The authors would like to thank the members of the Giachelli lab for their technical help and support. We thank the W. M. Keck Microscopy Center and the Keck Center manager, Dr. Nathanial Peters, for assistance in obtaining the confocal microscopy and widefield microscopy images. We also thank the UW Flow Core Facility and the Flow Core Facility manager, Aurelio Silvestroni, for technical support and assistance. Finally, we thank Hannah Bl&#252;mke for the support with illustration and graphic design.
Funding was provided through the National Institutes of Health grant R35 HL139602-01. We also acknowledge NIH S10 grant S10 OD016240 for instrument funding at the W. M. Keck Center as well as NIH grant 1S10OD024979-01A1 for instrument funding at the UW Flow Core Facility.

NOTE: All reagents used in this protocol can be found in the Table of Materials. Unless otherwise specified, all media were pre-equilibrated to 37 &#176;C before use. All centrifugation steps are performed at 37 &#176;C and by using the slowest acceleration/deceleration mode. Unless otherwise specified, supernatant is always removed using disposable Pasteur glass pipettes.
1. Thawing and propagation of human iPSCs
<ol>
	<li>One day prior to thawing iPSCs, coat a well of a 6-well plate with 1 mL of a basal membrane extract at a concentration of 200-300 &#181;g/mL. Place the well plate at 4 &#176;C overnight.</li>
	<li>The next day, thaw and transfer the cells into a 15 mL tube using a P1000 pipette. Dropwise, add 5-7 mL of DMEM/F-12 with 15 mM HEPES.</li>
	<li>Centrifuge the cells at 300 x g for 5 min. Gently remove the cells from the centrifuge, and be sure not to disturb the cell pellet.</li>
	<li>Carefully remove the supernatant with a Pasteur glass pipette and resuspend the cells in 1 mL of hiPSC-serum free medium (hiPSC-SFM containing 10 &#181;M of Rho kinase (ROCK) inhibitor Y-27632) using P1000 with wide bore tips.</li>
	<li>Aspirate the basal membrane extract from the well coated the previous day (step 1.1) and transfer 1 mL of hiPSC-SFM with 10 &#181;M Y-27632 into the well. Add resuspended iPSCs to the well to reach a final volume of 2 mL per well.</li>
	<li>Swirl the plate to evenly distribute iPSC aggregates before placing in an incubator at 37 &#176;C and 5% CO2.</li>
	<li>Perform full medium changes every other day using hiPSC-SFM (without the addition of ROCK inhibitor Y-27632). iPSCs usually reach 70-80% confluency after 3-4 days of propagation.</li>
</ol>
2. Passaging iPSCs
<ol>
	<li>One day prior to passaging iPSCs, coat wells of a 6-well plate with a basal membrane extract at a concentration of 200-300 &#181;g/mL. Place the well plate at 4 &#176;C for overnight. 
	NOTE: Confirm that cells have reached approximately 70-80% confluency. Ensure iPSCs do not get overconfluent (more than 80% confluency), as this will promote spontaneous differentiation.</li>
	<li>Begin passaging iPSCs by removing differentiated regions or regions with many dead iPSC aggregates under the stereomicroscope by using 10 &#181;L or 20 &#181;L pipette tips or a cell scraper. Differentiated regions will appear denser or whiter in color. 
	NOTE: Scraped-off cell aggregates may still partially attach to the well bottom.</li>
	<li>Wash the well bottom several times with a wide bore P1000 pipette to remove any aggregates that may still be partially attached to the well&#39;s bottom.</li>
	<li>Discard the spent medium in which the iPSCs have been cultivated that contains the detached iPSC aggregates. Repeat the washing step two more times with phosphate buffered saline (PBS).</li>
	<li>Next, add 1 mL of 5 U/mL Dispase per well. Edges of the iPSC colonies will lift off from the well plate, which can be observed under the stereomicroscope after 3-5 min incubation at room temperature.</li>
	<li>Carefully remove Dispase to avoid dislodging slightly detached aggregates and add 1 mL of DMEM/F-12 with 15 mM HEPES. Use a disposable cell lifter to slice the aggregates into small sizes. The consistency of iPSC aggregate sizes can be improved with a stem cell passaging tool.</li>
	<li>Wash off the sliced iPSC aggregates with the medium in the well and transfer the medium with aggregates into a 15 mL conical tube using a 5 mL serological pipette or P1000 with a wide bore tip.</li>
	<li>Rinse the well with DMEM/F-12 and transfer the medium to the 15 mL tube with the iPSC aggregates.</li>
	<li>Centrifuge the sliced iPSC aggregates at 200 x g for 3 min. Remove the supernatant with a Pasteur glass pipette and add 2 mL of hiPSC-SFM to dislodge and resuspend the iPSCs using a 5 mL serological pipette or P1000 with wide bore tips.</li>
	<li>Aspirate the leftover basal membrane extract from the pre-coated well plate(s) and add 1 mL of hiPSC-SFM to each well of the 6-well plate.</li>
	<li>Transfer iPSCs into new wells of a 6-well plate so that the final volume is 2 mL of hiPSC-SFM per well using a P1000 with wide bore tips. Depending on the iPSC line, split ratios need to be optimized. Here, a 1:6 split ratio was used.</li>
	<li>Check the well for floating aggregates and aggregate size under the stereomicroscope. Ideally, the aggregate size should be between 50-200 &#181;m.</li>
	<li>Swirl the well plate to distribute the cells evenly across the plate after the aggregates have been transferred and incubate at 37 &#176;C at 5% CO2 until 70-80% confluency.</li>
</ol>
3. Freezing back iPSCs
<ol>
	<li>To freeze iPSC cells back, passage cells as described above (see protocol step &#34;2. Passaging of iPSCs&#34;). After cells have been sliced into colonies and washed off the well bottom (step 2.10), spin down the sliced aggregates at 200 x g for 3 min. Aspirate the supernatant.</li>
	<li>Add 1 mL of serum-free cryopreservation medium per well of a 6-well plate to the 15 mL tube and resuspend the sliced aggregates using a P1000 with a wide bore tip.</li>
	<li>Transfer cells into prelabeled cryotubes. Close tubes and transfer tubes into a prechilled cryopreservation container at 4 &#176;C. Store cells at -80 &#176;C for 24-48 h.</li>
	<li>Transfer cryovials into liquid nitrogen for long-term storage. 
	NOTE: A schematic summary of the OC differentiation process is illustrated in Figure 1.</li>
</ol>
4. Embryoid body induction
<ol>
	<li>Cultivate and expand sufficient iPSCs as described above for EB induction. 
	NOTE: OC production can be upscaled by increasing the number of embryoid bodies, which in turn produce an overall higher yield of hematopoietic cells. One well of 70-80% confluent iPSCs yields approximately 8.4 x 105 cells per well of a 6-well plate well. 12,500 single-cell iPSCs are needed to form one embryoid body.</li>
	<li>Aspirate the spent medium from the iPSC cultures and rinse the iPSC colonies with D-PBS.</li>
	<li>Add 0.5 mL of prewarmed to room temperature single-cell dissociation reagent to each well of a 6-well plate and swirl the culture vessel to coat the entire well surface. Incubate the culture vessel at 37 &#176;C for 5-8 min.</li>
	<li>Remove the vessel from the incubator, aspirate the single cell dissociation reagent, and add 1 mL of the Stage 1 differentiation medium to the wells, consisting of hiPSC-serum free medium with 50 ng/mL human bone morphogenetic protein 4 (hBMP4), 50 ng/mL human vascular endothelial growth factor-165 (hVEGF), 20 ng/mL human stem cell factor (hSCF), and 10 &#181;M Y-27632.</li>
	<li>Gently detach cells by rinsing the well with the Stage 1 medium. Pool dissociated iPSCs into a 15 mL conical tube.</li>
	<li>Add 1 mL of Stage 1 differentiation medium to the wells and wash any remaining cells off or use a cell scraper for colonies that do not wash off easily.</li>
	<li>After transferring all cells to the tube, centrifuge at 200 &#215; g for 5 min at room temperature to form a cell pellet. Aspirate and resuspend the cells in a total of 2 mL of pre-equilibrated Stage 1 differentiation medium using a 5 mL serological pipette or a P1000 with a wide bore tip.</li>
	<li>Count cells using a hemocytometer or automated cell counting device. Add the medium to the 15 mL tube containing the single cell suspension to plate 12,500 cells per well in a round bottom ultra-low attachment 96-well plate in 100 &#181;L of the Stage 1 differentiation medium. 
	NOTE: Under the microscope, cells will appear as a single-cell suspension with cells dispersed throughout the well.</li>
	<li>Centrifuge the 96-well plates for 3 min at 100 x g. After centrifuging, cells should begin to resemble spheroids when viewed under the microscope. Place the plate in a 37 &#176;C incubator for 24 h.</li>
	<li>Change half of the medium on Day 1 and Day 2 with the Stage 1 differentiation medium. To improve efficiency, use a multichannel pipette to dispose of 50 &#181;L of the spent Stage 1 differentiation medium into a Petri dish.</li>
	<li>After disposing of the medium from the 96-well plate, check for EBs under the stereomicroscope that might have accidentally been removed. Transfer accidentally removed EBs to the 96-well plate using a P1000 pipette with wide bore tips.</li>
	<li>Add 50 &#181;L of fresh Stage 1 differentiation medium to each well of a round bottom ultra-low attachment 96-well plate using a multichannel pipette.</li>
</ol>
5. Hematopoietic differentiation
<ol>
	<li>One day prior to starting hematopoietic differentiation, coat a well of a 6-well plate with 1 mL of a basal membrane extract at a concentration of 200-300 &#181;g/mL. Place the well plate at 4 &#176;C overnight. 
	NOTE: Each well will receive 8 EBs at a later point of this protocol. Depending on the number of EBs prepared, coat wells correspondingly.</li>
	<li>Aspirate the excess basal membrane extract and prefill wells of the 6-well plate with 3 mL of the Stage 2 differentiation medium, consisting of a hematopoietic basal medium to which 2 mM Ultraglutamine, 55 &#181;M 2-mercaptoethanol, 25 ng/mL human interleukin 3 (hIL-3), and 100 ng/mL human macrophage colony-stimulating factor (hM-CSF) is added.</li>
	<li>Using a P1000 with wide bore tips, transfer 8 EBs into each well of the 6-well plate. After transferring, confirm by eye or under the stereomicroscope that 8 EBs are in each well. 
	NOTE: After 1 day, the floating EBs will adhere to the well bottom. In the following 5-7 days, a floating cell population comprising hematopoietic cells should become visible. The hematopoietic differentiation period can be varied, starting with 7 days. Differentiation for 10 days showed a hematopoietic population comprised of large CD45+, CD14+ and CD11b+ subpopulations.</li>
	<li>After 5 days of treatment with the Stage 2 differentiation medium, perform a medium change by removing the spent medium and dispensing it into a 50 mL conical tube. Pipette slowly to try to remove as little floating cells as possible and keep the shear stress as low as possible. Pipetting under a stereomicroscope can help avoid accidental removal of cells.</li>
	<li>Immediately add 1 mL of fresh Stage 2 differentiation medium to the wells. In order to recover any floating hematopoietic cells that may already be present at this point in the differentiation process, do not discard the dispensed medium. Rather, spin down the tube with the spent medium at 300 x g for 5 min, and aspirate the supernatant.</li>
	<li>Add fresh Stage 2 differentiation medium to the tube and resuspend in order to detach cells that might have been transferred with the spent medium.</li>
	<li>Add 2 mL of the fresh Stage 2 differentiation medium with the recovered cells into each well, which was previously filled with 1 mL of the Stage 2 differentiation medium.</li>
	<li>On Day 10 of hematopoietic differentiation, check for the presence of large numbers of floating hematopoietic cells. Harvest by collecting them in a 50 mL tube. They can either be frozen back in 10% DMSO, 50% FBS, and 40% medium or used immediately to differentiate the cells into OCs.</li>
</ol>
6. M-CSF maturation and OC differentiation
<ol>
	<li>Seed cells at a concentration of 200,000 cells/cm2 onto tissue culture treated well plates and treat with alpha-MEM, supplemented with 10% FBS and 50 ng/mL hM-CSF.</li>
	<li>To perform functional assays or imaging, detach the M-CSF matured cells 3 days after seeding by washing with pre-warmed PBS to 37 &#176;C using a P1000 with a wide bore tip. Repeated washing steps might be needed to fully detach cells.</li>
	<li>Transfer the detached cells into a 15 mL tube using a P1000 with a wide bore tip and centrifuge at 300 x g for 5 min. Discard the supernatant.</li>
	<li>Resuspend and dissolve the cell pellet with the OC differentiation medium, consisting of alpha-MEM supplemented with 10% FBS, 50 ng/mL hM-CSF, and 80 ng/mL hRANKL. Count the cells using a hemocytometer or automated cell counting device and reseed M-CSF matured cells at a density of 200,00-250,000 cells/cm2 in an appropriate culture ware for functional assays or imaging (i.e., bone resorption assays, coverslip slides for imaging, etc.).</li>
	<li>Differentiate with the OC differentiation medium for 7-9 days. Perform a complete well medium change with the fresh OC differentiation medium every 2-3 days. 
	NOTE: Multinucleated OCs typically appear after 5-7 days.</li>
</ol>

The authors declare no competing interests.

This protocol offers a reliable and robust method to differentiate iPSCs into OCs. Nevertheless, there are several pitfalls that can be encountered throughout the differentiation process. Human iPSC lines generated from cells of different tissue origins have successfully been differentiated using this protocol33. When freezing back iPSCs (see protocol step &#34;3. Freezing back iPSCs&#34;), one well at the point of passaging was frozen back into one cryovial. When thawing (see protocol step &#34;1. Thawing and propagation of human iPSCs&#34;), one cryovial was thawed into a single well of a 6-well plate. Different iPSC lines will behave slightly differently, and proliferation rates will vary. The split rate will need to be adjusted correspondingly.
When passaging iPSCs or dissociating iPSCs into a single-cell suspension for EB induction, it is important to remove any spontaneously differentiated colonies or aggregates of dead cells to improve the effectiveness and efficiency of mesodermal and hematopoietic differentiation. This can be done under a stereomicroscope by using a cell scraper, pipette tips from a P10 or P20 pipette, or a scraping tool built from a Pasteur pipette35.
As mentioned above, this protocol involves the dissociation of iPSC colonies into a single-cell suspension for EB induction, while with other protocols, iPSCs are left as aggregates to be centrifuged for EB induction36. EB size, cell number, shape, and morphology have all been reported to influence differentiation37,38,39. Thus, we hypothesize that the dissociation into a single-cell suspension allows for better EB-to-EB uniformity in size and shape after centrifugation and, hence, more consistent results of hematopoietic cell production31.
Other methods for EB induction have been described. Such methods that use a single-cell suspension in conjunction with ROCK inhibitor to improve cell survival have been reported to be advantageous in controlling EB size and differentiation outcome31.
The round-bottom 96-well plate EB induction method described in this protocol is suitable for large-scale production of EBs and allows for upscaling of OC production. More novel methods for hematopoietic differentiation without an embryoid body induction step with the potential to facilitate the differentiation process have recently been described32. Nonetheless, these protocols have not yet been established for OC differentiation33.
In the above-mentioned protocol, we describe the medium change on Day 5 of hematopoietic differentiation. A small number of floating cells may already appear around Day 4-5 of differentiation. In order to avoid discarding any floating cells, the medium should be collected in a tube and centrifuged before being discarded. The cell pellet at the bottom of the tube must then be dislodged with the fresh medium and should be transferred back to wells of a 6-well plate. The significance of the early floating cell population in OC differentiation still needs to be determined, however.
The production of hematopoietic cells can be assessed using flow cytometry. High yields of CD45+, CD14+ and CD11b+ cells are desirable for osteoclast differentiation33. Cryopreservation of harvested floating hematopoietic cells has been reported to be challenging with generally limited recovery rates and low cell viability40,41. By cryopreserving hematopoietic cells in a cryopreservation medium consisting of 50% of a serum-free hematopoietic cell expansion medium (see Table of Materials), 40% FBS and 10% DMSO, we were able to recover cells with a cell viability of approximately 90.3% &#177; 2.62 SD (n = 7) post thawing.
Osteoclastogenesis requires the fusion of multiple mononucleated OC precursors to form multinucleated osteoclasts that are capable of mineral and bone resorption. While murine OC-precursor cell lines simply need the addition of RANKL to induce OC formation42, human precursors require additional M-CSF for cell survival and proliferation43. OSCAR has recently been discovered as an additional receptor involved in OC differentiation, even though only type I collagen has thus far been identified as a ligand. While research of OSCAR with iPSC-derived OCs is still limited, supraphysiological in vitro RANKL and M-CSF concentrations in a murine cell line seem to bypass the necessity for OSCAR activation44, activation of OSCAR in vivo appears to be a necessary costimulatory signal for osteoclastogenesis45. An additional factor that needs to be considered is the well plate surface. Chemical46&#160;and physical47 surface properties are known to influence osteoclast differentiation and can either promote or hinder successful differentiation. A certain heterogeneity within the precursor population also appears critical for successful fusion of OCs, as different fusion-related factors such as CD47 and DC-STAMP act at different stages of osteoclast fusion48.
In conclusion, this protocol enables the differentiation of human OC from iPSCs to facilitate and accelerate OC research.

Osteoclasts (OCs) are hematopoietic-derived1,2, versatile cell types that are commonly used by researchers in areas such as bone disease research3,4, cancer research5,6, tissue engineering7,8, and endoprosthesis research9,10. Nevertheless, OC differentiation can be challenging as fusion of mononuclear precursors into multinucleated OCs is necessary to create functional OCs11. Several biological factors, such as receptor activator of NF-&#954;B ligand (RANKL) and macrophage colony-stimulating factor (M-CSF), are necessary for OC differentiation. M-CSF has been reported to have a positive effect on cell proliferation, cell survival, and RANK expression12,13,14. On the other hand, RANKL binds to RANK, which activates downstream signaling cascades that induce osteoclastogenesis. Activation is mediated via TNF receptor-associated factor 6 (TRAF6), which leads to the degradation of nuclear factor of kappa light polypeptide gene enhancer in B cells inhibitor, alpha (I&#954;B-&#945;), a binding protein that binds NF-kB dimers16,17. Hence, I&#954;B-&#945; degradation releases NF-kB dimers, which then translocate into the nucleus and induce the expression of the transcription factors c-Fos and Nuclear Factor of Activated T-Cells 1 (NFATc1). This, in turn, triggers the transcription of a multitude of OC differentiation-related proteins15,18. Upregulated proteins such as DC-Stamp and Atp6v0d2 mediate cell-cell fusion of OC precursors, leading to syncytium formation19,20,21.
With respect to human primary cells, CD34+ and CD14+ PBMCs are currently the most widely used cell types for differentiation into OCs22. However, this approach is limited by the heterogeneity within the CD34+ population of harvested cells from donors23 and their limited expandability. Human iPSCs present an alternative source for OCs. As they can be propagated indefinitely24, they allow for expandability and upscaling of OC production. This allows for the differentiation of large numbers of OCs, which facilitates OC research.
Several protocols for the differentiation of iPSCs into OCs have been published25,26,27. The entire differentiation process can be divided into an iPSC propagation part, a mesodermal and hematopoietic differentiation part, and OC differentiation. Propagation of iPSCs before the differentiation process allows for the upscaling of OC production prior to differentiation. Several approaches exist regarding mesodermal and hematopoietic differentiation. Traditionally, embryoid body (EB) formation has been used to differentiate hematopoietic cells, but monolayer-based approaches represent another hematopoietic differentiation strategy that does not require EB induction. Nevertheless, monolayer-based systems appear to require further optimization, as we and others have found EB-based approaches to be more robust for the differentiation of OCs.
Here, we describe the differentiation of OCs from human iPSCs using an EB-based protocol. This protocol was adapted from R&#246;ssler et al.26 and modified to increase robustness and allow for cryopreservation during the differentiation process. First, we harvested hematopoietic cells only once after 10 days of differentiation. Hematopoietic cells were then cryopreserved to allow for more flexibility during the differentiation process. Additionally, we increased the hematopoietic cell seeding density from 1 x 105 to 2 x 105 cells/cm2 for OC differentiation. A more recent human iPSC serum-free medium (hiPSC-SFM, see Table of Materials) was used, and coating of wells was performed with 200-300 &#181;g/mL of a basal membrane extract (see Table of Materials) instead of 0.1% gelatin. Penicillin/streptomycin was not added to the media.
The protocol by R&#246;ssler et al.26 was originally adapted from an iPSC to a macrophage differentiation protocol28 that uses EB formation for hematopoietic differentiation. While EB formation has been used for an extended time by researchers for hematopoietic differentiation29,30, several methods of EB induction have been described in the literature, such as spontaneous aggregation, centrifugation in a round-bottom well plate, hanging drop culture, bioreactor culture, conical tube culture, slow turning lateral vessel, and micromold gel culture31. This protocol uses centrifugation of dissociated iPSCs in a round-bottom well plate to bring single iPSC cells into proximity to each other and to allow for sphere (EB) formation, as described hereafter.

<table><tbody><tr><td>2-Mercaptoethanol</td><td>Sigma Aldrich</td><td>M6250-10ML</td><td></td></tr><tr><td>Antibody - Anti-Cathepsin K&amp;nbsp;</td><td>Abcam</td><td>ab19027</td><td></td></tr><tr><td>Antibody - APC-conjugated Anti-Human CD45</td><td>BD</td><td>555485</td><td></td></tr><tr><td>Antibody - APC-conjugated Mouse IgG1, &amp;kappa; Isotype Control</td><td>BD</td><td>555751</td><td></td></tr><tr><td>Antibody - BV711-conjugated Anti-Human CD14</td><td>BD</td><td>563372</td><td></td></tr><tr><td>Antibody - BV711-conjugates Mouse IgG2b, &amp;kappa; Isotype Control</td><td>BD</td><td>563125</td><td></td></tr><tr><td>Antibody - Goat Anti-Rabbit IgG H&amp;amp;L Alexa Fluor&amp;reg; 647</td><td>Abcam</td><td>ab150079</td><td></td></tr><tr><td>Antibody - PE-conjugated Anti-Human CD14</td><td>R&amp;amp;D Systems</td><td>FAB3832P-025</td><td></td></tr><tr><td>Antibody - PE-conjugated Anti-Human Integrin alpha M/CD11b</td><td>R&amp;amp;D Systems</td><td>FAB16991P-025</td><td></td></tr><tr><td>Antibody - PE-Cy7-conjugated Anti-Human CD34</td><td>BD</td><td>560710</td><td></td></tr><tr><td>Antibody - PE-Cy7-conjugated Mouse IgG1 &amp;kappa; Isotype Control</td><td>BD</td><td>557872</td><td></td></tr><tr><td>Antibody - PE/Cyanine5-conjugated Anti-Human CD11b</td><td>Biolegend</td><td>301308</td><td></td></tr><tr><td>Antibody - PE/Cyanine5-conjugated Mouse IgG1, &amp;kappa; Isotype Ctrl</td><td>Biolegend</td><td>400118</td><td></td></tr><tr><td>Antibody - PerCP-Cy5.5-conjugated Mouse IgG1 &amp;kappa; Isotype Control</td><td>BD</td><td>550795</td><td></td></tr><tr><td>Antibody - PerCpCy5.5-conjugated Anti-Human CD43</td><td>BD</td><td>563521</td><td></td></tr><tr><td>Bone Resorption Assay Kit</td><td>CosmoBioUSA</td><td>CSR-BRA-24KIT</td><td></td></tr><tr><td>Countess 3 Automated Cell Counter</td><td>ThermoFisher</td><td>16812556</td><td></td></tr><tr><td>Cultrex Stem Cell Qualified Reduced Growth Factor Basement Membrane Extract</td><td>R&amp;amp;D Sytems</td><td>3434-010-02</td><td>Basal membrane extract</td></tr><tr><td>DAPI</td><td>R&amp;amp;D Systems</td><td>5748/10</td><td></td></tr><tr><td>Dispase (5 U/mL)</td><td>STEMCELL Technologies</td><td>7913</td><td></td></tr><tr><td>DMEM/F-12 with 15 mM HEPES</td><td>Stem Cell</td><td>36254</td><td></td></tr><tr><td>DMSO</td><td>Sigma Aldrich</td><td>D2650</td><td></td></tr><tr><td>DPBS</td><td>Sigma Aldrich</td><td>D8537-500ML</td><td></td></tr><tr><td>Human Bone Morphogenetic Protein 4 (hBMP4)</td><td>STEMCELL Technologies</td><td>78211</td><td></td></tr><tr><td>Human IL-3</td><td>STEMCELL Technologies</td><td>78146.1</td><td></td></tr><tr><td>Human Macrophage Colony-stimulating Factor (hM-CSF)</td><td>STEMCELL Technologies</td><td>78150.1</td><td></td></tr><tr><td>Human Soluble Receptor Activator of Nuclear Factor-&amp;kappa;B Ligand (hsRANKL)</td><td>STEMCELL Technologies</td><td>78214.1</td><td></td></tr><tr><td>Human Stem Cell Factor (hSCF)</td><td>STEMCELL Technologies</td><td>78155.1</td><td></td></tr><tr><td>Human TruStain FcX (Fc Receptor Blocking Solution)</td><td>Biolegend</td><td>422301</td><td></td></tr><tr><td>Human Vascular Endothelial Growth Factor-165 (hVEGF165)</td><td>STEMCELL Technologies</td><td>78073</td><td></td></tr><tr><td>Invitrogen Rhodamine Phalloidin</td><td>Invitrogen</td><td>R415</td><td></td></tr><tr><td>MEM &amp;alpha;, nucleosides, no phenol red</td><td>ThermoFisher</td><td>41061029</td><td></td></tr><tr><td>mFreSR</td><td>STEMCELL Technologies</td><td>05855</td><td>Serum free cryopreservation medium</td></tr><tr><td>mTeSR Plus medium</td><td>STEMCELL Technologies</td><td>100-0276</td><td>Human iPSC-serum free medium (hiPSC-SFM)</td></tr><tr><td>Nunclon Sphera 96-Well, Nunclon Sphera-Treated, U-Shaped-Bottom Microplate</td><td>Thermo Scientific</td><td>174925</td><td>Round bottom ultra-low attachment 96-well plate</td></tr><tr><td>P1000 Wide Bore Tips</td><td>ThermoFisher</td><td>2079GPK</td><td></td></tr><tr><td>ROCK-Inhibitor Y-27632</td><td>STEMCELL Technologies</td><td>72304</td><td></td></tr><tr><td>StemSpan SFEM</td><td>StemCell</td><td>09650</td><td>Hematopoietic cell culture medium</td></tr><tr><td>TrypLE Select Enzyme (1X), no phenol red</td><td>Thermo Fisher</td><td>12563011</td><td>Single-cell dissociation reagent</td></tr><tr><td>Ultraglutamine</td><td>Bioscience Lonza</td><td>BE17-605E/U1</td><td></td></tr><tr><td>X-VIVO 15 Serum-free Hematopoietic Cell Medium</td><td>Bioscience Lonza</td><td>04-418Q</td><td>Hematopoietic basal medium</td></tr><tr><td>&amp;micro;-Slide 8 Well High</td><td>Ibidi</td><td>80806</td><td></td></tr></tbody></table>

differentiation and characterization of osteoclasts from human induced pluripotent stem cells

This protocol details the propagation and passaging of human iPSCs and their differentiation into osteoclasts. First, iPSCs are dissociated into a single-cell suspension for further use in embryoid body induction. Following mesodermal induction, embryoid bodies undergo hematopoietic differentiation, producing a floating hematopoietic cell population. Subsequently, the harvested hematopoietic cells undergo a macrophage colony-stimulating factor maturation step and, finally, osteoclast differentiation. After osteoclast differentiation, osteoclasts are characterized by staining for TRAP in conjunction with a methyl green nuclear stain. Osteoclasts are observed as multinucleated, TRAP+ polykaryons. Their identification can be further supported by Cathepsin K staining. Bone and mineral resorption assays allow for functional characterization, confirming the identity of bona fide osteoclasts. This protocol demonstrates a robust and versatile method to differentiate human osteoclasts from iPSCs and allows for easy adoption in applications requiring large quantities of functional human osteoclasts. Applications in the areas of bone research, cancer research, tissue engineering, and endoprosthesis research could be envisioned.

Monitoring cell morphology throughout the differentiation process 
All results described below were generated using the MCND-TENS2 iPSC line for OC differentiation. This iPSC line has previously been used in several studies32,33. Nevertheless, other iPSC lines have also been successfully used with this differentiation protocol.
Regular visual assessment reveals differing and distinct morphological characteristics of iPSCs throughout the differentiation process to OCs (Figure 2). iPSC colonies (Figure 2A) were dissociated into a single-cell suspension, which appears as individual cells throughout the round bottom well plate before centrifugation (Figure 2B). Following centrifugation (step 4.9 in the protocol), cells will collect in the center of the round bottom ultra-low attachment well plate and subsequently form spheres (embryoid bodies, EBs, Figure 2C). EBs will increase two to three times in size throughout the mesodermal differentiation process (Figure 2D) and become easily visible to the eye at the end of the 4-day differentiation period (Figure 2E). EBs can be seen adhering and fusing with the well plate bottom following the transfer into wells of a 6-well plate for hematopoietic differentiation (step 5.3 in the protocol, Figure 2F). After 7-8 days, large quantities of floating hematopoietic cells become visible in the culture medium (Figure 2G). After harvesting and replating hematopoietic cells, an M-CSF maturation phase follows, and OC differentiation is initiated (step 6.4 in the protocol). Within 5-6 days, multinucleated cells with a large transparent cell body first become visible (Figure 2H). A large number of mononuclear cells are still visible at this stage. After 2-3 more days of OC differentiation, OCs will further fuse with adjacent cells to form large polykarions with even more nuclei (Figure 2I).
Assessing the EB-derived hematopoietic population 
Hematopoietic differentiation can be performed for a variable length of time. Time periods starting with 7 days up to 9 weeks have been described in the literature. In this protocol, hematopoietic differentiation is performed for a period of 10 days. We found that 10 days of hematopoietic differentiation yielded low numbers of early CD34+ (0.53%, Figure 3A) and a larger number of midterm stage CD43+ (48.5%, Figure 3B) hematopoietic progenitors. More critically, sufficient quantities of CD45+ (96.2%, Figure 3C), CD14+ (33%, Figure 3D), and CD11b+ (35.9%, Figure 3E) HPCs were generated after a treatment period of 10 days to successfully differentiate them further into OCs. However, the cytokine treatment period for hematopoietic differentiation (step 5.4 in the protocol) may need to be adjusted and optimized based on the iPSC line to generate adequate amounts of CD45+, CD14+, and CD11b+ cells.
On average, 6 million hematopoietic cells were harvested after 10 days of differentiation from each well with 8 EBs.
Assessing OC morphology and activity 
Following OC differentiation, OCs can be assessed morphologically and functionally. OC precursors are reseeded onto chamber slides or coverslip slides following the M-CSF maturation step to improve image quality when staining for TRAP or Cathepsin K. Enzymatic TRAP staining following OC differentiation shows large, multinucleated, TRAP-positive OCs (Figure 4A). Additionally, slightly TRAP-positive mononuclear cells can be seen interspersed in between multinuclear OCs. Negative controls without the addition of RANKL do not display fused multinuclear OC. Nevertheless, a small number of slightly TRAP-positive mononuclear cells can be depicted (Figure 4B).
Confocal laser scanning microscopy (CLSM) images show OCs stained for Cathepsin K (turquoise) and F-actin (red) in conjunction with DAPI nuclear stain (blue) (Figure 4C, 4D). Large multinuclear OCs are visible when treated with RANKL according to the protocol, which depicts an extensive F-actin cytoskeletal structure and stain positive for Cathepsin K (Figure 4C). Negative controls without the addition of RANKL, on the other hand, do not show fused multinuclear cells.
OCs can be further assessed functionally by measuring the resorptive activity. Bone or mineral resorption assays can be used to determine the resorptive activity. Here, OC precursors were seeded onto calcium phosphate-coated wells and terminally differentiated. Large areas where the mineral coating was resorbed are visible in a bluish-gray color (Figure 4E). Resorption pits of different sizes can be identified. Not resorbed, the remaining calcium phosphate-coating is visible in brown. The presence of resorption pits confirms the identity of the differentiated multinucleated cells as OCs. Additionally, resorption pits can further be quantified in order to assess and compare the resorptive activity. The total area of resorbed mineral over the total well surface area can be quantified as a percentage measure. Additionally, the size and number of resorption pits can further be quantified. Untreated negative controls did not display resorption pits (Figure 4F).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66527/66527fig01.jpg" /> 
Figure 1: Schematic illustration of the osteoclast differentiation process from human iPSCs. Illustration drawn by Hannah Bl&#252;mke using Affinity Designer 2.1.1. The illustration utilizes previously used drawings33. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66527/66527fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66527/66527fig02.jpg" /> 
Figure 2: Microscopy images throughout the differentiation process of human iPSCs toward osteoclasts. (A) Undifferentiated iPSC colonies throughout propagation. (B) iPSCs in round bottom wells after dissociation into a single cell suspension prior to centrifugation. (C) Centrally collected single cell iPSCs following centrifugation. (D) EBs grow in size throughout the 4-day mesodermal differentiation period. (E) Visible embryoid bodies following mesodermal differentiation. (F) Following the transfer of embryoid bodies onto basal membrane extract coated 6-well plates, embryoid bodies can be seen adhering and fusing with the well bottom. (G) After 5-7 days of hematopoietic differentiation, a large number of floating hematopoietic cells can be observed in the medium. (H) Following M-CSF maturation, the first osteoclasts with 3-4 nuclei appear after 5-7 days of differentiation with RANKL. (I) At the end of osteoclast differentiation, large multinucleated cells can be observed. Scale bars: A, D, F, G = 200 &#181;m, B, C, H, I = 50 &#181;m, E = 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66527/66527fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66527/66527fig03.jpg" /> 
Figure 3: Surface marker analysis of embryoid body-derived hematopoietic cells using flow cytometry. Marker expression allows for the analysis of hematopoietic cells and identification of sub-populations after gating for singlets and live cells. (A) Ontogenetically early CD34+ hematopoietic progenitor cell population is very small to absent in conjunction with this protocol. (B) CD43+ cells make up approximately 50% of the entire population. (C) Later stage CD45+ hematopoietic progenitor cells make up the largest part of the hematopoietic population with 96.2%. (D, E) More direct CD14+ and CD11b+ OC precursors make up 33% and 36%, respectively. In red: unstained negative control, in blue: isotype controls, in yellow: cells stained with the respective marker antibody. Plots use previously published data33. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66527/66527fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66527/66527fig04.jpg" /> 
Figure 4: Morphological and functional assessment of iPSC-differentiated human osteoclasts. (A) TRAP staining following osteoclast differentiation shows large, multinucleated, TRAP positive osteoclasts. Slightly TRAP positive mononuclear cells can be seen interspersed in-between multinuclear osteoclasts. (B) Negative controls without the addition of RANKL stained for TRAP do not display fused multinuclear osteoclasts. Nevertheless, a small number of slightly TRAP positive, mononuclear cells can be depicted. (C) Confocal laser scanning microscopy images of osteoclast differentiated hematopoietic cells on a coverslip slide stained for F-actin (red) and Cathepsin K (turquoise) in conjunction with DAPI nuclear stain (blue) show large multinucleated Cathepsin K positive osteoclasts. (D) Negative controls without the addition of RANKL show similar to (B) mononuclear cells at a lower cell density. (E) Functional assessment can be performed by assessing the resorption activity of osteoclasts. For this, osteoclast differentiation is performed on a bone or mineral resorption assay. Tiled well images acquired with an inverted widefield microscope in phase contrast mode depict large resorption areas of the calcium phosphate mineral layer. Resorptive activity can be further quantified by measuring the resorption area over the total area. (F) Negative controls without the addition of RANKL do not show resorption areas. Scale bars: A, B = 100 &#181;m, C, D = 50 &#181;m, E, F = 1 mm. Images have been edited from previously published data33. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66527/66527fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Differentiation and Characterization of Osteoclasts from Human Induced Pluripotent Stem Cells at JoVE.com

Differentiation and Characterization of Osteoclasts from Human Induced Pluripotent Stem Cells

This protocol presents the differentiation of human osteoclasts from induced pluripotent stem cells (iPSCs) and describes methods for the characterization of osteoclasts and osteoclast precursors.

This protocol details the propagation and passaging of human iPSCs and their differentiation into osteoclasts. First, iPSCs are dissociated into a ...

differentiation-characterization-osteoclasts-from-human-induced

Nanyang Technological University

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Developmental Biology

1.5K Views.  University of Washington. This protocol presents the differentiation of human osteoclasts from induced pluripotent stem cells (iPSCs) and describes methods for the characterization of osteoclasts and osteoclast precursors. 

Video: Differentiation and Characterization of Osteoclasts from Human Induced Pluripotent Stem Cells

Modeling Osteosarcoma Using Li-Fraumeni Syndrome Patient-derived Induced Pluripotent Stem Cells

Li-Fraumeni syndrome (LFS) is an autosomal dominant hereditary cancer disorder. Patients with LFS are predisposed to a various type of tumors, including osteosarcoma--one of the most frequent primary non-hematologic malignancies in the childhood and adolescence. Therefore, LFS provides an ideal model to study this malignancy. Taking advantage of iPSC methodologies, LFS-associated osteosarcoma can be successfully modeled by differentiating LFS patient iPSCs to mesenchymal stem cells (MSCs), and then to osteoblasts--the cells of origin for osteosarcomas. These LFS osteoblasts recapitulate oncogenic properties of osteosarcoma, providing an attractive model system for delineating the pathogenesis of osteosarcoma. This manuscript demonstrates a protocol for the generation of iPSCs from LFS patient fibroblasts, differentiation of iPSCs to MSCs, differentiation of MSCs to osteoblasts, and in vivo tumorigenesis using LFS osteoblasts. This iPSC disease model can be extended to identify potential biomarkers or therapeutic targets for LFS-associated osteosarcoma.

Here, we present a protocol for the generation of induced pluripotent Stem Cells (iPSCs) from Li-Fraumeni Syndrome (LFS) patient derived fibroblasts, differentiation of iPSCs via mesenchymal stem cells (MSCs) to osteoblasts, and modeling in vivo tumorigenesis using LFS patient-derived osteoblasts.

Li-Fraumeni syndrome (LFS) is an autosomal dominant hereditary cancer disorder. Patients with LFS are predisposed to a various type of tumors, including ...

Cancer Research

Stimulation of Notch Signaling in Mouse Osteoclast Precursors

Notch signaling is a key component of multiple physiological and pathological processes. The nature of Notch signaling, however, makes in vitro investigation of its varying and sometimes contradictory roles a challenge. As a component of direct cell-cell communication with both receptors and ligands bound to the plasma membrane, Notch signaling cannot be activated in vitro by simple addition of ligands to culture media, as is possible with many other signaling pathways. Instead, Notch ligands must be presented to cells in an immobilized state. 
Variations in methods of Notch signaling activation can lead to different outcomes in cultured cells. In osteoclast precursors, in particular, differences in methods of Notch stimulation and osteoclast precursor culture and differentiation have led to disagreement over whether Notch signaling is a positive or negative regulator of osteoclast differentiation. While closer comparisons of osteoclast differentiation under different Notch stimulation conditions in vitro and genetic models have largely resolved the controversy regarding Notch signaling and osteoclasts, standardized methods of continuous and temporary stimulation of Notch signaling in cultured cells could prevent such discrepancies in the future.
This protocol describes two methods for stimulating Notch signaling specifically in cultured mouse osteoclast precursors, though these methods should be applicable to any adherent cell type with minor adjustments. The first method produces continuous stimulation of Notch signaling and involves immobilizing Notch ligand to a tissue culture surface prior to the seeding of cells. The second, which uses Notch ligand bound to agarose beads allows for temporary stimulation of Notch signaling in cells that are already adhered to a culture surface. This protocol also includes methods for detecting Notch activation in osteoclast precursors as well as representative transcriptional markers of Notch signaling activation.

Notch signaling is a form of cellular communication that relies upon direct contact between cells. To properly induce Notch signaling in vitro, Notch ligands must be presented to cells in an immobilized state. This protocol describes methods for in vitro stimulation of Notch signaling in mouse osteoclast precursors.

Notch signaling is a key component of multiple physiological and pathological processes. The nature of Notch signaling, however, makes in vitro ...

Differentiating Chondrocytes from Peripheral Blood-derived Human Induced Pluripotent Stem Cells

In this study, we used peripheral blood cells (PBCs) as seed cells to produce chondrocytes via induced pluripotent stem cells (iPSCs) in an integration-free method. Following embryoid body (EB) formation and fibroblastic cell expansion, the iPSCs are induced for chondrogenic differentiation for 21 days under serum-free and xeno-free conditions. After chondrocyte induction, the phenotypes of the cells are evaluated by morphological, immunohistochemical, and biochemical analyses, as well as by the quantitative real-time PCR examination of chondrogenic differentiation markers. The chondrogenic pellets show positive alcian blue and toluidine blue staining. The immunohistochemistry of collagen II and X staining is also positive. The sulfated glycosaminoglycan (sGAG) content and the chondrogenic differentiation markers COLLAGEN 2 (COL2), COLLAGEN 10 (COL10), SOX9, and AGGRECAN are significantly upregulated in chondrogenic pellets compared to hiPSCs and fibroblastic cells. These results suggest that PBCs can be used as seed cells to generate iPSCs for cartilage repair, which is patient-specific and cost-effective.

We present a protocol to generate a chondrogenic lineage from human peripheral blood (PB) via induced pluripotent stem cells (iPSCs) using an integration-free method, which includes embryoid body (EB) formation, fibroblastic cells expansion, and chondrogenic induction.

In this study, we used peripheral blood cells (PBCs) as seed cells to produce chondrocytes via induced pluripotent stem cells (iPSCs) in an ...

Isolation, Culture, and Differentiation of Bone Marrow Stromal Cells and Osteoclast Progenitors from Mice

Bone marrow stromal cells (BMSCs) constitute a cell population routinely used as a representation of mesenchymal stem cells in vitro. They reside within the bone marrow cavity alongside hematopoietic stem cells (HSCs), which can give rise to red blood cells, immune progenitors, and osteoclasts. Thus, extractions of cell populations from the bone marrow results in a very heterogeneous mix of various cell populations, which can present challenges in experimental design and confound data interpretation. Several isolation and culture techniques have been developed in laboratories in order to obtain more or less homogeneous populations of BMSCs and HSCs invitro. Here, we present two methods for isolation of BMSCs and HSCs from mouse long bones: one method that yields a mixed population of BMSCs and HSCs and one method that attempts to separate the two cell populations based on adherence. Both methods provide cells suitable for osteogenic and adipogenic differentiation experiments as well as functional assays.

In this article, we present methods to isolate and differentiate bone marrow stromal cells and hematopoietic stem cells from mouse long bones. Two different protocols are presented yielding different cell populations suitable for expansion and differentiation into osteoblasts, adipocytes, and osteoclasts.

Bone marrow stromal cells (BMSCs) constitute a cell population routinely used as a representation of mesenchymal stem cells in vitro. They reside within ...

Differentiation of Newborn Mouse Skin Derived Stem Cells into Germ-like Cells In vitro

Studying germ cell formation and differentiation has traditionally been very difficult due to low cell numbers and their location deep within developing embryos. The availability of a "closed" in vitro based system could prove invaluable for our understanding of gametogenesis. The formation of oocyte-like cells (OLCs) from somatic stem cells, isolated from newborn mouse skin, has been demonstrated and can be visualized in this video protocol. The resulting OLCs express various markers consistent with oocytes such as Oct4 , Vasa , Bmp15, and Scp3. However, they remain unable to undergo maturation or fertilization due to a failure to complete meiosis. This protocol will provide a system that is useful for studying the early stage formation and differentiation of germ cells into more mature gametes. During early differentiation the number of cells expressing Oct4 (potential germ-like cells) reaches ~5%, however currently the formation of OLCs remains relatively inefficient. The protocol is relatively straight forward though special care should be taken to ensure the starting cell population is healthy and at an early passage.

Differentiation of Newborn Mouse Skin Derived Stem Cells into Germ-like Cells In vitro

In recent years the formation of germ cells using in vitro culture methods has been demonstrated using several different types of somatic stem cells. This article will visually present the differentiation of newborn mouse derived stem cells to early oocyte-like cells.

Studying  germ cell formation and differentiation has traditionally been very difficult  due to low cell numbers and their location deep within developing ...

Biology

Differentiation of Functional Osteoclasts from Human Peripheral Blood CD14+ Monocytes

Osteoclasts (OCs) are bone-resorbing cells that play a pivotal role in skeletal development and adult bone remodeling. Several bone disorders are caused by increased differentiation and activation of OCs, so the inhibition of this pathobiology is a key therapeutic principle.Two key factors drive the differentiation of OCs from myeloid precursors: macrophage colony-stimulating factor (M-CSF) and receptor activator of nuclear factor kappa-B ligand (RANKL). Human circulating CD14+ monocytes have long been known to differentiate into OCs in vitro. However, the exposure time and the concentration of RANKL influence the differentiation efficiency. Indeed, protocols for the generation of human OCs in vitro have been described, but they often result in a poor and lengthy differentiation process. Herein, a robust and standardized protocol for generating functionally active mature human OCs in a timely manner is provided. CD14+ monocytes are enriched from human peripheral blood mononuclear cells (PBMCs) and primed with M-CSF to upregulate RANK. Subsequent exposure to RANKL generates OCs in a dose- and time-dependent manner. OCs are identified and quantified by staining with tartrate acid-resistant phosphatase (TRAP) and light microscopy analysis. Immunofluorescence staining of nuclei and F-actin is used to identify functionally active OCs. In addition, OSCAR+CD14&#8722; mature OCs are further enriched via flow cytometry cell sorting, and OC functionality quantified by mineral (or dentine/bone) resorption assays and actin ring formation. Finally, a known OC inhibitor, rotenone, is used on mature OCs, demonstrating that adenosine triphosphate (ATP) production is essential for actin ring integrity and OC function. In conclusion, a robust assay for differentiating high numbers of OCs is established in this work, which in combination with actin ring staining and an ATP assay&#160;provides a useful in vitro model to evaluate OC function and to screen for novel therapeutic compounds that can modulate the differentiation process.

Differentiation of Functional Osteoclasts from Human Peripheral Blood CD14+ Monocytes

Osteoclasts are key bone-resorbing cells in the body. This protocol describes a reliable method for the in vitro differentiation of osteoclasts from human peripheral blood monocytes. This method can be used as an important tool to further understand osteoclast biology in homeostasis and in diseases.

Osteoclasts (OCs) are bone-resorbing cells that play a pivotal role in skeletal development and adult bone remodeling. Several bone disorders are caused ...

Immunology and Infection

Osteoclast Derivation from Mouse Bone Marrow

Osteoclasts are highly specialized cells that are derived from the monocyte/macrophage lineage of the bone marrow. Their unique ability to resorb both the organic and inorganic matrices of bone means that they play a key role in regulating skeletal remodeling. Together, osteoblasts and osteoclasts are responsible for the dynamic coupling process that involves both bone resorption and bone formation acting together to maintain the normal skeleton during health and disease.
As the principal bone-resorbing cell in the body, changes in osteoclast differentiation or function can result in profound effects in the body. Diseases associated with altered osteoclast function can range in severity from lethal neonatal disease due to failure to form a marrow space for hematopoiesis, to more commonly observed pathologies such as osteoporosis, in which excessive osteoclastic bone resorption predisposes to fracture formation.
An ability to isolate osteoclasts in high numbers in vitro has allowed for significant advances in the understanding of the bone remodeling cycle and has paved the way for the discovery of novel therapeutic strategies that combat these diseases. 
Here, we describe a protocol to isolate and cultivate osteoclasts from mouse bone marrow that will yield large numbers of osteoclasts.

Osteoclasts are the principal bone-resorbing cell in the body. An ability to isolate osteoclasts in large numbers has resulted in significant advances in the understanding of osteoclast biology. In this protocol, we describe a method for isolation, cultivating and quantifying osteoclast activity in vitro.

Osteoclasts are highly specialized cells that are derived from the monocyte/macrophage lineage of the bone marrow. Their unique ability to resorb both the ...

Isolation, Purification, and Differentiation of Osteoclast Precursors from Rat Bone Marrow

Osteoclasts are large, multinucleated, and bone-resorbing cells of the monocyte-macrophage lineage that are formed by the fusion of monocytes or macrophage precursors. Excessive bone resorption is one the most significant cellular mechanisms leading to osteolytic diseases, including osteoporosis, periodontitis, and periprosthetic osteolysis. The main physiological function of osteoclasts is to absorb both the hydroxyapatite mineral component and the organic matrix of bone, generating the characteristic resorption appearance on the surface of bones. There are relatively few osteoclasts compared to other cells in the body, especially in adult bones. Recent studies have focused on how to obtain more mature osteoclasts in less time, which has always been a problem. Several improvements in the isolation and culture techniques have developed in laboratories in order to obtain more mature osteoclasts. Here, we introduce a method that isolates bone marrow in less time and with less effort compared to the traditional procedure, using a special and simple device. With the use of density gradient centrifugation, we obtain large amounts of fully differentiated osteoclasts from rat bone marrow, which are identified by classical methods.

Osteoclasts are tissue-specific macrophage polykaryons derived from the monocyte-macrophage lineage of hematopoietic stem cells. This protocol describes how to isolate bone marrow cells so that large quantities of osteoclasts are obtained while reducing the risk of accidents found in traditional methods.

Osteoclasts are large, multinucleated, and bone-resorbing cells of the monocyte-macrophage lineage that are formed by the fusion of monocytes or ...

Characterizing Surface Antigens and Proliferation Ability in hiPSC-Driving MSCs

Comparison of Two Representative Methods for Differentiation of Human Induced Pluripotent Stem Cells into Mesenchymal Stromal Cells

Mesenchymal stromal cells (MSCs) are adult pluripotent stem cells which have been widely used in regenerative medicine. As somatic tissue-derived MSCs are restricted by limited donation, quality variations, and biosafety, the past 10 years have seen a great rise in efforts to generate MSCs from human induced pluripotent stem cells (hiPSCs). Past and recent efforts in the differentiation of hiPSCs into MSCs have been centered around two culture methodologies: (1) the formation of embryoid bodies (EBs) and (2) the use of monolayer culture. This protocol describes these two representative methods in deriving MSC from hiPSCs. Each method presents its advantages and disadvantages, including time, cost, cell proliferation ability, the expression of MSC markers, and their capability of differentiation in vitro. This protocol demonstrates that both methods can derive mature and functional MSCs from hiPSCs. The monolayer method is characterized by lower cost, simpler operation, and easier osteogenic differentiation, while the EB method is characterized by lower time consumption.

Author Spotlight: Advancements in iPSCs and Genetic Disease Research 

This protocol describes and compares two representative methods for differentiating hiPSCs into mesenchymal stromal cells (MSCs). The monolayer method is characterized by lower cost, simpler operation, and easier osteogenic differentiation. The embryoid bodies (EBs) method is characterized by lower time consumption.

Mesenchymal stromal cells (MSCs) are adult pluripotent stem cells which have been widely used in regenerative medicine. As somatic tissue-derived MSCs are ...

Monitoring Pluripotent Stem Cell Differentiation by Image-Based ML Strategy

A Live-cell Image-Based Machine Learning Strategy to Monitor Pluripotent Stem Cell Differentiation

Pluripotent stem cell (PSC) technologies have been widely used in drug discovery, disease modeling, and regenerative medicine.&#160;However, available PSC-to-functional cell differentiation systems are impeded by problems of severe line-to-line and batch-to-batch variability. Precise control of cell differentiation in real time is therefore important. In this protocol, we describe a non-invasive and intelligent strategy that overcomes the variability in cell differentiation by using bright-field image-based machine learning. Taking PSC-to-cardiomyocyte differentiation as an example, this methodology provides detailed information for control of the initial PSC state, early assessment and intervention in differentiation conditions, and elimination of the misdifferentiated cell contamination, together realizing consistently high-quality differentiation from PSCs to functional cells. In principle, this strategy can be extended to other cell differentiation or reprogramming systems with multiple steps to support cell manufacturing, as well as to further our understanding of the mechanisms during cell fate conversion.

Author Spotlight: Enhancing PSC-to-Functional Cell Differentiation Using ML Models Based on Live-Cell Bright-Field Imaging

Available pluripotent stem cell (PSC)-to-functional cell differentiation systems are currently impeded by problems of severe line-to-line and batch-to-batch variability. Here, using cardiac differentiation as the main example, we present a protocol to intelligently monitor and modulate the process of PSC differentiation based on image-based machine learning.

Pluripotent stem cell (PSC) technologies have been widely used in drug discovery, disease modeling, and regenerative medicine.&#160;However, available ...

Differentiation and Characterization of Osteoclasts from Human Induced Pluripotent Stem Cells

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Modeling Osteosarcoma Using Li-Fraumeni Syndrome Patient-derived Induced Pluripotent Stem Cells

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