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Mazzitelli, C.; , .; Maravic, T.; Mazzoni, A.; Breschi, L.; Fidler, A. Biofilm in Endodontics. Encyclopedia. Available online: (accessed on 22 February 2024).
Mazzitelli C,  , Maravic T, Mazzoni A, Breschi L, Fidler A. Biofilm in Endodontics. Encyclopedia. Available at: Accessed February 22, 2024.
Mazzitelli, Claudia, , Tatjana Maravic, Annalisa Mazzoni, Lorenzo Breschi, Aleš Fidler. "Biofilm in Endodontics" Encyclopedia, (accessed February 22, 2024).
Mazzitelli, C., , ., Maravic, T., Mazzoni, A., Breschi, L., & Fidler, A. (2022, April 06). Biofilm in Endodontics. In Encyclopedia.
Mazzitelli, Claudia, et al. "Biofilm in Endodontics." Encyclopedia. Web. 06 April, 2022.
Biofilm in Endodontics

Sonic, ultrasonic and Er:Yag laser agitation, in general, offer better biofilm removal when compared to conventional irrigation methods delivered by syringe and needle. The choice of the right irrigation solution is an important factor for removal of the endodontic biofilm, with water and saline being less effective compared to NaOCl and CHX. However, due to heterogeneity in methodologies, it is difficult to compare adjuvant endodontic techniques with one another and give recommendations for the most efficient method in biofilm removal.

biofilm endodontics irrigation sonic ultrasonic laser

1. Biofilm Cultivation

1.1. Monospecies Biofilm

The endodontic bacteria are usually organized in biofilm communities which are present not only in the main canal, but in the overall root canal system [1]. The extracellular matrix of the biofilm offers bacteria higher survival rates in challenging growth and environmental conditions [2]. In order to mimic conditions that are well established within infected root canals, authors put their efforts into growing E. faecalis biofilms since it is considered to be the leading pathogen associated with failed endodontic treatment. Among the microorganisms commonly isolated in the endodontic space, this microorganism represents the leading pathogen largely associated with failed endodontic treatment [3].
According to the reviewed articles, potential double origins, lab-adapted strains [4][5][6][7][8][9][10][11][12][13][14][15][16][17][18][19][20][21][22][23][24][25][26][27][28] and clinically isolated E. faecalis were noted [29]. E. faecalis strain isolated from root canal of pulpless teeth is available, but only two authors reported using it [10][12], while the majority of the studies used strains of E. faecalis isolated from different tissues and fluids.
When choosing the strain of E. faecalis, dental researchers should be aware of its genetic heterogeneity which is observed inside a single population, as well as that different strains of E. faecalis can be detected in the oral cavity of one individual [30][31].

1.2. Multispecies Biofilm

Mixed endodontic infections are more common than infections caused by a single microorganism [2]. Collecting subgingival plaque from dental patients is a method used to grow multispecies biofilm [32][33]. Another approach for multispecies biofilm sampling is an intraoral contamination process by wearing a custom-made appliance for 6–8 days. However, this method is very subjective as the patients voluntarily carry the appliance and follow the diet recommendations [33]. A simplified, lab-adapted, dual-species biofilm model of E. faecalis and Streptococcus mutans was introduced, as well as a three-species biofilm composed of E. faecalis, Streptococcus mitis and Campylobacter rectus [25][34]. Only one study investigated the removal of dual-species biofilm composed of vancomycin resistant E. faecalis and Candida albicans [17]. Niazi et al. (2014) used a biofilm consisting of five different species of microorganisms [35].

1.3. Biofilm Mimicking

Few studies included in this entry used non-bacterial approaches to test different methods of biofilm removal. Macedo et al. (2014) proposed a hydrogel model to provide visualization of biofilm removal by ultrasonic techniques. As it stated, viscoelastic properties of hydrogel can be compared to the one of bacterial biofilm and therefore it may be suitable for replacing bacterial biofilm in in vitro studies [36]. Joy et al. (2015) applied layers of stained collagen to the dentin surface and analyzed digital images of its removal by ultrasonic irrigation [37].

1.4. Substrate and Period of Incubation

The majority of research included in this entry uses human [4][5][6][7][9][37][10][29][38][12][13][14][15][16][17][19][20][22][23][24][25][26][27][28][33][39][40] and animal-bovine dentin [8][41] as substrate for biofilm growth and formation.
A general pattern in preparing samples for bacterial inoculation was observed among the studies which included both human and bovine dentin: after examining the extracted teeth, root canals were enlarged and shaped using endo files, with sodium hypochlorite (NaOCl) serving as an irrigant and EDTA used for smear layer removal. In contrast, Meire’s et al. (2012) presented a different approach since the crowns were firstly cut and dentin slices of standardized thickness were obtained for further testing [20]. Similarly, Bao et al. (2017) used a split tooth model which, after biofilm removal efforts have been made, allows dissembling the tooth and gaining a clear insight into the dentin surface [39]. Another methodology observed in the reviewed studies focuses on use of bovine dentin sections that serve as a substrate for multispecies biofilm cultivation. These sections were incorporated within an orthodontic device and worn by a volunteer allowing oral bacteria to accumulate on the dentin surfaces [41].
Hydroxyapatite (HA) discs are frequently used in dental research and possess the affinity towards bacteria colonization [42][43]. Consequently, both Noiri et al. (2008) and Shen et al. (2010) used hydroxyapatite discs for E. faecalis biofilm cultivation [32][11].
Further, six studies included in this research used root canal models as substrate for biofilm formation. In the most recent studies, root canal models were created using CAD technology and 3D printing. The goal of 3D printing is to create a desirable, transparent and anatomically standardized model which would allow an insight into real-time interaction between irrigants and biofilm removal [21][44].
Time plays an important role in biofilm formation, allowing bacteria to aggregate and form a network of polymer strands. Scanning electron microscope (SEM) investigations revealed that after 1 week of incubation, a biofilm-like structure can be observed on dentin surface. After 2, 3 and 4 weeks, biofilm becomes thicker and thus more challenging to remove. Mature biofilm with characteristic honey-comb like structures can be observed after 6 weeks of incubation [45].
In studies reviewed in this entry, the researchers determined the incubation period based on data available from the literature and their personal preference. However, as pointed out earlier, various incubation periods result in different maturity and thickness of the biofilm, which eventually can lead to unequal effort towards biofilm removal.

2. Biofilm Removal Techniques

2.1. Sonic Devices

Sodium hypochlorite (NaOCl) is one of the most commonly used irrigants in endodontic practice. Authors are in agreement that sonic energizing with different concentrations of NaOCl offers greater biofilm disruption than sonic energizing with water or saline [4][15]. Furthermore, sonic energizing with NaOCl was found to be an effective and promising technique in biofilm reduction in many different studies. [4][5][21][44]. Maden et at. (2017) developed a prototype device which using low electric current is able to sonically agitate the NaOCl solution. This device was able to significantly reduce biofilm in comparison to other sonic devices [19].
Chlorhexidine (CHX) is also a popular irrigant due to its antimicrobial effect [46]. It has been shown that the antimicrobial effect of sonic irrigation with 2% chlorhexidine was superior when compared to sonic saline irrigation. Additionally, it was concluded that longer exposure time to irrigants (up to 3 min) and use of CHX–Plus contributed to higher number of dead bacterial cells [32].
Alternative irrigants used in the reviewed studies were microbubble-emulsion (ME) and QMiX solution. Halford et al. (2012) examined the synergistic effect of ME and sonic agitation. This combination provided bacteria reduction 3 mm from the apical terminus, but left a considerable number of viable bacteria 1 mm from the apical terminus [47]. Interestingly, EndoActivator in combination with QMiX solution provides more favorable antibiofilm efficacy than NaOCl needle irrigation. However, as stated by the researchers, this result may also be due to chemical properties of QMIX solution in which the detergent plays an important role in weakening the biofilm structure [4].

2.2. Ultrasonic Devices

Passive ultrasonic irrigation (PUI) is a term used in endodontics for describing irrigation of root canal system without additional shaping of the canal wall [48]. With the intention to avoid possible confusion and misunderstanding, PUI will be referred to as “ultrasonic irrigation” in further text. In contrast to previously discussed studies where EndoActivator is the most commonly used sonic device, authors used different units in an attempt to enhance biofilm removal by ultrasonic agitation of irrigants.
With the aim of investigating purely mechanical effects of ultrasonic devices, only saline or distilled water was used during biofilm removal. Ultrasonic agitation of saline had proven to be more efficient in multispecies biofilm removal than simple irrigation with saline delivered by syringe and needle. This result can be due to pure mechanical effect of the ultrasonic agitation, since no antibacterial agent was used [34]. The results are in agreement with a similar research [14], that reported bacterial reduction using a comparable approach in monospecies biofilm elimination. Similarly, Grundling et al. (2011) and Hartmann et al. (2019) stated that ultrasonic irrigation with distilled water offers significant biofilm reduction when compared to manual agitation of saline with hand files. Furthermore, this research was based on a microscopy evaluations (SEM) method and confirmed a significant difference in apical and middle thirds between manually agitated saline and ultrasonic irrigation with distilled water [8][49].
NaOCl can also be used as an irrigant during ultrasonic agitation. Bhuva et al. (2010) demonstrated that ultrasonic irrigation with NaOCl is superior to saline needle/syringe irrigation in biofilm removal at all three levels of the root canal [7]. Comparatively, other studies noted similar results, although the evaluation method of biofilm removal was different and included plate counting (CFU method) [23][26][27]. In addition, it was shown that ultrasonic NaOCl irrigation offers better bacterial reduction than ultrasonic irrigation with water or saline, which can be explained by the antimicrobial effect of NaOCl [12][15]. Both the ultrasonic device and GentleWave system were effective in reducing the bacteria inside the root canal space [50].
Similarly to NaOCl, CHX can be ultrasonically agitated. Cherian et al. (2016) investigated the effectiveness of ultrasonic agitation of CHX and compared it to CHX syringe irrigation. It was concluded that ultrasonically delivered CHX provides significant bacterial reduction in comparison to syringe CHX irrigation [9]. Furthermore, Shen at al. (2010) compared the antimicrobial efficacy of CHX with CHX-Plus, both ultrasonically agitated, and found a significant difference in the number of cells killed. CHX-Plus was more efficient in biofilm reduction, which can be contributed to the chemistry of the antimicrobial agent itself [32]. Yet, when observing the research, it should be noted that HA discs were used as substrate for multispecies biofilm formation, which is notably different when compared to the morphology of the root canal system. Similarly, when activated ultrasonically, enzymes are more efficient in biofilm removal compared to saline alone [35].
Lastly, ultrasonic effect within simulated biofilm and root canal models was also investigated. For this purpose, Macedo et al. (2014) introduced a transparent root canal model with isthmus and lateral canals which were filled with hydrogel. As a result, the main canals were better cleaned with water used as an irrigant rather than NaOCl. Different from lateral canals, isthmi were equally well rinsed regardless of the agent used for ultrasonic irrigation [36]. Another study used root canal models to investigate fluid dynamics generated by syringe irrigation and both continuous and intermittent ultrasonic technique [18]. Continuous ultrasonic agitation was found to be significantly better in biofilm removal compared to syringe irrigation and intermittent ultrasonic technique. The superior action of continuous ultrasonic agitation can be due to complete oscillating amplitude of the ultrasonic tip inside the root canal which, consequently, generates maximum acoustic microstreaming. Unlike complete oscillating amplitude achieved by the continuous tip, the intermittent ultrasonic tip comes in occasional contact with the canal wall, thus resulting in weakened microstreaming effect [18]. Very recently, Mohmmed et al. explored the effect of different agitation methods using NaOCl as irrigant within 3D printed root canals [21]. The results indicated an effective biofilm removal with NaOCl ultrasonic agitation especially when compared to sonic and syringe irrigation. Additionally, microscopic images evaluations showed that 1 mm from the apex manual and sonic treatment left the biofilm intact, while complete biofilm removal at the same level was associated with ultrasonic agitation of NaOCl [21][44].

2.3. Er:Yag Laser Group

Er:Yag Laser

In one of the pioneer studies which investigated the effect of Er:Yag laser in biofilm removal, Noiri et al. (2008) directly irradiated hydroxyapatite discs that had previously been contaminated with multispecies biofilm.
However, similarly to the previous study, a uniform irradiation of dentin discs was possible due to the laboratory setup of the experiment. Complex root canal morphology in clinical conditions represents a greater challenge in biofilm removal, but, nonetheless, the results of the mentioned studies confirm beneficial effect of laser irradiation in attempts to remove biofilms.
A more relevant clinical approach was proposed by Cheng et al. (2012) who compared the results of biofilm removal using different techniques and irrigants. Although conventional 5.25% NaOCl syringe irrigation of canals was effective in eliminating the bacteria from the surface of root canals, CFU counting revealed it was not able to successfully remove E. faecalis from deep dentin layers. By applying Er:Yag laser and NaOCl as irrigant, better biofilm reduction was achieved deep inside dentinal tubules, thus suggesting that Er:Yag laser supports penetration of NaOCl. Furthermore, the research emphasized the importance of the synergistic effect of NaOCl Er:Yag laser agitation since it showed better results in comparison to NaOCl syringe irrigation or saline Er:Yag agitation [10].

Er,Cr:YSGG Laser

The pure effect of Er,Cr: YSGG laser on biofilm removal without the presence of irrigant/dry canal was investigated by Cheng et al. (2012) [10].
Surprisingly, other studies found no difference in biofilm removal between Er,Cr: YSGG 4% NaOCl agitation and 4% NaOCl syringe irrigation [15]. On the other hand, Seet et al. (2012) discovered that Er,Cr: YSGG 4% NaOCl agitation offers better biofilm eradication compared to 4% NaOCl syringe irrigation [5]. Interestingly, both authors used the identical E. faecalis strain, same period of incubation and the same irrigant concentration and time of agitation. Even though Seet et al. (2012), Betancourt et al. (2019) and Suer et al. (2020) used lower laser power settings compared to Chriso et al. (2016), they still found superior results in biofilm removal which were associated with Er,Cr: YSGG laser agitation of the irrigant, rather than conventional syringe irrigation [28][51].

Photon-Induced Photoacoustic Streaming (PIPS)

The goal of PIPS is to enhance biofilm removal by creating photoacoustic shockwaves that would travel through the root canal system which is filled with an irrigant [52]. When applying the PIPS technique, the laser tip is usually positioned in the access cavity (pulp chamber or canal entrance). Many authors are consistent in their methodologies with an emphasis that, during studies, the position of the tip was limited to the access cavity only, without further insertion towards the root canal [38][12][40][41]. Instead, De Meyer et al. (2017) inserted the PIPS tip into the canal, 6 mm short of the working length, only to discover equal effect of PIPS, regardless of the position of the laser tip [34].
In general, the PIPS technique is considered to be superior to conventional syringe/needle irrigation, regardless of the irrigant used [4][29][12][25][34][40]. Confocal laser scanning microscopy images taken by Al Shahrani et al. (2014) revealed that conventional NaOCl irrigation leaves viable bacteria deep inside dentinal tubules, while PIPS with NaOCl offers deeper penetration of the irrigant, consequently killing more bacteria [12].
When comparing PIPS to sonic agitation, Ordinola-Zapata et al. (2014) demonstrated that PIPS significantly reduces the number of bacteria within bovine root canal models when NaOCl was used as irrigant [40]. Contrarily, Balic et al. (2016) and Hage et al. (2019) concluded that both PIPS and sonic irrigation of NaOCL remove biofilm evenly from the root canal [4][53].
Up to the present time, it has been confirmed that, compared to ultrasonic techniques, PIPS offers enhanced biofilm removal in the apical part of root canals [33]. SEM images from different studies confirm PIPS superiority over ultrasonic methods in biofilm reduction inside root canals [41]. Moreover, by evaluating treatment results by CLSM and CFU, Nelaakantan et al. (2015) concluded that PIPS agitation of NaOCl and etidronic acid provides better biofilm removal when compared to conventional and ultrasonic techniques with the same irrigants [22]. Furthermore, PIPS was more efficient than sonic devices in removing hydrogel from the isthmus when using only water as irrigant [54].
Only one study compared the effect of PIPS to Er,Cr:YSGG laser in dual-species biofilm removal. The research used saline as irrigant and therefore it was possible to estimate solely the physical effect of lasers. Er,Cr:YSGG laser agitation of non-antimicrobial agent performed better at E. faecalis and C. albicans biofilm removal in comparison to PIPS [17].
Lastly, Golob et al. (2017) suggested a modified PIPS protocol, which offered promising results in disinfection of root canals [38]. Unlike the classic PIPS protocol, the authors introduced PIPS with EDTA, prior to NaOCl irrigation, and removed the mineralized part of the smear layer, opening dentinal tubules, thus enabling deeper penetration of NaOCl. Additionally, in order to increase the safety of the PIPS treatment, laser energy was reduced by 50% and no difference was found in biofilm removal between higher and lower power settings.

3. Evaluation of Biofilm Removal

The most frequently used methods for evaluating biofilm removal efficacy include counting of colony forming units (CFU) and analysis of scanning electron microscope (SEM) images, while confocal laser scanning microscopy (CLSM), polymerase chain reaction (PCR) and transmission electron microscopy (TEM) are found to be less mentioned in the reviewed studies. Additionally, it was found that some authors used more than one means of evaluation while assessing the success of biofilm removal [6][9][10][11][12][22][27].

3.1. CFU—Plate Counting

Methods of obtaining samples for further microbial analysis differ among the studies. It is suggested that after treatment protocol, root canals are filled with sterile saline, followed by syringe aspiration, centrifugation and counting of CFU [24]. Similarly, paper points leave the integrity of the dentin surface intact and have also been used in collecting samples for bacteriological evaluation [10][12]. On the other hand, Hedstrom files [14], round dental [16], Gates Gliden burs [9][22] and Peeso reamer [23] allowed researchers to retrieve dentin samples from various depths and use them for later analysis. Regardless of the sampling technique used, the CFU method provides information on the number of viable bacteria found either on the root canal surface or at various dentin depths.

3.2. SEM

An innovative proposal introduced by Bhuva et al. (2010) involves SEM image observation and analysis by endodontists with different levels of experience [7]. Briefly, a scoring system was created in relation to percentage of root canal which was covered with biofilm and dentists rated the SEM images according to their personal opinion and observations. A similar approach in SEM analysis was also used a few years later by Bhardway et al. (2013) and Ordinola-Zapata et al. (2014) [13][41]. The observed level of magnification used varies, ranging from 40 up to 10.000× [7][9][12][21][39]
Overall, SEM allows visualization of morphological structures of biofilms, their amount and distribution on dentin surface, as well as in deeper dentin layers. [8][10] However, it should be noted that sample preparation for SEM analysis might result in changes of the biofilm’s extracellular polymer matrix [2].

3.3. CLSM

Based on the reviewed papers, it was noted that the main advantage of using CLSM techniques is the researcher’s capability to distinguish viable and dead cells within biofilms. When observing the CLSM images taken after the treatment, live cells are usually seen as green, while dead cells are painted red [32][12][21]. Additionally, 3D reconstruction can be achieved and the ratio between live and dead cells can also be determined [22].


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Update Date: 07 Apr 2022