Sample collection and preparation
The ethics committee of Shanxi Provincial People’s Hospital approved the study protocol [Provincial Medical Opinions (2019) No. 3]. Distal and palatal roots of freshly extracted human molars were used for this in vitro study. The selected teeth were untreated and had completely formed apices and root canals. 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. 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.
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 ofthe Enterococcus 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 vitro E. 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 subjected to three cycles involving 3% NaOCl (2 mL each) activation using the SWEEPS mode for 20 seconds, 0.9% sterile physiological saline irrigation (2 mL each) for 20 seconds, and a rest interval of 20 seconds. 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.
Table 1. Parameters for Er:YAG laser irrigation of root canals using the shock wave-enhanced emission photoacoustic streaming mode and the photon-induced photoacoustic streaming mode
Mode
|
Pulse (mJ)
|
Frequency (Hz)
|
Power (W)
|
λ (μm)
|
Water and air
|
SWEEPS
|
20
|
15
|
0.3
|
2.94
|
Turn off
|
PIPS
|
20
|
15
|
0.3
|
2.94
|
Turn off
|
Er:YAG: erbium-doped yttrium aluminum garnet, PIPS: photon-induced photoacoustic streaming, SWEEPS: shock wave-enhanced emission photoacoustic streaming
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 clearance rate by using the following formula: bacterial clearance 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 clearance rate, the bacterial clearance efficiency of each method was evaluated. For more intuitive demonstration of the bacterial clearance efficiency, 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 clearance rates are presented as medians and interquartile ranges. A p-value of ≤0.05 was considered statistically significant.