Sample collection and preparation
The ethics committee of Shanxi Provincial People’s Hospital approved the study protocol [Provincial Medical Opinions (2019) No. 3]. Palatal roots of the maxillary first molars and distal roots of the mandibular first molars were used for this in vitro study for the following reasons. First, the required sample capacity of this experiment was large, so palatal and distal roots were collected to ensure a sufficient sample size. Second, the diameter of the palatal and distal roots is relatively large, which is convenient for the preparation of the Enterococcus faecalis biofilm model. The selected teeth were untreated and had completely formed apices and root canals. All samples were scanned using KaVo 3D eXam cone-beam CT (KaVo, Germany) to observe the morphology of the root canal system from different directions. Type Ⅰ (1-1) palatal and distal root canal systems were selected, excluding curved and other root canal system types. Any calculi and periapical soft tissues were first removed by using an ultrasonic scaler. All samples were then placed in a saline-filled test tube and stored at 4 °C until use. For the study, the teeth were decoronated using a diamond bur. The standard length of the remaining root was 12 mm. The working length (WL) was set as the standard length minus 1 mm (i.e., 11 mm). K-files (#10 and #15; Dentsply Maillefer, Ballaigues, Switzerland) were used to create a glide path for the WL, and ProTaperNext rotary files (Dentsply Sirona, York, PA, USA) were used to shape the canals up to size X3 (0.3 mm, 7%) according to the manufacturer’s instructions. The shaping of all root canals in this study was performed by the same operator. First the 10 # and 15 # K-files were used in order to establish the root canal access, and then X1, X2, and X3 Ni-Ti files were used to shape the root canal in turn; all shaping methods and parameters were selected in accordance with the manufacturer's instructions. Using a Gates Glidden drill #5, a groove 4 mm in length, 1 mm deep, and 0.4 mm wide was made in the wall of each root canal. This groove served as a coronal reservoir for irrigated placement. After the use of each instrument, the canals were irrigated with 2.5 mL 3% sodium hypochlorite (NaOCl) solution (Sigma-Aldrich Corporation, St. Louis, MO, USA), which was delivered via a syringe with a 27-G side-vented needle. After completing the preparation, the canals were sequentially irrigated with 5 mL of 17% ethylenediaminetetraacetic acid (EDTA; Ultradent Products, Inc., South Jordan, UT, USA) and 5 mL of 3% NaOCl for 5 minutes each; this ensured removal of the smear layer. The canals were then flushed for 15 minutes with 20 mL of 0.9% physiological saline for the removal of any residual EDTA or NaOCl solution. The samples were dried at room temperature. To prevent bacterial leakage, we sealed the apical third of all roots with a composite resin and coated the entire root surface with nail polish. Finally, five teeth were randomly selected and subjected to scanning electron microscopy (SEM) (EVO MA10; ZEISS, Oberkochen, Germany) for confirmation of smear layer removal.
The following scoring criteria were used for smear layer removal: 1, no smear layer, 100% dentinal tubules open; 2, small amount of scattered smear layer, 80% dentinal tubules open; 3, thin smear layer, 60% dentinal tubules open; 4, a portion of the root canal wall covered with a thick smear layer; and 5, root canal wall completely covered by a smear layer. Two experts in the field of dental pulp disease evaluated the SEM images using a double-blind method. A score of 3 or lower met the smear removal standard. SEM images were used for two reasons—first, the smear layer was composed of assorted debris and infectious substances; however, in this study the single bacterial infection model (i.e. the E. faecalis infection model) had to be established. Thus, it was important to remove as much of the smear layer as possible to minimize the possibility of the presence of bacteria other than E. faecalis in the root canal system. Secondly, the presence of the smear layer in would have hindered the colonization of E. faecalis in the dentin tubules and led to failure of establishing the E. faecalis infection model.
The samples were subsequently placed in glass test tubes filled with 0.9% physiological saline sterilized at 121 °C in a 1.5-Mpa autoclave (LS-150LD; Binjiang Medical Equipment Ltd., Jiangyin, China) for 30 minutes. An inoculating loop was used to collect a loopful of the liquid near the root canal in the test tube. This liquid was inoculated on a sterile plate with blood agar medium and placed in a Tri-Gas incubator (HF-100; Heal Force Bio-meditech Holdings, Ltd., Shanghai, China) for 24 hours. The effects of sterilization were determined by observing the colony growth on the plate.
Establishment of the E. faecalis infection models
A standard E. faecalis strain (ATCC 29212), which was procured from stocks in the Microbiology Laboratory of Shanxi Provincial People’s Hospital, was activated and formulated into a bacterial suspension, the concentration of which was adjusted to 1.0 MCF on an electronic turbidimeter (BioMerieux, Mercy l’Etoile, France). Five tooth samples were placed in a glass tube containing 1 mL of Enterococcus broth (HB0133-2; Haibo Biotechnology Co., Ltd, Qingdao, China) and 1 mL of the E. faecalis suspension (in total, 25 tubes), which were incubated for 4 weeks at 37 °C in a Tri-Gas incubator (HF-100; Heal Force). Every 48 hours, the liquid in the tube was changed. At the time of culture solution replacement, 1 mL of liquid near the root canal was collected and incubated for 24 hours on a plate containing blood agar medium. The presence of other bacteria was ruled out by analyzing the formed colonies using a fully automated rapid mass spectrometry detection system (Microflex LT/SH; Bruker Daltonik, Bremen, Germany). After 4 weeks, in vitroE. faecalis infection models were successfully established. To confirm E. faecalis colonization, five samples were randomly selected and observed by using SEM.
Bacterial sampling and counting before irrigation
In a biosafety cabinet (HFSafe 1200; Heal Force Bio-meditech Holdings, Ltd.), the culture solution in the canals was carefully blotted by using sterile paper tips. The canals were then rinsed with 1 mL of 0.9% sterile saline to flush out unattached bacteria. Thereafter, three sterile paper tips saturated with 0.9% saline were successively inserted up to the WL and repeatedly rubbed against the inner canal walls. After 1 minute, the paper tips were placed in 1 mL of 0.9% sterile physiological saline and shaken on a vortex mixer (XW-80A; Jingke Industrial Co., Ltd., Shanghai, China) for 5 minutes for deployment of the bacterial suspension. This suspension was serially diluted with physiological saline (up to 10-6), with a volume ratio of 1:10. To count the bacteria, 0.1-mL aliquots containing appropriate dilutions of each sample were spread onto blood agar plates and incubated for 24 hours at 37 °C in the Tri-Gas incubator (HF-100; Heal Force Bio-meditech Holdings, Ltd.). The colony-forming unit (CFU) number in the entire plate was then counted and recorded.
Irrigation protocols
We randomly divided the 120 roots into two experimental groups and one control group (n = 40 each) according to the irrigation protocol: 3% NaOCl activation with an Er:YAG laser (LightWalkers ATS; Fotona, Ljubljana, Slovenia) using the SWEEPS mode for 60 seconds (SWEEPS group); 3% NaOCl activation with an Er:YAG laser (LightWalkers ATS) using the PIPS mode for 60 seconds (PIPS group), and 3% NaOCl irrigation without activation for 60 seconds (control group).
The parameters for the SWEEPS and PIPS modes are presented in Table 1. In the SWEEPS group, the canals were first subjected to 3% NaOCl (2 mL each) activation using the SWEEPS mode for 20 seconds, followed by a rest interval of 20 seconds, 0.9% saline (2 ml each) activation using the SWEEPS mode for 20 seconds, and then another rest interval of 20 seconds. This procedure was repeated three times. The same regimen was followed in the PIPS group, where NaOCl was activated using the PIPS mode. In the control group, the canals were subjected to three 20-second cycles of 3% NaOCl (2 mL each) irrigation via a syringe with a 27-G side-vented needle without any activation, 0.9% sterile physiological saline irrigation (2 mL each) for 20 seconds, and a rest period of 20 seconds.
The No. 99128 fiber tip was used in the SWEEPS mode. This fiber tip has a diameter of 0.60 mm, a length of 12 mm, and a flat end. The No. 89036 fiber tip was used in the PIPS mode. This fiber tip has a diameter of 0.60 mm and a length of 9 mm. Unlike that of the fiber tip used in SWEEPS mode, the end of the fiber tip in PIPS mode is tapered. During activation, the fiber tip was placed into the coronal reservoir which was 10 mm from the WL. The SWEEPS and PIPS fiber tips are shown in Figure 1a and b.
Bacterial sampling and counting after irrigation
The method used for bacterial sampling and counting after irrigation was the same as that used before irrigation. On the basis of the obtained values, we calculated the bacterial reduction rate by using the following formula: bacterial reduction rate (%) = (E − F)/E × 100. In this paper, E and F represent the number of bacterial colonies before and after irrigation, respectively. Based on the bacterial reduction rate, the bacterial clearance efficacy of each method was evaluated. For more intuitive demonstration of the bacterial clearance efficacy, the number of remaining bacteria in the irrigated samples from the three groups was observed by SEM.
Statistical analysis
All data were statistically analyzed using SPSS, version 22.0 (IBM, Chicago, IL, USA). We used the Kruskal–Wallis test to detect statistically significant differences between groups. Intergroup comparisons were conducted using nonparametric one-way analysis of variance. The bacterial reduction rate are presented as medians and interquartile ranges. A p-value of ≤0.05 was considered statistically significant.