Differentiation of Functional Osteoclasts from Human Peripheral Blood CD14⁺ Monocytes

Buffy coats obtained from the Scottish National Blood Transfusion Service (Edinburgh) and leucocyte cones obtained from NHS Blood and Transplant (Newcastle) are provided to University of Glasgow researchers in a fully anonymized (non-identifiable) form from fully consenting NHS blood donors. The buffy coat and leukocyte cone blood components are produced from an NHS standard blood donation given at an NHS blood donor center in Scotland or England. The blood donor gives informed consent at the time of the blood donation for surplus blood not used in standard NHS clinical practice to be used for approved medical research studies. The ethical approval from the NHS Research Ethics Committee and the signed donor consent forms to use these blood donations are held by the NHS blood donation service. Approval to access and use these consented blood donations in approved medical research studies was sought and gained using the standard internal application and review process of the National Blood Transfusion Service (Scotland) and NHS Blood and Transport (England). No further NHS REC approval or internal University of Glasgow ethical committee approval was required to use the blood components for the approved medical research studies.

1. General notes prior to starting

1. Proceed with all work with blood with caution. Consider the potential hazards of various infectious agents that may be present in the samples.
2. Conduct all the work cautiously in the biosafety laboratory under sterile conditions while wearing gloves and lab coats.
3. Perform biosafety disposal according to the local guidelines.
4. Obtain appropriate consent and ethical approvals prior to sample collection according to local authority regulations.
5. Generally, 1 mL of fresh blood will yield 1 million PBMCs, and CD14⁺ monocytes account for approximately 10%-30% of PBMCs. In comparison, 10 mL of a leukocyte cone can contain 5 x 10⁸-15 x 10⁸ PBMCs. For further details on PBMC isolation, refer to a previous protocol³⁷.

2. Isolation of peripheral blood mononuclear cells (PBMCs) from whole blood

1. Collect the required volume of fresh blood from healthy donors into lithium heparin collection tubes.
   ​NOTE: Other collection tubes with appropriate anticoagulants can also be used (e.g., sodium heparin tubes). For higher cell counts, leukocyte cones or buffy coats can be used.
2. To isolate the PBMCs, transfer the blood into a new 50 mL tube, and dilute it with sterile 1x phosphate-buffered saline (PBS) in a 1:1 or 1:3 ratio for fresh blood or leukocyte cones/buffy coats, respectively.
3. Gently mix the cells several times by inversion.
4. Prepare 15 mL tubes containing 3 mL of density gradient medium. Slowly layer 8-10 mL of diluted blood on the top of the density gradient medium, and centrifuge at 400 x g for 30 min at room temperature (RT) with no brake.
   NOTE: Apply the blood carefully on top of the density gradient medium to prevent mixing. Mixing might result in the loss of PBMCs.
5. Carefully discard the top layer (containing the plasma) with a Pasteur pipette, collect the underneath interphase layer containing the PBMCs (white, ring-like structure), and transfer this layer into a new 50 mL tube.
6. Suspend the cells with sterile 1x PBS up to 50 mL, and wash off the residual density gradient medium by centrifuging at 300 x g for 10 min at RT with full brake.
7. To remove residual platelets, repeat the process with an additional slower spin at 200 x g for 10 min at RT with no brake.
8. Optional: To remove the carryover of red blood cells, dilute red blood cell lysis buffer (10x, Table of Materials) 1:10 in distilled water, and apply 3 mL of the diluted buffer onto the pellet. Mix, and incubate for 3 min. Wash the pellet in up to 50 mL of 1x PBS, and centrifuge at 300 x g for 10 min at RT with full brake.
9. Resuspend the isolated and purified PBMCs in 20 mL of 1x PBS and count them using a hemacytometer or following other standard methods.
   NOTE: The cell dilution and cell counting methods must be adjusted accordingly based on the cell density and counting device used.

3. Enrichment of CD14⁺ monocytes from PBMCs

1. Isolate CD14⁺ monocytes from the PBMCs with a human CD14⁺ selection kit according to the manufacturer's protocol (Table of Materials).
   NOTE: Classical CD14⁺CD16⁻ monocytes are the main source of OC precursors³⁵; alternative purification methods can be considered.
2. Transfer 1 x 10⁷ PBMCs into a suitable polystyrene round-bottom tube (i.e., that fits the selection kit magnet), and pellet the cells at 300 x g for 5 min.
3. Discard the supernatant, resuspend the cell pellet in cell separation buffer (PBS, 2% fetal bovine serum [FBS], 1 mM ethylenediaminetetraacetic acid [EDTA]) to a final concentration of 1 x 10⁸ cells/mL, and incubate with 10 µL of antibody cocktail per 100 µL with the lid on for 10 min.
   NOTE: The cells are resuspended at 1 x 10⁸ cells/mL; adjust the volumes accordingly.
4. After incubation, add 10 µL of the magnetic nanoparticle beads per 100 µL, and incubate for 3 min with the lid on.
   NOTE: Adjust the volumes of the antibody cocktail and magnetic beads to obtain a concentration of 10 µL/mL.
5. Top up the volume to 2.5 mL with the cell separation buffer, place the tube into a magnet (without the lid), and incubate for 3 min. Discard the negative cell population by one continuous move by inversion while the tube is still in the magnet.
   NOTE: If using >2 x 10⁸ PBMCs and a larger magnet, top up to 5 mL or 10 mL with cell separation buffer following the manufacturer's instructions.
6. Remove the tube from the magnet, and wash the enriched CD14⁺ monocytes attached to the magnetic beads by resuspending them in 2.5 mL of the cell separation buffer. Incubate for 3 min inside the magnet as before, discard the negative fraction, and repeat one more time.
7. Centrifuge all the collected cells at 300 x g for 5 min, discard the supernatant, and resuspend the cells in 5 mL of alpha minimum essential medium (α-MEM; Table of Materials) supplemented with 1% L-glutamine, 1% penicillin/streptomycin (complete α-MEM), and 10% FBS.
   ​NOTE: A post-enrichment purity check via flow cytometry is recommended, and ≥96% purity should be expected. Additional washing steps (step 3.6) can increase the purity.

4. OC differentiation in vitro

1. Count the enriched CD14⁺ monocytes using a hemacytometer.
2. Pellet the cells at 300 x g for 5 min, and resuspend at 1 x 10⁶ cells/mL in complete α-MEM supplemented with 10% FBS.
3. To differentiate the OCs, add M-CSF at a final concentration of 25 ng/mL to the cell suspension.
   NOTE: For 1 mL of cell suspension, add 0.25 µL of M-CSF from a stock concentration of 100 µg/mL.
4. Mix by pipetting thoroughly to homogenize the cell suspension and plate 100 µL/well in a flat-bottom 96-well plate to a final cell density of 1 x 10⁵ cells/well.
5. Add 200 µL/well of sterile distilled water into the wells around the plated cells to prevent medium evaporation and edge effects in the culture system.
6. Incubate the cells overnight, for approximately 18-20 h, at 37 °C with 5% CO₂.
7. After overnight incubation, carefully remove half of the medium (50 µL/well) by aspiration using a P200 pipette, avoiding touching the bottom of the well, and replace with fresh warm complete α-MEM containing 10% FBS, 25 ng/mL M-CSF, and 50 ng/mL RANKL for a final concentration of 25 ng/mL
   NOTE: For 1 mL of medium, add 0.25 µL of M-CSF and 0.5 µL of RANKL from a stock concentration of 100 µg/mL.
8. Change the media every 3 days and differentiate the cells into OCs for 7-14 days (Figure 1).
   ​NOTE: Keep the M-CSF and RANKL concentrations consistent throughout the culture.

Figure 1: OC differentiation workflow. Schematic overview of CD14⁺ monocyte isolation from PBMCs and differentiation into mature OCs in the presence of M-CSF and RANKL for 7-14 days. RT = room temperature. Image created with BioRender.com. Please click here to view a larger version of this figure.

5. TRAP staining for osteoclasts

1. Carefully remove the medium, fix the differentiated adherent OCs with 100 µL/well of the previously prepared fixative solution, and incubate for 1 min. Do not touch the bottom of the wells to avoid scratching the adherent cells.
   NOTE: The fixative solution is prepared as follows: 12.5 mL of citrate solution (included in the TRAP staining kit), 32.5 mL of acetone, and 5 mL of 37% formaldehyde.
2. Wash the wells three times with 300 µL of sterile distilled water. Tap the plates dry after washing.
3. Prepare a staining solution according to the manufacturer's instructions (Table of Materials); add 5 µL of Fast Garnet and 5 µL of sodium nitrite to make the Fast Garnet solution; mix by inversion, and incubate for 3 min at RT. Prepare 1 mL of staining solution by mixing 900 µL of sterile distilled water, 10 µL of naphthol, 40 µL of acetate solution, 50 µL of tartrate solution, and 10 µL of Fast Garnet solution.
4. Add 100 µL/well of freshly prepared staining solution, and incubate the plate at 37 °C in the dark for 20 min.
5. After incubation, remove the staining solution by inversion, and wash the plate three times with 300 µL/well of distilled water.
6. Remove the excess water by tapping the plates on paper towels. Leave the plates open and protected from the light to air dry overnight.
   NOTE: Dry plates can be stored for up to 6 months. Sometimes, residual buffers can promote the development of mold, which is visible under the microscope; this can be removed at any time by washing the affected wells with distilled water and letting them air dry again.
7. Take images at 10x or 20x using a brightfield microscope with a tile option to capture the entire well surface.
8. Manually count the OCs identified as TRAP⁺ purple stained cells with more than three nuclei using an image analysis software with a cell counter plugin.
   ​NOTE: The numbers of TRAP⁺ OCs per well are donor dependent and may vary from ~200-1,600 OCs/well, with an average of about 1,000 OCs/well. Moreover, to analyze the data, the OC numbers should be determined in three different wells (technical replicates), and the average should be calculated for each condition and for each biological replicate.

6. Bone resorption assay

1. Plate the freshly enriched CD14+ monocytes on 96-well osteo-assay plates coated with calcium phosphate at 1 x 10⁵ cells/well, and differentiate the OCs for 7-14 days, as indicated in steps 4.1-4.8, and changing the medium every 3 days.
   NOTE: Dentine/ivory or bovine cortical bone slices can be used in place of osteo-assay plates. If so, the total time of the culture should be prolonged to 14-21 days due to the more complex substrate to be resorbed.
2. At the end time point, carefully remove the medium, avoiding touching the bottom of the well, and lyse the cells with 10% sodium hypochlorite solution. Wash the wells three times with distilled water.
3. Scan the dry plates using a brightfield microscope, and analyze/quantify the acquired images of the resorption pits using an image analysis software.

7. Actin ring fluorescent staining

1. Plate 100 µL/well of the isolated CD14+ monocytes in an 18-well chamber slide at a cell density of 1 x 10⁵ cells/well. Differentiate the OCs in the presence of M-CSF and RANKL as described before (steps 4.1-4.8), including changing the medium every 3 days.
2. At the end time point, gently remove the medium, and wash each well twice with 200 µL/well of pre-warmed PBS, pH 7.4. Do not let the wells dry in between any of the steps.
3. Fix the sample with 100 µL/well of 4% formaldehyde solution in PBS, and incubate for 10 min at RT on an orbital shaker with gentle shaking.
   NOTE: Methanol can disrupt actin during the fixation process. Therefore, it is best to avoid any methanol-containing fixatives. The preferred fixative is methanol-free formaldehyde. The orbital shaker used in this step of the protocol was set at a power setting of 3 out of 10.
4. Wash twice with 200 µL/well of PBS, permeabilize the cells with 100 µL/well of 0.1% Triton X-100 solution diluted in PBS, and incubate for 10 min at RT on an orbital shaker with gentle shaking.
5. Wash twice with 200 µL/well of PBS. To block non-specific binding and increase the signal, add 100 µL/well of blocking solution made with 2% bovine serum albumin (BSA)/PBS solution. Incubate for 20 min at RT on an orbital shaker with gentle shaking.
6. Remove the blocking solution, and add 100 µL/well of fluorescently conjugated phalloidin solution diluted in 2% BSA/PBS solution. Incubate for 20 min at RT on an orbital shaker with gentle shaking and protected from light.
   NOTE: Adjust the concentration of the phalloidin dye according to the manufacturer's recommendations.
7. Wash twice with 200 µL/well of PBS, stain the nuclei with 100 µL/well of a solution of PBS containing 300 nM DAPI, and incubate for 10-15 min at RT on an orbital shaker with gentle shaking and protected from light.
   NOTE: DAPI is diluted in distilled water to make a 14.3 mM (5 mg/mL) DAPI stock solution. The stock solution is further diluted to the final concentration of 300 µM. Finally, the 300 µM DAPI solution is diluted one more time in PBS to a final concentration of 300 nM.
8. After 10-15 min, remove the DAPI solution, and replace it with 100 µL/well PBS.
   NOTE: Depending on the chamber slides chosen, either store with an appropriate volume of PBS (100 µL/well for 18-well chamber slides), or mount the slide with coverslips and an appropriate mounting medium. The chamber slides can be stored for up to 1 week in the fridge. For 18-well chamber slides, use a 50-100 µL staining volume and 200-300 µL for washing. Scale up for the other chamber slide sizes accordingly. To avoid evaporation, keep the coverslips inside a covered container during the incubation times. The use of an orbital shaker is recommended but not essential.
9. Visualize the staining using appropriate immunofluorescence or confocal microscopes and magnifications between 4x to 40x.

8. Enrichment of mature OCs and OC precursors via flow cytometry sorting

1. Resuspend the freshly enriched CD14⁺ monocytes at 1 x 10⁶ cells/mL, and differentiate them into mature OCs in the presence of M-CSF and RANKL in the same way as described above (steps 4.1-4.8).
   NOTE: When scaling up from a 96-well plate to a bigger plate size, follow Table 1; these volumes are calculated starting from a 1 x 10⁶ cells/mL solution and provide an optimum density for cell-cell fusion.
2. On day 7, wash the wells once with warm PBS, and add 50 µL to 1 mL (volume determined by the plate size used) of accutase. Incubate the cells at 37 ˚C with 5% CO₂ for 20 min.
3. After incubation, check the plates under a light microscope to see if the cells have detached. Further, detach the cells by tapping the plates on all sides and pipetting them up and down.
4. Collect the cell suspension in a 15 mL conical tube. Wash the wells with warm PBS (no Ca²⁺, no Mg²⁺), and combine them with the cell suspension. Repeat steps 8.2-8.3 once or twice until most of the cells have detached.
   NOTE: Accutase, the recommended method prior to surface staining for flow cytometric analysis, does not detach very large OCs. The recovery rate is ~50%-70%.
5. Centrifuge the cells at 300 x g for 5 min, resuspend the cell pellet in 1 mL of PBS, and count the cells by trypan blue exclusion.
6. Resuspend the cells at 1 x 10⁶ cells/mL, remove 100 µL corresponding to 1 x 10⁵ cells, and transfer those cells into a new polypropylene test tube. Add 200 µL of the cell sorting buffer, and set this aside on ice as the unstained control.
7. Stain the remainder of the cells with live/dead dye diluted at 1:750 for 10 min at RT and protected from light.
   NOTE: Live/dead staining needs to be performed in the absence of FBS to avoid high background staining.
8. Top up the 15 mL collection tube containing the live/dead stained cell suspension with warm cell sorting buffer (1x PBS, no Ca²⁺, no Mg²⁺, 1% FBS, and 5 mM EDTA), and centrifuge at 300 x g for 5 min to pellet the cells.
   NOTE: A high concentration of EDTA and a low concentration of FBS are recommended in the sorting buffer to prevent cell clumps.
9. Remove a volume corresponding to 1 x 10⁵ cells, and transfer them into a new polypropylene test tube for the OSCAR isotype control. Transfer all the remaining cells into another polypropylene test tube for staining and cell sorting.
   NOTE: Polypropylene test tubes are used for sorting as the cells are less likely to adhere to these tubes than to polystyrene tubes.
10. Spin the tubes at 400 x g for 5 min to pellet the cells, and discard the excess supernatant by inversion.
11. Resuspend the cell pellet in an antibody master mix solution prepared following Table 2. Stain the OSCAR isotype control tube with CD14 antibody and OSCAR isotype control in place of the OSCAR antibody.
12. Incubate the cells at 4 ˚C protected from the light for 30 min.
13. After 30 min, wash the cells by adding five volumes of the cell sorting buffer, and centrifuge at 400 x g for 5 min at 4 ˚C.
14. Resuspend the cells in 300-1,000 µL of cold cell sorting buffer, and acquire the cells using a flow cytometry sorting machine fitted with a 100μM nozzle.
    NOTE: OCs are very sticky cells, so it is important to filter them through a sterile 70 µm membrane prior to sorting.
15. Gate the OCs and pre-OCs as CD14⁻OSCAR⁺. Set the OSCAR⁺ gate based on the OSCAR isotype control tube.
16. Collect the sorted cells in polypropylene test tubes containing complete α-MEM supplemented with 20% FBS at 8 ˚C.
17. After sorting, pellet the cells by centrifugation at 300 x g for 5 min at RT, count the cells, and resuspend for downstream applications.
    ​NOTE: Usually, to get ~1 x 10⁵ sorted pre-OCs/OCs, start from ~10 x 10⁶ cells plated at day 0. The low recovery rate is influenced by cell loss during detachment with accutase and by the processing for staining and sorting. It is recommended to carry out the whole procedure using sterile buffers and reagents, and work under sterile conditions.

9. ATP assay for mitochondrial activity

1. Incubate the enriched CD14⁺ monocytes in the presence of M-CSF and RANKL in a 96-well plate in the same way as described before (steps 4.1-4.8). Plate four extra conditions in triplicates to use as controls.
2. Conduct the ATP assay with the luminescence ATP detection assay kit according to the manufacturer's manual. Briefly, to prepare the ATP solution, add 10 mL of the substrate buffer solution to the lyophilized substrate, and leave it to incubate at RT for 30 min.
   NOTE: Different methods can be used to measure the intracellular ATP production. Herein, we have used the detection of ATP production by luminescence.
3. During the incubation, prepare and add the controls directly to the control wells as follows: 2-Deoxy-D-glucose (2DG) at 10 mM and 100 mM, oligomycin at 1 µM, and 100 mM 2DG in combination with 1 µM oligomycin. Incubate for 30 min at 37 °C with 5% CO₂.
   NOTE: The 2DG blocks glycolysis, while oligomycin is an inhibitor of oxidative phosphorylation. Combining these two inhibitors results in the complete loss of ATP production via glycolysis and oxidative phosphorylation, thus meaning they serve as an internal control for the ATP assay. For 10 mM and 100 mM 2DG controls, add 0.5 µL and 5 µL of 2M 2DG stock solution per 100 µL culture well, respectively. For 1 µM oligomycin, dilute the 5 mM stock solution 1:100 in medium, and add 2 µL/well to the dedicated control wells. For the last control, add 5 µL of 2DG and 2 µL of diluted oligomycin solution per well.
4. Add 50 µL of the ATP solution to each well to stop the reaction, and incubate at RT on a shaker at 700 rpm for 5-10 min protected from light.
5. Transfer 100 µL of the supernatant to a 96-well white-bottom plate specific for the ATP assay, and read the plate using a luminescence reader.