig.\u0026nbsp;\u003cspan class=\"InternalRef\"\u003e2\u003c/span\u003e) were performed in a strict sequence of steps to ensure process accuracy. First, the specimens\u0026rsquo; root canals were filled with sterilized PBS. Subsequently, PBS was carefully aspirated from the canals using a micropipette with sterilized tips. Following this step, renewed PBS was introduced into the canals to aid in the removal of any remaining planktonic cells.\u003c/p\u003e\n\u003cp\u003eThe specimens were then transferred to 5 mL Eppendorf tubes with conical bottoms and manipulated using sterilized tweezers. Root canals were carefully placed facing the bottom of the tubes, which were then tightly sealed. The specimens were then vigorously agitated in a vortex (Gehaka, S\u0026atilde;o Paulo, SP, Brazil) at maximum speed for 1 min. This mechanical agitation detaches the biofilm that has formed on the root canal walls.\u003c/p\u003e\n\u003cp\u003eThe next step of the biofilm collection process involved placing 5 mL Eppendorf tubes containing the specimens on a rack and immersing them in an ultrasonic bath (Digital Ultrasonic Cleaner, Kondortech, S\u0026atilde;o Carlos, SP, Brazil) at room temperature (approximately 25\u0026deg; C). This set was immersed in an ultrasonic bath for 480 s to ensure that the biofilm was completely removed from the root canal walls.\u003c/p\u003e\n\u003cp\u003eTo remove the detached biofilm from the interior of the root canal, centrifugation was performed at 700 g for 1 min. Finally, the pellet formed at the bottom of the 5 mL Eppendorf was collected for further counting of CFU. The protocol was standardized to maintain bacterial viability, as demonstrated by Robertson et al., \u003cspan class=\"CitationRef\"\u003e2019\u003c/span\u003e, who demonstrated bacterial viability while using a vortex at maximum power and centrifugation at 7000 g for 7 min.\u003c/p\u003e\n\u003c/div\u003e\n\u003cdiv id=\"Sec9\" class=\"Section2\"\u003e\n\u003ch2\u003e2.7 Second Biofilm Collection\u003c/h2\u003e\n\u003cp\u003eFollowing the collection of RCS biofilms, the same specimens were subjected to a new collection protocol using the same protocol. This strategy was used to determine whether there would be a significant amount of biofilm remaining from the first collection, as well as whether the proposed protocol would require only one or two collections.\u003c/p\u003e\n\u003cp\u003eThe biofilms collected in both protocols were quantified by counting CFU. The difference between the first and second biofilm collections was statistically determined, as described below.\u003c/p\u003e\n\u003c/div\u003e\n\u003cdiv id=\"Sec10\" class=\"Section2\"\u003e\n\u003ch2\u003e2.8 Absorbent paper point collection\u003c/h2\u003e\n\u003cp\u003eFor comparison with the new proposed method, biofilms were collected from the RCS using the absorbent paper point method described by Alfirdous et al. (\u003cspan class=\"CitationRef\"\u003e2022\u003c/span\u003e), with adjustments for the number of paper points. The number of paper tips used for collection increased in proportion to the diameter of the root canal, ranging from one and three units.\u003c/p\u003e\n\u003cp\u003eBefore inserting the paper points into the root canals, each one was saturated with PBS. After the paper points were extracted from the root canals, PBS was drained from them using a pipette with sterilized tips. This procedure eliminated any planktonic bacteria that were present inside the canals.\u003c/p\u003e\n\u003cp\u003eFollowing this preliminary step, the absorbent paper points were resoaked with fresh PBS. Five absorbent paper points with appropriate working diameters (taper 50) were inserted sequentially into the root canals. This strategy ensured that the points were consistently and uniformly placed within the root canal structure.\u003c/p\u003e\n\u003cp\u003eAll biofilm-soaked points were placed in 1.5 mL Eppendorf tubes containing 1 mL of PBS and mixed thoroughly in a vortex mixer for 1 min. These tubes, designed to facilitate subsequent specimen manipulation and analysis, served as storage vessels for subsequent steps, such as serial dilution followed by counting CFUs.\u003c/p\u003e\n\u003c/div\u003e\n\u003cdiv id=\"Sec11\" class=\"Section2\"\u003e\n\u003ch2\u003e2.9 Microbiological analysis\u003c/h2\u003e\n\u003cdiv id=\"Sec12\" class=\"Section3\"\u003e\n\u003ch2\u003e2.9.1 Colony-forming units (CFU)\u003c/h2\u003e\n\u003cp\u003eBiofilms extracted from all experimental groups\u0026rsquo; root canals were serially diluted at a ratio of 1:9. According to this protocol, aliquots of 0.1 mL of each suspension were aseptically inoculated into Petri dishes containing MH Agar medium. This inoculation was performed using a Drigalski spatula and the spread plate methodology (Tortora et al., \u003cspan class=\"CitationRef\"\u003e2018\u003c/span\u003e). The Petri dishes were then incubated under controlled conditions at 37\u0026deg;C for 24 h. After incubation, the colony-forming units (CFU) per square millimeter (CFU/ mm\u0026sup2;) were carried out (Tortora et al., \u003cspan class=\"CitationRef\"\u003e2018\u003c/span\u003e).\u003c/p\u003e\n\u003cp\u003eTo count CFU, the image analysis software ImageJ (Schneider et al., \u003cspan class=\"CitationRef\"\u003e2012\u003c/span\u003e) was employed. To aid analysis, the photograph corresponding to the last dilution was taken. Then, the ImageJ software imported this photograph and converted it to an 8-bit resolution image. Calibration was performed using a 180/250 correlation and a black background was selected. The next step involved isolating a specific region using the circumference option. The analysis focused on particles ranging from 10 to 130 pixels\u0026sup2;. Consequently, the number of colonies was expressed in pixels2.\u003c/p\u003e\n\u003cp\u003eCFU counting can also be done visually with trained and calibrated readers; however, using software greatly reduces the possibility of errors.\u003c/p\u003e\n\u003c/div\u003e\n\u003cdiv id=\"Sec13\" class=\"Section3\"\u003e\n\u003ch2\u003e2.9.2 Scanning electron microscopy analysis of bacterial adhesion\u003c/h2\u003e\n\u003cp\u003eTwo specimens from each experimental group were designated for SEM analysis. Before the SEM analysis, slits were precisely cut into distal and mesial regions of the specimens using a chisel. In the next step, the specimens were cleaved into two halves with a chisel for further analysis.\u003c/p\u003e\n\u003cp\u003eThe prepared specimen halves were immersed in a series of ethanol solutions at increasing concentrations to facilitate dehydration. This set featured ethanol concentrations of 30%, 50%, 70%, 80%, 85%, and 100%. The specimens were immersed in each solution for 1 h, for a total of two hours in absolute ethanol. After dehydration, the specimens were ready for SEM analysis (Huang et al., 2017; Li et al., 2018).\u003c/p\u003e\n\u003cp\u003eThe specimens were appropriately labeled and mounted on cylindrical structures known as stubs. Double-sided carbon adhesive tape was used to hold the specimens in place on the stubs. A fine coating of electrically conductive metal, typically gold or gold\u0026ndash;palladium alloy, was then applied to the specimens using vacuum metallization equipment (SDC 050, Bal-Tec AG, Liechtenstein). This final step was executed to ensure optimal conductivity and structural integrity during the SEM analysis, with specific parameters properly adjusted to ensure precise data acquisition and interpretation. These parameters include 0.01 mbar of pressure, 40 mA of current to facilitate electron emission and interaction with the sample, 50 mm of working distance, and a coverage time of 110 s, which determines the duration of the interaction between the sample and the electron beam. A subsequent metal deposition step with an average thickness of 20\u0026ndash;30 nm ensured a controlled and uniform metal coating.\u003c/p\u003e\n\u003cp\u003eAfter sample preparation, the specimens were transferred to a scanning electron microscope (JSM 5410, JEOL Ltd, Tokyo, Japan) for examination. The specimens\u0026rsquo; apical, middle, and cervical thirds were selected as areas of interest. To capture varying levels of detail, magnifications of 500, 2000, and 5000x were used. The acquired images were analyzed with a focus on two distinct aspects. First, the assessment confirmed the extension of bacterial adhesion to dentinal tubules, which provided information about interactions at the biofilm\u0026ndash;tubule interface. Second, the study assessed the effectiveness of biofilm removal techniques in the root canal.\u003c/p\u003e\n\u003c/div\u003e\n\u003c/div\u003e\n\u003cdiv id=\"Sec14\" class=\"Section2\"\u003e\n\u003ch2\u003e2.10 Statistical Analysis\u003c/h2\u003e\n\u003cp\u003eFor statistical analysis, the Shapiro\u0026ndash;Wilk test was used to verify data distribution. It was discovered that measurements were not normally distributed and thus were transformed into log\u003csub\u003e10\u003c/sub\u003e values before the one-way parametric method was used to compare the groups. The significance level was set to 5%.\u003c/p\u003e\n\u003c/div\u003e"},{"header":"3 RESULTS","content":"\u003cdiv id=\"Sec16\" class=\"Section2\"\u003e\n\u003ch2\u003e3.1 Evaluation of Collection Efficiency Considering CFU Counting\u003c/h2\u003e\n\u003cp\u003eThe evaluation of collection efficiency produced comparable results in terms of CFU counting for all experimental groups (Table\u0026nbsp;\u003cspan class=\"InternalRef\"\u003e2\u003c/span\u003e). Surprisingly, the collection conducted under the group biofilm collection protocol showed higher CFU values. Conversely, the absorbent paper point collection and 2\u0026deg; biofilm collection protocols produced significantly lower CFU values (Fig.\u0026nbsp;\u003cspan class=\"InternalRef\"\u003e3\u003c/span\u003e).\u003c/p\u003e\n\u003cp\u003eIt is worth noting that the biofilm collection protocol effectively removed the biofilm from the root canal, with only a few bacteria found inside the dentinal tubules. Therefore, we believe that the second collection is unnecessary to ensure the experiments\u0026rsquo; effectiveness.\u003c/p\u003e\n\u003cdiv class=\"gridtable\"\u003e\n\u003ctable id=\"Tab2\" border=\"1\"\u003e\u003ccaption\u003e\n\u003cdiv class=\"CaptionNumber\"\u003eTable 2\u003c/div\u003e\n\u003cdiv class=\"CaptionContent\"\u003e\n\u003cp\u003eNumber of colonies forming units (CFU) in different biofilm collection protocols. Distinct letters in the same column indicate statistically significant differences between the groups.\u003c/p\u003e\n\u003c/div\u003e\n\u003c/caption\u003e\n\u003cthead\u003e\n\u003ctr\u003e\n\u003cth align=\"left\"\u003e\n\u003cp\u003eExperimental groups\u003c/p\u003e\n\u003c/th\u003e\n\u003cth align=\"left\"\u003e\n\u003cp\u003eThe number of colonies forming units (CFU) log\u003csub\u003e10\u003c/sub\u003e\u003c/p\u003e\n\u003c/th\u003e\n\u003c/tr\u003e\n\u003c/thead\u003e\n\u003ctbody\u003e\n\u003ctr\u003e\n\u003ctd align=\"left\"\u003e\n\u003cp\u003eBiofilm Collection Protocol\u003c/p\u003e\n\u003c/td\u003e\n\u003ctd align=\"left\"\u003e\n\u003cp\u003e7.38\u0026thinsp;\u0026plusmn;\u0026thinsp;6.40\u003csup\u003eA\u003c/sup\u003e\u003c/p\u003e\n\u003c/td\u003e\n\u003c/tr\u003e\n\u003ctr\u003e\n\u003ctd align=\"left\"\u003e\n\u003cp\u003e2nd Biofilm Collection Protocol\u003c/p\u003e\n\u003c/td\u003e\n\u003ctd align=\"left\"\u003e\n\u003cp\u003e6.20\u0026thinsp;\u0026plusmn;\u0026thinsp;5.67\u003csup\u003eB\u003c/sup\u003e\u003c/p\u003e\n\u003c/td\u003e\n\u003c/tr\u003e\n\u003ctr\u003e\n\u003ctd align=\"left\"\u003e\n\u003cp\u003eAbsorbent Paper Point Collection\u003c/p\u003e\n\u003c/td\u003e\n\u003ctd align=\"left\"\u003e\n\u003cp\u003e6.05\u0026thinsp;\u0026plusmn;\u0026thinsp;5.68\u003csup\u003eB\u003c/sup\u003e\u003c/p\u003e\n\u003c/td\u003e\n\u003c/tr\u003e\n\u003c/tbody\u003e\n\u003c/table\u003e\n\u003c/div\u003e\n\u003c/div\u003e\n\u003cdiv id=\"Sec17\" class=\"Section2\"\u003e\n\u003ch2\u003e3.2 Analysis of Collected Biofilms by Scanning Electron Microtomography (SEM)\u003c/h2\u003e\n\u003cp\u003eQualitative evaluation of photomicrographs obtained through SEM analysis revealed significant differences among all experimental groups, each distinguished by the specific collection techniques used. The PBS control and absorbent paper point collection groups revealed dense biofilm at the periphery of the dentinal tubules. In stark contrast, the proposed biofilm collection protocol effectively removed bacterial biofilm content from within the dentinal tubules. Notably, the absorbent paper point group\u0026rsquo;s collection showed a significant accumulation of biofilm adhering across all assessed regions, which was similar to the results observed in the control group (Fig.\u0026nbsp;\u003cspan class=\"InternalRef\"\u003e4\u003c/span\u003e).\u003c/p\u003e\n\u003c/div\u003e"},{"header":"4 DISCUSSION","content":"\u003cp\u003e \u003cdiv class=\"BlockQuote\"\u003e \u003cp\u003eBiofilms are closely associated with RCS infections, and their complexity, combined with the nature of biofilms, makes disinfection of this system extremely difficult. Much of the research in the field of endodontics has focused on the characterization of microbial biofilms formed in RCS, as well as clinical methods for killing and eliminating them. Biofilms must be studied in endodontics microbiology to understand the pathogenic potential of the microorganisms involved in the infection and to evaluate new disinfection strategies for RCS (Jhajharia et al., \u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e2015\u003c/span\u003e; Neelakantan et al., \u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e2017\u003c/span\u003e). Recent research has shown that biofilm development is complex, requiring a delicate balance of bacterial proliferation and the production of extracellular polymeric substances (EPS). Therefore, the treatment of biofilm infections is challenging and requires strategies tailored to the stage of biofilm development (He et al., \u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e2024\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eTraditionally, bacterial biofilms are removed from root canals using sterilized absorbent paper points, allowing bacteria absorption. The proposed method aims to simulate the \u003cem\u003ein vivo\u003c/em\u003e collection procedure (Alfirdous et al., \u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e2022\u003c/span\u003e). It is important to note \u003cem\u003ethat in vivo\u003c/em\u003e investigations gather a distinct set of RCS decontamination protocols involving physical process instrumentation (Alfirdous et al., \u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e2022\u003c/span\u003e) and chemical methods using irrigating solutions with or without activation (Căpută et al., \u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e2019\u003c/span\u003e; Susila, Minu, \u003cspan citationid=\"CR50\" class=\"CitationRef\"\u003e2019\u003c/span\u003e). Nonetheless, certain areas of the RCS present challenges to effective paper point absorption. Therefore, to bring the experimental approach closer to the clinical reality of endodontic infections, a methodology for optimal collection of microbial biofilms in RCS was proposed.\u003c/p\u003e \u003cp\u003eTo propose the biofilm collection protocol in the RCS, several parameters were evaluated and consulted in the literature, including the preservation of cell viability when the biofilms were subjected to various experimental steps. The cellular viability of the collected biofilms was assessed individually in three stages: after vortexing and ultrasonic bath protocols, and then after collection centrifugation. Finally, the cell viability of the biofilms was assessed following their sequential exposure to the three processes. The study found bacterial viability in all phases tested (data not shown). Robertson et al. (\u003cspan citationid=\"CR62\" class=\"CitationRef\"\u003e2019\u003c/span\u003e) reported similar results, using vortexing and centrifugation at 7,000 g for 7 min to maintain bacterial viability. The authors used the Live/Dead kit to determine the number of dead and living cells. Chen et al. (\u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e2021\u003c/span\u003e) achieved similar results by centrifugation at 5,000 g for 15 min to form pellets for analysis while preserving bacterial viability.\u003c/p\u003e \u003cp\u003eThe results from the collection using absorbent paper points and the second collection after the initial collection step were statistically similar. On the other hand, the initial biofilm collection protocol had significantly higher CFU values than the other two protocols. The CFU values discovered in the second collection are statistically insignificant for analysis, indicating that the second collection is irrelevant in terms of CFU values when considering the cost, time, and labor required for its execution. These findings highlight the effectiveness of the proposed collection method when compared to collection using absorbent paper points.\u003c/p\u003e \u003cp\u003eThe traditional method of collecting biofilm using sterilized absorbent paper points has limitations in accessing the entire root canal structure, resulting in flaws in the bacterial sampling step (Swimberghe, Coenye, et al., 2019; Hoedke et al., \u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e2021\u003c/span\u003e; Alfirdous et al., \u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e2022\u003c/span\u003e). The inherent limitation of this method is its tendency to predominantly capture planktonic bacteria while failing to effectively retrieve biofilm-associated bacteria (Neelakantan et al., \u003cspan citationid=\"CR29\" class=\"CitationRef\"\u003e2015\u003c/span\u003e; Neuhaus et al., \u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e2016\u003c/span\u003e). To address this limitation, a new collection protocol that includes vortex mixing, ultrasonic bath immersion, and centrifugation is proposed. The primary goal of this research is to develop a collection methodology that not only ensures the effective removal of biofilm material from the radicular canal but also protects the viability and integrity of bacterial cells by preventing lysis and death. In this regard, it is a pioneering study because the proposal addresses both the efficiency of material removal from the radicular canal using a three-step process and longterm bacterial viability.\u003c/p\u003e \u003cp\u003eTo demonstrate the efficacy and reliability of the proposed collection protocol, a comparison with a consolidated method was conducted. It is also important to note that the variability and often subjective nature of operator skills can have a significant impact on the outcomes of various collection methods. Thereby, the operator\u0026rsquo;s skills make an expressive contribution to potential discrepancies in results. To achieve reliable and unbiased results, the ideal collection method must be highly reproducible, separating the obtained results from operator-dependent factors. Conversely, techniques that use instruments to mechanically dislodge biofilms run the risk of lysing bacterial cells, compromising the accuracy of the results. In contrast, the centrifuge-based collection protocol used in this study proved advantageous because it avoids bacterial lysis (Chen et al., \u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e2021\u003c/span\u003e). The methodology used allows for the preservation of cell viability during the detachment of biofilm material from the root walls, as demonstrated in the findings of this study.\u003c/p\u003e \u003cp\u003eIt is important to note that there are several classic and contemporary techniques for removing biofilms from the RCS to eliminate pathogenic microorganisms and, as a result, disinfecting the canal in the practice of endodontic therapy (Josic et al., \u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e2022\u003c/span\u003e). However, the goal of this article is not to evaluate RCS disinfection; rather, it is to provide researchers with an experimental model so that such disinfection strategies can be evaluated \u003cem\u003ein vivo\u003c/em\u003e with greater representativeness of biofilm collection, reproducibility, and without interfering with biofilm viability, thereby avoiding undesirable results.\u003c/p\u003e \u003cp\u003eComparative analysis of SEM images provided valuable qualitative information about the efficacy of the various collection methods tested. There is a similarity in terms of CFU when comparing the absorbent paper point group to the second collect protocol group, which collects proc planktonic bacteria preferentially. Qualitative SEM analyses revealed the presence of bacterial cells in the dentinal tubules following the paper tip collection process, whereas the proposed protocol revealed a significant lack of bacteria in these regions. These findings are significant because many protocols for disinfecting endodontic canals have varying effectiveness in penetrating dental tubules (Josic et al., \u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e2022\u003c/span\u003e). Therefore, the effective \u003cem\u003ein vivo\u003c/em\u003e evaluation of these protocols via CFU counting is dependent on the efficient collection of biofilms in these deeper regions.\u003c/p\u003e \u003c/div\u003e \u003c/p\u003e"},{"header":"5 CONCLUSION","content":"\u003cp\u003eIn conclusion, this study found that a new protocol for collecting biofilm from RCS \u003cem\u003ein vivo\u003c/em\u003e was effective. The current study supports the efficacy of the devised collection protocol over the conventional paper point method, establishing it as the best approach for thoroughly retrieving bacterial biofilm from within the RCS. The protocol\u0026rsquo;s robust performance is highlighted by its inherent reproducibility and avoidance of operator-dependent variations. Visual analysis with scanning electron microscopy provides compelling evidence, revealing almost no bacterial presence following collection. It should be noted that the proposed protocol has the advantage of being able to collect biofilm from deeper areas of RCS, which allows it to be used in different experimental protocols. However, in this study, teeth with anatomical variations in the RCS were not evaluated during biofilm collection, which may limit future studies of teeth with such characteristics.\u003c/p\u003e"},{"header":"DECLARATIONS","content":"\u003cp\u003e\u003cstrong\u003eAuthor Contributions\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eRafael da Silva Goulart: methodology (lead); conceptualization (lead); writing\u0026ndash;original draft (lead); formal analysis (lead); writing\u0026ndash;review and editing (equal). Mariana Oliveira-Silva: methodology (supporting) and review and editing (supporting). Yara Teresinha Correa Silva Sousa: conceptualization (supporting)and review and editing (equal). Carlos Eduardo Saraiva Miranda: conceptualization (lead); writing\u0026ndash;original draft (lead); writing\u0026ndash;review and editing (equal). Andr\u0026eacute; Pitondo-Silva: conceptualization (lead); writing\u0026ndash;original draft (lead); writing\u0026ndash;review and editing (equal).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eData Availability\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eNo data were used to support this study.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eConflicts of Interest\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors declare no conflicts of interest.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eEthical approval\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eEthical approval for the use of bovine teeth in this study was obtained from the Animal Use Ethics Committee of the University of Ribeir\u0026atilde;o Preto (Ribeir\u0026atilde;o Preto, SP, Brazil) [approval number 01/2017]. There was no patient involvement in the study.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eFunding\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eRafael da Silva Goulart is a Ph.D. student fellow of Coordena\u0026ccedil;\u0026atilde;o de Aperfei\u0026ccedil;oamento de Pessoal de N\u0026iacute;vel Superior (CAPES); therefore, this study was financed in part by the CAPES (grant no 88887.493929/2020-00).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAcknowledgments\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThis work was reviewed by a native English speaker with expertise in Endodontics from the ENAGO (https://www.enago.com.br/).\u003c/p\u003e"},{"header":"REFERENCES","content":"\u003col\u003e\n\u003cli\u003eAbu Hasna, A., Khoury, R. D., Toia, C. C., Gon\u0026ccedil;alves, G. B., de Andrade, F. B., Talge Carvalho, C. A., Ribeiro Camargo, C. H., \u0026amp; Carneiro Valera, M. (2020). \u003cem\u003eIn vitro\u003c/em\u003e Evaluation of the Antimicrobial Effect of N-acetylcysteine and Photodynamic Therapy on Root Canals Infected with \u003cem\u003eEnterococcus faecalis.\u003c/em\u003e Iranian endodontic journal, 15(4), 236\u0026ndash;245.\u003c/li\u003e\n\u003cli\u003eAlfirdous, R. A., Alquiria, T. A., Jacinto, R. C., \u0026amp; Martinho, F. C. (2022). A modified dentine infection model with fluorescent lipopolysaccharide and lipopolysaccharides sampling technique to compare XP-Endo finisher and passive ultrasonic irrigation. International endodontic journal, 55(10), 1081\u0026ndash;1090. \u003c/li\u003e\n\u003cli\u003eAli, L., Goraya, M. U., Arafat, Y., Ajmal, M., Chen, J. L., \u0026amp; Yu, D. (2017). Molecular Mechanism of Quorum-Sensing in \u003cem\u003eEnterococcus faecalis\u003c/em\u003e: Its Role in Virulence and Therapeutic Approaches. International journal of molecular sciences, 18(5), 960. \u003c/li\u003e\n\u003cli\u003eAnderl, J. N., Franklin, M. J., \u0026amp; Stewart, P. S. (2000). Role of Antibiotic Penetration Limitation in Klebsiella pneumoniae Biofilm Resistance to Ampicillin and Ciprofloxacin. Antimicrobial Agents and Chemotherapy, 44(7), 1818\u0026ndash;1824.\u003c/li\u003e\n\u003cli\u003eArias-Moliz, M. T., Ferrer-Luque, C. M., Espigares-Garc\u0026iacute;a, M., \u0026amp; Baca, P. (2009). \u003cem\u003eEnterococcus faecalis\u003c/em\u003e biofilms eradication by root canal irrigants. Journal of endodontics, 35(5), 711\u0026ndash;714.\u003c/li\u003e\n\u003cli\u003eBerber, V. B., Gomes, B. P., Sena, N. T., Vianna, M. E., Ferraz, C. C., Zaia, A. A., \u0026amp; Souza-Filho, F. J. (2006). Efficacy of various concentrations of NaOCl and instrumentation techniques in reducing \u003cem\u003eEnterococcus faecalis\u003c/em\u003e within root canals and dentinal tubules. International endodontic journal, 39(1), 10\u0026ndash;17.\u003c/li\u003e\n\u003cli\u003eBrittan, J. L., Sprague, S. V., Macdonald, E. L., Love, R. M., Jenkinson, H. F., \u0026amp; West, N. X. (2016). \u003cem\u003eIn vivo\u003c/em\u003e model for microbial invasion of tooth root dentinal tubules. \u003cem\u003eJournal of applied oral science : revista FOB\u003c/em\u003e, \u003cem\u003e24\u003c/em\u003e(2), 126\u0026ndash;135. \u003c/li\u003e\n\u003cli\u003eCamargo, C. H. R., Siviero, M., Camargo, S. E. A., de Oliveira, S. H. G., Carvalho, C. A. T., \u0026amp; Valera, M. C. (2007). Topographical, Diametral, and Quantitative Analysis of Dentin Tubules in the Root Canals of Human and Bovine Teeth. Journal of Endodontics, 33(4), 422\u0026ndash;426. \u003c/li\u003e\n\u003cli\u003eCăpută, P. E., Retsas, A., Kuijk, L., Ch\u0026aacute;vez de Paz, L. E., \u0026amp; Boutsioukis, C. (2019). Ultrasonic Irrigant Activation during Root Canal Treatment: A Systematic Review. Journal of endodontics, 45(1), 31\u0026ndash;44. \u003c/li\u003e\n\u003cli\u003eChen, J., Tomasek, M., Nu\u0026ntilde;ez, E., \u0026amp; Gau, V. (2021). 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Comparative evaluation of endodontic irrigants against \u003cem\u003eEnterococcus faecalis\u003c/em\u003e biofilms. Journal of endodontics, 32(6), 527\u0026ndash;531. \u003c/li\u003e\n\u003cli\u003eEvans, M., Davies, J. K., Sundqvist, G., \u0026amp; Figdor, D. (2002). Mechanisms involved in the resistance of \u003cem\u003eEnterococcus faecalis\u003c/em\u003e to calcium hydroxide. International endodontic journal, 35(3), 221\u0026ndash;228. \u003c/li\u003e\n\u003cli\u003eGomes, B. P., Pinheiro, E. T., Sousa, E. L., Jacinto, R. C., Zaia, A. A., Ferraz, C. C., \u0026amp; de Souza-Filho, F. J. (2006). \u003cem\u003eEnterococcus faecalis\u003c/em\u003e in dental root canals detected by culture and by polymerase chain reaction analysis. Oral surgery, oral medicine, oral pathology, oral radiology, and endodontics, 102(2), 247\u0026ndash;253. \u003c/li\u003e\n\u003cli\u003eHaapasalo, M., \u0026amp; \u0026Oslash;rstavik, D. (1987). \u003cem\u003eIn vitro\u003c/em\u003e Infection and of Dentinal Tubules. Journal of Dental Research, 66(8), 1375\u0026ndash;1379.\u003c/li\u003e\n\u003cli\u003eHaapasalo, M., Shen, Y., Wang, Z., \u0026amp; Gao, Y. (2014). Irrigation in endodontics. British dental journal, 216(6), 299\u0026ndash;303.\u003c/li\u003e\n\u003cli\u003eHe, W., Liu, H., Wang, Z., Tay, F. R., \u0026amp; Shen, Y. (2024). The Dynamics of Bacterial Proliferation, Viability, and Extracellular Polymeric Substances in Oral Biofilm Development. Journal of Dentistry, 104882.\u003c/li\u003e\n\u003cli\u003eHoedke, D., Kaulika, N., Dommisch, H., Schlafer, S., Shemesh, H., \u0026amp; Bitter, K. (2021). Reduction of dual-species biofilm after sonic- or ultrasonic-activated irrigation protocols: A laboratory study. International endodontic journal, 54(12), 2219\u0026ndash;2228.\u003c/li\u003e\n\u003cli\u003eJhajharia, K., Parolia, A., Shetty, K. V., \u0026amp; Mehta, L. K. (2015). Biofilm in endodontics: a review. Journal of International Society of Preventive \u0026amp; Community Dentistry, 5(1), 1.\u003c/li\u003e\n\u003cli\u003eJett, B. D., Huycke, M. M., \u0026amp; Gilmore, M. S. (1994). Virulence of enterococci. Clinical microbiology reviews, 7(4), 462\u0026ndash;478.\u003c/li\u003e\n\u003cli\u003eJohnson, E. M., Flannagan, S. E., \u0026amp; Sedgley, C. M. (2006). Coaggregation interactions between oral and endodontic \u003cem\u003eEnterococcus faecalis\u003c/em\u003e and bacterial species isolated from persistent apical periodontitis. Journal of endodontics, 32(10), 946\u0026ndash;950.\u003c/li\u003e\n\u003cli\u003eJosic, U., Mazzitelli, C., Maravic, T., Fidler, A., Breschi, L., \u0026amp; Mazzoni, A. (2022). Biofilm in endodontics: In vitro cultivation possibilities, sonic-, ultrasonic-and laser-assisted removal techniques and evaluation of the cleaning efficacy. Polymers, 14(7), 1334.\u003c/li\u003e\n\u003cli\u003eKaufman, B., Sp\u0026aring;ngberg, L., Barry, J., \u0026amp; Fouad, A. F. (2005). Enterococcus spp. in endodontically treated teeth with and without periradicular lesions. Journal of endodontics, 31(12), 851\u0026ndash;856. \u003c/li\u003e\n\u003cli\u003eLeoni, G. B., Versiani, M. A., P\u0026eacute;cora, J. D., \u0026amp; Dami\u0026atilde;o de Sousa-Neto, M. (2014). Micro\u0026ndash;Computed Tomographic Analysis of the Root Canal Morphology of Mandibular Incisors. Journal of Endodontics, 40(5), 710\u0026ndash;716.\u003c/li\u003e\n\u003cli\u003eLove, R. M., \u0026amp; Jenkinson, H. F. (2002). Invasion of dentinal tubules by oral bacteria. Critical reviews in oral biology and medicine: an official publication of the American Association of Oral Biologists, 13(2), 171\u0026ndash;183. \u003c/li\u003e\n\u003cli\u003eNair, P. N. R., Henry, S., Cano, V., \u0026amp; Vera, J. (2005). Microbial status of apical root canal system of human mandibular first molars with primary apical periodontitis after \u0026ldquo;one-visit\u0026rdquo; endodontic treatment. 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Microbiological Aspects of Root Canal Infections and Disinfection Strategies: An Update Review on the Current Knowledge and Challenges. Front Oral Health.;2:672887.\u003c/li\u003e\n\u003cli\u003eZhao, H., Chu, M., Huang, Z., Yang, X., Ran, S., Hu, B., Zhang, C., \u0026amp; Liang, J. (2017). Variations in oral microbiota associated with oral cancer. Scientific reports, 7(1), 11773. \u003c/li\u003e\n\u003cli\u003eChen J, Tomasek M, Nu\u0026ntilde;ez E, Gau V. Method for concentrating viable microorganisms for microbial load determination and eliminating uncertainty from matrix effects from urine and whole blood. MethodsX. 2021;8:101451.\u003c/li\u003e\n\u003cli\u003eRobertson, J., McGoverin, C., Vanholsbeeck, F., Swift, S.. Optimisation of the protocol for the LIVE/DEAD\u0026reg; BacLightTM bacterial viability kit for rapid determination of bacterial load. Frontiers in microbiology. 10, 801. 2019\u003c/li\u003e\n\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[{"identity":"2d106320-8cef-4721-b25d-5f4baf08fe35","identifier":"10.13039/501100002322","name":"Coordenação de Aperfeiçoamento de Pessoal de Nível Superior","awardNumber":"88887.493929/2020-00","order_by":0}],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":true,"hideJournal":true,"highlight":"","institution":"Universidade de Ribeirão Preto","isAcceptedByJournal":false,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"researchsquare","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":true,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"/submission","title":"Research Square","twitterHandle":"researchsquare","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"","reportingPortfolio":"","inReviewEnabled":false,"inReviewRevisionsEnabled":true},"keywords":"bacterial biofilms, root canal system, endodontic infection, colony-forming units, biofilm collection","lastPublishedDoi":"10.21203/rs.3.rs-4006763/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-4006763/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003ch2\u003eAim\u003c/h2\u003e \u003cp\u003eThe goal of this study was to standardize a new protocol for collecting biofilm from the interior of the root canal system (RCS) for \u003cem\u003ein vivo\u003c/em\u003e testing.\u003c/p\u003e\u003ch2\u003eMethodology:\u003c/h2\u003e \u003cp\u003eIn this study, 44 bovine incisors were used. The samples were divided into three experimental groups: 14 teeth, 12 for counting colony-forming units (CFU), and two samples for scanning electron microscopy (SEM). The first group was used for the biofilm collection protocol proposed here, the second group for the 2nd Biofilm Collection Protocol collection, and the third group for biofilm collection with an absorbent paper tip. Two additional teeth were used as sterilization controls to ensure that the experiments were free of contamination. The coronal region was removed and standardized at 15 mm. They were fitted with a Protaper up to the F5 insert, and the apical foramen was sealed with composite resin. The roots were stabilized with acrylic resin in a 1.5 mL Eppendorf tube. The specimens were sterilized and then inoculated with \u003cem\u003eEnterococcus faecalis\u003c/em\u003e NTCT 775 every 24 h for 21 days. Following the period, each group underwent biofilm collection protocols, and CFU and scanning electron microscopy (SEM) data were analyzed. Shapiro\u0026ndash;Wilk and one-way ANOVA tests were used to determine statistically significant differences between groups.\u003c/p\u003e\u003ch2\u003eResults\u003c/h2\u003e \u003cp\u003eThe biofilm collection protocol group had the most CFUs, with extremely high values when compared with the other groups when converted to Log10. The results of the One-Way ANOVA test revealed that the 2nd collection protocol and absorbent paper tip collection groups were statistically similar (p\u0026thinsp;\u0026gt;\u0026thinsp;0.05), whereas the biofilm collection protocol group was not.\u003c/p\u003e\u003ch2\u003eConclusion\u003c/h2\u003e \u003cp\u003eThe biofilm collection protocol proposed in this study was effective at collecting microorganisms from within the RCS. Compared to the biofilm collection protocol with paper cones, the \u003cem\u003ein vivo\u003c/em\u003e collection protocol from bovine teeth yielded significantly more CFUs. Thus, the proposed protocol significantly increases the bacterial load of biofilms collected from the RCS sample, bringing the experiments closer to the reality of endodontic infections.\u003c/p\u003e","manuscriptTitle":"Optimized protocol for collecting root canal biofilms for in vitro studies","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2024-03-04 08:25:01","doi":"10.21203/rs.3.rs-4006763/v1","editorialEvents":[{"type":"communityComments","content":0}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"researchsquare","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":true,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"/submission","title":"Research Square","twitterHandle":"researchsquare","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"","reportingPortfolio":"","inReviewEnabled":false,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"db0a8d39-1bab-460e-9e33-57fec77c4e86","owner":[],"postedDate":"March 4th, 2024","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"posted","subjectAreas":[{"id":29094312,"name":"Dentistry"},{"id":29094313,"name":"General Microbiology"}],"tags":[],"updatedAt":"2024-03-04T08:25:02+00:00","versionOfRecord":[],"versionCreatedAt":"2024-03-04 08:25:01","video":"","vorDoi":"","vorDoiUrl":"","workflowStages":[]},"version":"v1","identity":"rs-4006763","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-4006763","identity":"rs-4006763","version":["v1"]},"buildId":"8U1c8b4HqxoKbykW_rLl7","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}