Alizarin

For fixed samples, all specimens (at 10 dpf and 30 dpf and three month old fish) were euthanized with an overdose of MS 222 and subsequently fixed for 12 h in neutral buffered 4 % paraformaldehyde. All specimens were stained for 15 min with 0.01 % ARS (3,4-Dihydroxy-9,10-dioxo-2-anthracenesulfonic acid sodium salt, from Sigma-Aldrich, St. Louis, MO) dissolved in 70 % ethanol [40 (link)]. For a better visualization of the mineralized structures in adult fish, specimens were macerated with 3 % KOH for 12 h and subsequently dissected.
For vital staining, three ARS concentrations (0.005, 0.01 and 0.05 %) were prepared in embryo medium [3 ]. The pH was adjusted to 7.4 with KOH. No precipitated ARS occurred in any of the three concentrations.
For the study of bone development, the specimens were transferred with a minimum volume of embryo medium to a new 24-well plate [3 ] with 3 ml of staining solution or new embryo medium (control). The animals remained in the staining solution for 15 min. Staining was performed once a day from 6 to 10 dpf, in each of the three ARS solutions described above. 0.2 % calcein [28 (link)] was used as a reference dye for mineral staining. In this case, larvae were stained for 10 min, as previously described [28 (link)]. Following staining with ARS, larvae were rinsed in embryo medium 3 times for 5 min, while larvae stained with calcein had to be rinsed at least 3 times for 10 min with embryo medium. In all cases, we assured that no dye residues were externally visible after the last rinsing period. If so, additional rinsing was conducted.
Stress levels were assessed by observing variations in the opercular movement frequency, as previously described [34 (link)], upon fish immersion during the first minute of staining and for 1 min at end of the staining period, before rinsing. The remaining period (remaining staining periods, and washing steps) prior to skeletal tissue imaging, took place in a dark environment to avoid stress. However, our personal observations suggest that there is no apparent effect on staining efficiency or fish health if animals remain exposed to light.
For regeneration studies, 5 adult specimens (3 month old) were exposed for 15 min to 0.01 % ARS solution prepared in system water prior to amputation and every 24 h thereafter, until 96 h post amputation (hpa). Adult fish were rinsed 3 times after each staining event for 5 min also in system water.
After ARS and calcein staining, larvae and adult fish were kept for periods no longer than 30 min prior to imaging. All specimens were anaesthetised up to 0.6 mM Tricaine solution (MS222; Sigma, St. Louis, MO) prior to microscopy analysis. Imaging was performed under green (510–550 nm) and blue (450–480 nm) fluorescent light to image ARS and calcein staining, respectively, and under visible light for total length (TL) measurements. Images were captured using a Leica MZ6 stereo microscope (Leica Microsystems, Germany) equipped for epifluorescence together with a F-View II camera, and Cell^Fv2.7 software (Olympus Soft Imaging Solutions GmbH, Germany). Higher magnifications of skeletal structures were visualised using an Axio Imager Z2 microscope equipped with a digital AxioCam ICc3 camera (Zeiss, Germany).
Tg(fli1:egfp) transgenic fish [51 (link)] were used to validate the suitability of ARS vital staining applied to GFP labelled fish during the regeneration of the caudal fin rays and the development of caudal vertebrae.
ARS staining was also used to detect skeletal deformities. The analysed deformities were not induced, but developed under regular rearing conditions. All fish were photographed using the equipment and the procedures described above.

Normal human impacted third molars (n = 18) were collected from sixteen adults (18–20 yr of age) at the Dental Clinic of the National Institute of Dental & Craniofacial Research (NIDCR) under approved guidelines set by NIH Office of Human Subjects Research and University of Southern California IRB. Root apical papilla was gently separated from the surface of the root, minced and digested in a solution of 3 mg/ml collagenase type I (Worthington Biochemicals Corp., Freehold, NJ) and 4 mg/ml dispase (Roche Diagnostic/Boehringer Mannheim Corp., Indianapolis, IN) for 30 minutes at 37°C. Single cell suspensions of SCAP were obtained by passing through a 70 µm strainer (Falcon, BD Labware, Franklin Lakes, NJ), seeded at 1×10⁴ into 10 cm culture dishes (Costar, Cambridge, MA), and cultured with alpha-Modification of Eagle's Medium (GIBCO/Invitrogen, Carlsbad, CA) supplemented with 15% FBS (Equitech-Bio Inc., Kerrville, TX), 100 µM L-ascorbic acid 2-phosphate (WAKO, Tokyo, Japan), 2 mM L-glutamine (Biosource/Invitrogen), 100 U/ml penicillin and 100 µg/ml streptomycin at 37°C in 5% CO₂. To assess colony-forming efficiency, day 10 cultures were fixed with 4% formalin, and then stained with 0.1% toluidine blue. Aggregates of ≥50 cells were scored as colonies. The proliferation rate of sub-confluent cultures (first passage) of SCAP was assessed by BrdU incorporation for 6 hours, using BrdU staining Kit (Zymed/Invitrogen). Conditions for the induction of calcium accumulation were as reported previously [18] (link). Calcium accumulation was detected by 2% Alizarin Red S (pH 4.2) staining. The induction of adipogenesis was as previously reported [8] (link). DPSCs and PDLSCs were isolated and cultured as previously described [8] (link), [9] (link). In some experiments, SCAP, DPSCs, and PDLSCs were obtained from the same donor or donors. All primary cells used in this study were at 1–3 passages. For each experiment, same passage of SCAP, DPSCs, and PDLSCs were used.

Calcium deposition was detected with alizarin red staining solution (Merck Millipore) according to the manufacturer’s instructions. Briefly, after 20 days of culture, cells were washed, fixed with ethanol 70%, incubated for 20 min with alizarin red solution, and rinsed before microscopy observation.

Cells were stained using an Alizarin red staining kit according to the manufacturer’s instructions (Solarib, China). In brief, the cells were fixed with 4% paraformaldehyde for 30 min and were rinsed with PBS. Then, the cells were stained with 0.1% (w/v) Alizarin red for 15 min. The matrix mineralization deposition appeared red, and the stained cells were photographed after washing with deionized water for 15 min. 10% ice acetic acid desorbed alizarin red for 15 min. The samples in each treatment group were added with 100 µL of ammonia and reacted at room temperature for 5 min. OD values were then determined at 405 nm.