Introduction.
Conventional therapy of endodontically induced periapical lesions aims at eliminating the infection as completely as possible. However, there are indications that despite adequate instrumentation of the root canal, bacteria can be left in the root canal system in some cases (Chong & Pitt Ford 1992, Oguntebi 1994, Peters et al . 1995). These microorganisms and/or their by-products may interfere with the periapical healing process and can be a major cause for failures.
For this reason, experimental studies have been initiated to explore the potential of different bacteria to migrate into root canal dentine and to test the antibacterial efficacy of various intracanal medicaments (Akpata & Blechman 1982, Haapasalo & Ørstavik 1987, Ørstavik & Haapasalo 1990, Safavi et al . 1990, Buck et al . 1999, Komorowski et al . 2000, Siqueira et al . 2000). Haapasalo & Ørstavik (1987) were the first to introduce a model for infection and disinfection of root dentine using artificially infected bovine root segments. Circumpulpal root dentine was sampled and conventional microbiological culture techniques were utilized to identify bacteria. The plate count method applied in most studies gives information on the number of microorganisms able to divide at a sufficient rate to form colonies under selected laboratory conditions. However, several reports described bacteria that decreased their culturability under starvation conditions and maintained metabolic activity (Mason et al . 1986, Kaprelyants et al . 1993, Oliver 1995). Also, bacterial cell wall remnants from dead bacteria still present in dentinal tubules may interfere with periapical healing from an immunological point of view. In comparison to the classical cultivation of microorganisms, rapid direct methods detecting different physiological states of bacteria become increasingly important.
Recently, a sensitive assay for monitoring bacterial viability was introduced. It contains the nucleic acid dye Syto 9, capable of penetrating bacterial cells with intact cell walls and the counterstain propidium iodide that only labels dead bacterial cells. This two-colour fluorescent assay has already been applied in microbiological experiments with promising results (Bogosian et al . 1998, Decker & Weiger 1998, Weiger et al . 1999).
The aim of this in vitro study was to characterize the vitality status of bacteria that penetrated dentinal tubules of human root dentine by fluorescence labelling in order to differentiate between viable and dead microorganisms.

Materials and methods.

Root specimens.
Twenty-four straight roots from teeth extracted for orthodontic or periodontal reasons were selected. The teeth were either not carious or had only small carious lesions and were stored in hydrogen peroxide (3%) for a maximum of 2 months. After cleaning the root surface with curettes, the roots were removed 2–3 mm below the cemento-enamel junction. A root segment with a length of about 7 mm was prepared by sectioning the root tip. Each root canal was enlarged to size 60 with Hedstroem files under irrigation with physiological saline. The smear layer was removed with 37% phosphoric acid by leaving it in the root canal for 30 s. The specimens were sterilized in the autoclave for 20 min at 121 C. Under sterile conditions the outer root surface was coated with nail varnish and the specimens placed in a sterile vial for 1 day. Sterility was then checked by incubating each specimen in 5 mL sterile Schaedler broth at 37 C for 24 h.
As target microorganisms, Streptococcus sanguinis (Nr. 20068, Deutsche Sammlung von Mikroorganismen und Zellkulturen, Braunschweig, Germany) and Enterococcus faecalis (ATCC 29212) were selected. The root specimens were transferred into 5 mL Schaedler broth (BBL, Becton Dickinson Systems, USA) inoculated with 200 L of a 24-hour-old bacterial suspension containing either S. sanguinis (12 root specimens) or E. faecalis (12 root specimens). Under strict asepsis, the bacterial suspension was changed every second day for a period of 8 weeks. The specimens were incubated at 37 C. The purity of the infection was checked on days 28 and 56.

Study design.
The study design is outlined in Figure 1. The root dentine specimens were randomly split into the control group and the test group. Bacterial samples from root dentine (rd) were taken on days 28 (week 4), 56 (week 8, control group) and 84 (week 12, test group). On days 28 and 56, 1 mL from the environmental bacterial suspension (sus) harbouring the root specimens was taken for microbiological analysis.
At week 4, all 24 root canals were irrigated with 1 mL physiological saline and cleaned using sterile paper points to remove microorganisms attached to the root canal walls. Subsequently, the root canals were prepared from size 60 to 90. For this purpose, Hedstroem files size 25 were used to obtain fine dentine chips, which were collected in a tube filled with 1 mL Schaedler broth. The Hedstroem files size 25 were agitated thoroughly in this solution several times. Finally, two paper points were used to take up remaining dentine shavings on the root canal walls and then added to the Schaedler broth. The smear layer was again removed with 37% phosphoric acid to allow bacterial penetration into dentinal tubules.

Figures 1. Study design (rd, root dentine; sus, bacterial suspension).

At week 8, the collection of ‘rd’ samples was limited to the 12 root specimens of the control group (Fig. 1). For this purpose, the root canals were enlarged to size 120. The test group consisted of the other 12 root specimens. Their root canals were irrigated with 1 mL of physiological saline and cleaned using sterile paper points. Afterwards, pure calcium hydroxide powder was mixed with distilled water in a powder-liquid ratio of 1 g mL –1 (pH = 13) and then condensed into the root canal with paper points. The cervical and apical openings of the filled root canals were closed with Cavit (ESPE, Seefeld/Oberbayern, Germany) and the root specimens stored in a wet gauze at 37 C for an additional 28 days.
At week 12, the calcium hydroxide was removed by rinsing the root canals twice with 5 mL of distilled water. After the first rinse the root canal walls were cleaned with sterile paper points. Finally, the root canals were enlarged to size 120 to provide ‘rd’ samples in the same way as described above.

Microbiological analysis.
The ‘rd’ and ‘sus’ samples were processed in the laboratory immediately. The portions of viable bacteria (PVB rd , PVB sus ) and the number of colony-forming units (CFU rd , CFU sus ) were determined as follows:

PVB rd and PVB sus

The samples were centrifuged for 5 min at 8240 g. After washing with sterile saline, a second centrifugation step followed. The pellet was then resuspended in 300 L staining solution containing two fluorescent stains, Syto 9 and propidium iodide (Live/Dead BacLight Bacterial Viability Kit, MoBiTec, Göttingen, Germany). These two substances allowed the differentiation between viable and dead microorganisms. After 15 min, the samples were again centrifuged and the stained pellet was analysed by a fluorescence photomicroscope (Zeiss, Jena, Germany) with a suitable filter combination (FITC: 450– 490 nm; Rhodamin: 540 nm) allowing to count the number of green (= viable) and red (= dead) bacterial cells in 10 visual fields at a magnification of 640 . PVB represents the number of viable microorganisms related to the total number of stained microorganisms (= sum of viable and dead bacteria). PVB was given as a percentage.

CFU rd and CFU sus

The original samples were diluted up to a concentration of 10 –5 for S. sanguinis and 10 –6 for E. faecalis . Portions of 20 L were inoculated onto Schaedler agar supplemented with sheep blood and Vitamin K 1 . Following anaerobic incubation (Anaerocult A, Merck, Darmstadt, Germany) for 48 h at 37 C, visible colonies were counted as colony-forming units (CFU) and related to 1 mL of the original sample.

Data analysis.
The CFU values were log transformed. The mean values of PVB rd and CFU rd with the corresponding 95% confidence intervals were calculated. As the ‘rd’ samples were of particular interest, the following differences PVB rd and CFU rd were given for each sample:

• Control group: PVB rd (week 8) – PVB rd (week 4) log CFU rd (week 8)
  – log CFU rd (week 4), if growth was present.
• Test group: PVB rd (week 12) – PVB rd (week 4). log CFU rd (week 12)
  – log CFU rd (week 4), if growth was present.

Positive values indicate that PVB rd or CFU rd increased within the given period of time, whilst for negative values PVB rd or CFU rd decreased.

Table 1. Mean values with the corresponding 95% confidence intervals of the portion of viable bacteria (PVBrd) and the number of colony-forming units (CFUrd) in root dentine (rd) collected after 4, 8 and 12 weeks.


Table 2. Comparison between baseline PVBrd values (week 4) and PVBrd values recorded at week 8 (control group) and at week 12 (test group). The individual differences  PVBrd = PVBrd (ti) - PVBrd (tbaseline) are depicted (d: mean value). Positive values indicate that PVBrd increased within the given period of time, whilst for negative values PVBrd decreased.


Table 3. Comparison between baseline CFUrd values (week 4) and CFUrd values recorded at week 8 (control group) and at week 12 (test group). The individual differences  log CFUrd = log CFUrd (ti) - log CFUrd (tbaseline) are depicted (d: mean value). Positive values indicate that CFUrd increased within the given period of time, whilst for negative values CFUrd decreased.

Results.
Fluorescent-labeled bacteria were found in all ‘rd’ samples During the period of infection, PVB sus ranged from 55 to 75% ( S. sanguinis ) and from 35 to 60% ( E. faecalis ), respectively. Likewise, viable as well as dead microorganisms were identified in root dentine at any time. In the majority of the cases, PVB rd values were lower than PVB sus values, regardless of the bacterial strain and the point of time.
In the control group, mean values for PVB rd did not markedly differ at week 4 and at week 8 for S. sanguinis and E. faecalis (Table 1). The individual PVB rd values varied between 13 and 61%. The differences PVB rd and CFU rd are specified in Figures 2 and 3. Mean values for CFU rd ranged between log 6.5 and 8.6 (Table 1).
In the test group, PVB rd decreased from 36% to 27% (mean values) after placement of calcium hydroxide in specimens infected with S. sanguinis (Table 1). The individual PVB rd values ranged from 13 to 52%. The differences PVB rd substantiated the decrease of PVB rd in most samples, indicating that viable S. sanguinis cells were still present in root dentine after 12 weeks, although in culture no growth was detectable in any sample (Figs 2, 3). By contrast, the majority of the samples infected with E. faecalis demonstrated somewhat higher PVB rd values after calcium hydroxide treatment than initially after 4 weeks (Fig. 2). CFU rd varied to a certain extent when comparing week 12 with week 4 (Fig. 3).

Discussion.
In the present study, extracted human teeth were selected as the test specimens, although bovine teeth were used in the majority of in vitro investigations related to infected root dentine (Haapasalo & Ørstavik 1987, Ørstavik & Haapasalo 1990, Vahdati et al . 1993, Perez et al . 1996, Siqueira & De Uzeda 1996, Heling & Chandler 1998, Komorowski et al . 2000). Bovine root dentine, however, differs from human dentine; dentinal tubules of bovine teeth have wider diameters (Perez et al . 1993) and in some cases giant tubules are present. This may affect the microenvironment of bacteria (e.g. diffusion of nutrients or medicaments into dentinal tubules) and their vitality status. Similarly, the presence of intact root cementum can limit bacterial ingrowth from the pulpal side in deeper layers of root dentine, due to limited availability of the nutrient source (Haapasalo & Ørstavik 1987, Adriaens et al . 1988). This indicates that microorganisms in dentinal tubules may be different in a nutrient-rich or nutrient-deficient environment. Therefore, infected human root specimens seem to be more suitable for investigating the microbial vitality status than infected bovine root dentine.
Under stressful conditions (e.g. starvation), bacterial populations may retain certain physiological functions although they do not grow under conventional culture conditions. These viable but non-culturable cells (VBNC) may survive and recover or go in a prelytic or lytic state. Undoubtedly, the existence of VBNC can only be suggested. An approach to substantiate the VBNC theory is to combine different viability tests examining, for example, respiration, enzymatic activitiy, membrane potential or membrane permeability. In the present study bacterial viability was characterized by conventional culturing and by applying the two stains, Syto 9 and propidium iodide. The latter provides information on the cytoplasmic membrane permeability that is a key element of cell viability. The two stains differ in their spectral characteristics and their ability to penetrate bacterial cell membranes. Viable bacteria with intact bacterial cell membranes stain fluorescent green. However, information is scarce as to whether or not these labelled organisms are able to divide under the environmental conditions prevailing in the root canal system. In growth cultures the majority of S. sanguinis cells that stained fluorescent green are viable, i.e. they form colonies (Decker & Weiger 1998). In contrast, microorganisms with damaged cell membranes accumulate propidium iodide in the cell body and stain fluorescent red; they are scored dead. Experimental results with S. sanguinis as the test organism showed that these cells were not capable of dividing after conventional culturing (Bogosian et al . 1998, Decker & Weiger 1998). A recent study demonstrated bacterial cells identified by propidium iodide to be permeabilized and irreversibly damaged (Nebe-von Caron et al . 1998).
Streptococcus sanguinis and Enterococcus faecalis were chosen as target microorganisms. Both are frequently encountered in infected root canal systems, and particularly Enterococcus faecalis is a species commonly recovered in teeth with failed root canal treatment (Siren et al . 1997, Sundqvist et al . 1998, Molander et al . 1998). In the present study smear layer was removed after root canal preparation of the specimens to facilitate bacterial migration into root dentine from the pulpal side as previously shown for S. sanguinis (Perez et al . 1996). Our experiments demonstrated that an infection time of 4 weeks allowed bacteria to penetrate human root dentine up to a depth of at least 150 m. The removal of infected circumpulpal root dentine with small files (size 25) resulted in very fine dentine shavings. These facilitated the microscopic analysis of the stained samples, namely the detection of green fluorescent bacteria, due to the reduced autofluorescence of the dentine dust, allowing the accurate determination of PVB rd . It appeared unlikely that the ‘rd’ samples still contained bacteria attached to the root canal walls, because the rinsing procedure prior to ‘rd’ sampling occurred under favourable conditions (straight and short root canal prepared to size 60 [at week 4] or 90 [at week 8] with a wide apical opening). Similarly, calcium hydroxide was thoroughly removed from the root canals at week 12. Preliminary experiments demonstrated that calcium hydroxide could neither be detected macroscopically nor microscopically on root canal walls. Furthermore, it might be speculated whether or not bacteria were pushed deep into dentine during preparation of the root canal walls with Hedström files. It is possible that some bacteria were pushed into the superficial dentine layers and were not included in the analysis.
Viable bacterial cells multiplying under the selected laboratory conditions, as well as dead microorganisms, were identified in all root dentine samples regardless of the bacterial strain used. However, whether all viable and green fluorescent microorganisms were capable of proliferating remains unanswered. Nevertheless, the portion of viable bacteria found in root dentine was reduced compared to the one present in the bacterial suspension in the root canal lumen. Most probably, some of the bacteria attached to the inner surfaces of dentinal tubules could not resist the specific conditions prevailing in root dentine, whilst they would have survived as planktonic cells in the surrounding bacterial suspension.
As expected, the degree of tubule infection varied from sample to sample. For that reason, the baseline dentine samples collected after 4 weeks served as individual controls for the samples taken after 8 weeks (control group) and 12 weeks (test group), respectively. When analysing each root specimen of the control group, the parameters PVB rd and CFU rd substantiated that the degree of infection established within the first 4 weeks largely corresponded to that re-established in the following 4 weeks after ‘rd’ sampling (weeks 5–8). Therefore, it was justified to take the ‘rd’ samples at week 4 as basis for those collected at week 12 in the test group. In these cases, no infected circumpulpal dentine was removed at week 8, as calcium hydroxide should demonstrate its efficacy toward infected root canal dentine.
With S. sanguinis , no growth on blood agar plates was observed after exposure of root dentine to calcium hydroxide for 4 weeks. However, in all these ‘negative’ ‘rd’ samples green fluorescent and viable bacterial cells were detected. Obviously, these were not culturable under the selected laboratory conditions and have not been detected in other comparable studies so far. These S. sanguinis cells may play a role in treatment failure when nutrients are available. With E. faecalis , PVB rd tended to increase, indicating the presence of a large number of viable E. faecalis cells in the root dentine despite calcium hydroxide treatment. These results confirmed various investigations showing that E. faecalis resists calcium hydroxide treatment (Haapasalo & Ørstavik 1987, Ørstavik & Haapasalo 1990, Siqueira & De Uzeda 1996).

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