Sample collection and stem cell isolation
Teeth were collected from the outpatient clinic of the Excellence Medical unit of the National Research Centre. All teeth were premolars indicated for extraction as a part of the Orthodontic treatment plan of the children involved. All cases were approved by the National Research Centre’s ethical committee. All donors were apparently healthy aged from 16 to 20 years. The collected human dental pulp tissues were obtained from the extracted premolars teeth following the protocols described by Eduardo et al. (2008) and Gronthos et al. (2000). Prior to extraction, patients received chlorhexidine mouth rinse to reduce oral microbial flora. Extraction was as atraumatic as possible in complete aseptic condition. The freshly extracted premolar was then grooved using a diamond stone at nearly the level of the cemento-enamel junction under sterile conditions using sterile saline coolant to facilitate later splitting. The groove surrounded the whole crown till half the depth of dentin without exposing the pulp. The tooth was immediately placed in a sterile tube containing phosphate buffer solution (PBS) and transferred to the laboratory in less than 2 h using a sterile cement spatula under the laminar; the crown was split; and the pulp was gently removed in sterilized conditions using sterile tweezers, a small excavator, and K-files and then immediately placed in PBS.
Sample preparation
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The extirpated pulp tissues were washed in PBS three times, and the specimens were placed in a small plate and minced into small pieces 1 mm3 using sterile scissors and surgical blades.
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The minced pieces were collected in sterile, labeled 1.5-ml Eppendorf tubes.
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Enzyme digestion was carried out according to Gronthos et al. (2000). The digested solution used was 3 mg/ml collagenase type I (Sigma-Aldrich, USA) for 60 min at 37 °C.
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The digestive reaction was stopped by the addition of culture media. The culture media consisted of alpha modified Eagle’s medium (αMEM) with l-glutamine (Gibco, Invitrogen Life Technologies, USA) supplemented with 10% fetal bovine serum (Gibco, Invitrogen Life Technologies, USA), antibiotics, penicillin/streptomycin (penicillin G100 unit/ml and streptomycin 100 μg/ml), and finally antimycotic agent (Fungizone, 0.25 μg/ml).
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Single-cell suspensions were obtained by passing the digested tissues through a 70-μm cell strainer (Becton/Dickinson, USA). The strainer also ensured the removal of debris.
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The tubes were centrifuged for 20 min at room temperature to obtain a cell pellet of pulp-derived cells.
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The supernatant was discarded, and then the cells in the pellet were resuspended in complete culture media by successive pipetting.
At this point, 1 ml of the tube was taken and 0.2% Trypan Blue stain (Sigma–Aldrich, UK) was mixed with the cell suspension and incubated for 5 min at room temperature, then placed on a Nubarau hemocytometer. The plates were labeled by cell type and date and incubated at 37 °C in a humidified atmosphere of 5% CO2. The medium was changed once every 4 days.
Sub-culturing “passaging”
Passaging was performed when the primary cell culture of adherent cells reached 70% confluence and was named passage zero (P0), and later passages were named accordingly.
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Primary cell cultures of adherent cells were detached by treatment with a sterile solution of trypsin/EDTA for 5–10 min at 37 °C in the incubator and shaken intermittently.
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The used trypsin was prepared with the dissolution of 2500 mg trypsin powder in 100 ml PBS solution and filtered onto 2-mm filters. Fetal bovine (100 μl) serum was added to inactivate the trypsin.
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The cells were then collected by centrifugation at 200 rpm for 10 min, and the cell pellet obtained was resuspended in 1-ml complete medium and divided into two plates (passaging) both followed by immersion in complete culture medium to increase cell numbers.
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Thus, the primary cell culture was propagated and expanded in repeated cell cultures.
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Cells were sub-cultured every other week, and the culture medium was replaced every 3 days over a 10–14-day period. All the previous procedures were conducted under aseptic conditions in an air-filtered laminar flow safety cabinet using sterile instruments.
Experimental groups
The culture was divided into 3 groups according to the LLLT energy density of irradiation:
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Group I: no irradiation (control)
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Group II: irradiation at dose 0.5 J/cm2for 20 s
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Group III: irradiation at dose 1 J/cm2for 20 s
Low-level laser therapy irradiation
- Stem cells were seeded in a 96-well plate at a density of 1 × 104 cells/well and allowed to attach overnight.
- Low-level laser therapy irradiation (LLLT) irradiation was applied in two doses into the 96 wells by the diode laser device (970 nm) through the fiber optic (SiroLaser fibers 320) at a distance from the opening of the wells to be accurate for equal distribution of the laser irradiation.
The energy density (fluence) calculated was equal to:
$$ \frac{\mathrm{Power}\times \mathrm{time}}{\mathrm{Area}}\kern0.48em \mathrm{J}/{\mathrm{cm}}^2 $$
-All doses were executed in continuous wave and noncontact mode
- Prior to irradiation, the media of all wells were replaced by fresh media supplemented with 15% FBS, to ensure the removal of any dead cells.
Assessing cell morphology and proliferation capability
Daily follow-up of stem cells was done to ensure cell viability. Estimation of the cell morphology and proliferation of pulp-derived stem cells were monitored in primary cultures and sub-cultures. The proliferation capacity was judged by close follow-up of confluence rate, i.e., culture plates reaching 70% confluence according to culture days. Cultures from DPSC were monitored using an inverted light microscope (Olympus, USA) with a digital camera for capturing images (Nikon, Japan). Furthermore, the cells were tested for the ability to form colonies where MTT assay was performed to compare the proliferation of the cells. A suspension of 100 cell/ml was cultured in a 3.5-cm dish in complete culture media, and the cultures were observed under an inverted light microscope. Aggregates with more than 50 cells were scored as colonies.
MTT assay
MTT assay according to the standard procedure after the cells were irradiated was performed. An indirect method measuring the metabolic activity of mitochondrial enzymes was used to determine viable cell number in the cultures. The assay is based on the cellular conversion of a tetrazolium salt [MTT: 3(-4, 5-dimethylthiazol-2-yl) 2, 5-diphenyltetrazolium bromide] to formazan. Control and treated cells were incubated with MTT (0.2 mg/ml) diluted in the appropriate media, for 60 min at 37 °C in 96-well plates. The culture medium was removed and formazan was solubilized. The extent of reduction of MTT to formazan within cells was quantified by using a spectrophotometer at a wavelength of 572 nm. Absorbance is directly proportional to the number of living cells in culture. Cell proliferation was measured by MTT assay which was performed after 24 h, 48 h, and 72 h of irradiation according to the cell proliferation kit protocol (Vybrant, Invitrogen). The experiment was repeated in triplicates. This protocol is optimized for dental pulp stem cells.
