During the initial steps of mineralization, cells produce extracellular matrix proteins and release matrix vesicles that accumulate calcium and phosphate to facilitate apatite nucleation. In this report, we compare the nucleation of minerals by two selected human cell models, osteoblastic hFOB 1.19 and osteosarcoma Saos-2 cells. Their mineralization profiles were analyzed by different methods.
Put all materials under the laminar, and sterilize them under UV light. Change the medium to 10 milliliters of fresh culture medium with fetal bovine serum. Resting cells are incubated without stimulators, whereas stimulated cells are treated with both ascorbic acid and beta-glycerophosphate.
First, gently drop ascorbic acid to final concentration of 50 micrograms per milliliter onto the surface of the culture medium to stimulate the cells. Next, add beta-glycerophosphate to final concentration of 7.5 millimolar in culture medium to stimulate the cells. Gently stir the culture dish, and put it into the incubator for seven days.
After seven days of incubation, wash the cell cultures with phosphate buffered saline, PBS. Add five milliliters of 2%alizarin red, and incubate the plate for 30 minutes to stain the minerals. Wash three times with PBS.
Carefully add PBS to the dish wall. Try not to destroy the minerals. Observe the minerals under an inverted light microscope.
Collect the medium from cell cultures, and wash the cells with PBS. Digest either resting or stimulated-for-seven-days cells with 500 units per milliliter collagenase solution at 37 degrees for three hours. Then, mechanically scrape the cells.
Transfer them to plastic microcentrifuge tubes. Pass them through a syringe 10 times. Centrifuge the samples at 500 g for five minutes.
Discard the supernatant, and suspend the cell pellet in 500 microliters of synthetic cartilage lymph. Transfer the hydroxyapatite, fluorapatite, and chlorapatite powders from the bottles on the UV transilluminator with spatula, and use as controls. Transfer the cell lysates from the plastic tubes with plastic tips, and place carefully on the UV transilluminator.
Suspend 2.5 milligrams of hydroxyapatite, fluorapatite, and chlorapatite powders in 500 microliters of deionized water. Incubate at 37 degrees for an hour. Take Formvar/carbon nickel grids from the box with antistatic forceps, and place it on a porcelain multi-well plate.
Drop 10 microliters of the hydroxyapatite suspension on the grids. Do the same for chlorapatite and fluorapatite. Dry the samples for 30 minutes at room temperature.
Prepare resting and stimulated cells for embedding. Collect medium from the cell cultures. Wash the cells in physiological desensitization medium.
Fix the cells with five milliliters of a mixture of 3%paraformaldehyde and 1%glutaraldehyde in 100 millimolar sodium phosphate buffer for an hour at room temperature under the fume hood. Wash the cells with five milliliters of 100 millimolar sodium phosphate buffer. Gently remove the buffer after washing.
In the dark room, postfix the samples with two milliliters of 1%osmium tetroxide in 100 millimolar sodium phosphate buffer. Incubate the samples for 20 minutes at room temperature under the fume hood. Remove osmium tetroxide, and utilize it.
Wash the cells with five milliliters of 100 millimolar sodium phosphate buffer. Then, dehydrate the samples in five milliliter aliquots of a graded ethanol solution series at room temperature, 25, 50, 75, and 90%ethanol. Finally, use absolute ethanol twice, and incubate for 30 minutes and 12 hours.
Mechanically scrape the cells from the culture dishes. Collect the cells into plastic microcentrifuge tubes. Centrifuge the samples at 130 g for one minute.
Remove the supernatants. Suspend the cells in a mixture of LR White resin and absolute ethanol at a volume ratio of one to two. Mix well, and incubate for 30 minutes at room temperature.
Centrifuge the samples at 130 g for one minute. Remove the supernatants. Repeat the previous step using a one-to-one mixture of LR White resin and absolute ethanol.
Mix well, and incubate for 30 minutes at room temperature. Centrifuge the samples at 130 g for one minute. Remove the supernatants.
Add pure LR White resin to the samples, and incubate for one hour at room temperature in plastic tubes. Centrifuge the samples at 130 g for one minute. Remove the supernatants.
Add again pure LR White resin. Place 500 microliters of each sample into gelatin capsules. Note, the samples are labeled using a small sheet of paper and a pencil so that the resin does not destroy the labels.
Close the gelatin capsules. Put the capsules into plastic microcentrifuge tubes. Centrifuge at 130 g for one minute in a swing-out rotor.
Remove the capsules from the plastic tubes using a vacuum pump. Move the samples to the oven, and polymerize at 56 degrees for 48 hours. Prepare the blocks by mounting them into the holder.
Put the holder into the ultramicrotome. Attach the diamond knife. Fill it with deionized water.
Remember, clean the blade from the incidental air bubbles. Then, cut sections using a diamond knife on the deionized water bath. Set the scraps using bovine eyelash.
Then, place them on the shiny side of the Formvar/carbon nickel grid, and dry them. In the dark room, counterstain the grids of synthetic apatites and cell samples with 2.5%uranyl acetate in ethanol for 20 minutes at room temperature under fume hood. Wash the grids in 50%ethanol, then in deionized water, and dry it at room temperature for 24 hours.
Finally, put the grids into the box. Prepare the beryllium holder for the observation of minerals and cells. Use antistatic tools.
Remove the pair of screws. Lift the beryllium plate and beryllium washer away from the rest of the retainer. Mount the grid, shiny side up, on the holder.
Carefully place the beryllium washer and beryllium plate. Screw tightly the fastening screws. Put the holder into the vacuum chamber, and turn on the vacuum pump.
Once a vacuum is achieved, gently insert the holder into the imaging chamber and turn on the beam. On the fluorescent monitor, set the aperture parameters of the microscope. Perform image astigmatism correction.
Set the zoom, focus, frame, and take a transmission electron microscopy image. Move the detector into the imaging chamber. Adjust the sharpness of the image in the focus mode.
Select a point in the sample for X-ray microanalysis, and collect spectra. Select elements such as calcium, fluorine, chlorine, and phosphorus to make ion mapping and perform perform maps of the selected elements. Stimulated Saos cells mineralized more efficiently than hFOB osteoblasts.
Only fluorapatites can be observed under UV light excitation. HFOB cell lysates were more fluorescent than Saos, suggesting that fluorapatites are more abundant in these cells. Stimulated Saos cells produce more vesicles containing minerals compared to hFOB cells and resting Saos cells.
Synthetic apatites had different mineral forms. The procedure applied above showed the presence of vesicles in hFOB and Saos cells under resting and stimulated conditions. The red points indicate the calcium, green phosphorus, and blue fluorine distributions.
Under stimulated conditions, there was a strong overlap between calcium and phosphorus distributions in vesicles produced by Saos cells and between fluorine and phosphorus in vesicles produced by hFOB cells. In conclusion, our findings indicate that the vesicles are key determinants of mineral nucleation, especially at the cellular level. The comparison of vesicles released from Saos-2 cells, which mineralize more efficiently, and vesicles released from hFOB 1.19 cells support the hypothesis that matrix vesicles are mineral-filled vesicles.
This leaves open the question whether the presence of matrix vesicles is required to induce apatite nucleation and how this may alter physiological to pathological mineralization.
