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...

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>2-Mercaptoethanol</td>
			<td>Sigma Aldrich</td>
			<td>M6250-10ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Antibody - Anti-Cathepsin K&#160;</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, &#954; 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, &#954; Isotype Control</td>
			<td>BD</td>
			<td>563125</td>
			<td></td>
		</tr>
		<tr>
			<td>Antibody - Goat Anti-Rabbit IgG H&#38;L Alexa Fluor&#174; 647</td>
			<td>Abcam</td>
			<td>ab150079</td>
			<td></td>
		</tr>
		<tr>
			<td>Antibody - PE-conjugated Anti-Human CD14</td>
			<td>R&#38;D Systems</td>
			<td>FAB3832P-025</td>
			<td></td>
		</tr>
		<tr>
			<td>Antibody - PE-conjugated Anti-Human Integrin alpha M/CD11b</td>
			<td>R&#38;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 &#954; 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, &#954; Isotype Ctrl</td>
			<td>Biolegend</td>
			<td>400118</td>
			<td></td>
		</tr>
		<tr>
			<td>Antibody - PerCP-Cy5.5-conjugated Mouse IgG1 &#954; 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&#38;D Sytems</td>
			<td>3434-010-02</td>
			<td>Basal membrane extract</td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>R&#38;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-&#954;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 &#945;, 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>&#181;-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 ...

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

Research

JoVE Journal

Developmental Biology

1.4K 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

Derivation of Highly Purified Cardiomyocytes from Human Induced Pluripotent Stem Cells Using Small Molecule-modulated Differentiation and Subsequent Glucose Starvation

Human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) have become an important cell source to address the lack of primary cardiomyocytes available for basic research and translational applications. To differentiate hiPSCs into cardiomyocytes, various protocols including embryoid body (EB)-based differentiation and growth factor induction have been developed. However, these protocols are inefficient and highly variable in their ability to generate purified cardiomyocytes. Recently, a small molecule-based protocol utilizing modulation of Wnt/&#946;-Catenin signaling was shown to promote cardiac differentiation with high efficiency. With this protocol, greater than 50%-60% of differentiated cells were cardiac troponin-positive cardiomyocytes were consistently observed. To further increase cardiomyocyte purity, the differentiated cells were subjected to glucose starvation to specifically eliminate non-cardiomyocytes based on the metabolic differences between cardiomyocytes and non-cardiomyocytes. Using this selection strategy, we consistently obtained a greater than 30% increase in the ratio of cardiomyocytes to non-cardiomyocytes in a population of differentiated cells. These highly purified cardiomyocytes should enhance the reliability of results from human iPSC-based in vitro disease modeling studies and drug screening assays.

Here, we describe a robust protocol for human cardiomyocyte derivation that combines small molecule-modulated cardiac differentiation and glucose deprivation-mediated cardiomyocyte purification, enabling production of purified cardiomyocytes for the purposes of cardiovascular disease modeling and drug screening.

Human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) have become an important cell source to address the lack of primary cardiomyocytes ...

Three-dimensional Quantification of Dendritic Spines from Pyramidal Neurons Derived from Human Induced Pluripotent Stem Cells

Dendritic spines are small protrusions that correspond to the post-synaptic compartments of excitatory synapses in the central nervous system. They are distributed along the dendrites. Their morphology is largely dependent on neuronal activity, and they are dynamic. Dendritic spines express glutamatergic receptors (AMPA and NMDA receptors) on their surface and at the levels of postsynaptic densities. Each spine allows the neuron to control its state and local activity independently. Spine morphologies have been extensively studied in glutamatergic pyramidal cells of the brain cortex, using both in vivo approaches and neuronal cultures obtained from rodent tissues. Neuropathological conditions can be associated to altered spine induction and maturation, as shown in rodent cultured neurons and one-dimensional quantitative analysis 1. The present study describes a protocol for the 3D quantitative analysis of spine morphologies using human cortical neurons derived from neural stem cells (late cortical progenitors). These cells were initially obtained from induced&#160;pluripotent stem cells. This protocol allows the analysis of spine morphologies at different culture periods, and with possible comparison between induced&#160;pluripotent stem cells obtained from control individuals with those obtained from patients with psychiatric diseases.

Dendritic spines of pyramidal neurons are the sites of most excitatory synapses in mammalian brain cortex. This method describes a 3D quantitative analysis of spine morphologies in human cortical pyramidal glutamatergic neurons derived from induced&#160;pluripotent stem cells.

Dendritic spines are small protrusions that correspond to the post-synaptic compartments of excitatory synapses in the central nervous system. They are ...

Large-Scale Production of Cardiomyocytes from Human Pluripotent Stem Cells Using a Highly Reproducible Small Molecule-Based Differentiation Protocol

Maximizing the benefit of human pluripotent stem cells (hPSCs) for research, disease modeling, pharmaceutical and clinical applications requires robust methods for the large-scale production of functional cell types, including cardiomyocytes. Here we demonstrate that the temporal manipulation of WNT, TGF-&#946;, and SHH signaling pathways leads to highly efficient cardiomyocyte differentiation of single-cell passaged hPSC lines in both static suspension and stirred suspension bioreactor systems. Employing this strategy resulted in ~ 100% beating spheroids, consistently containing &#62; 80% cardiac troponin T-positive cells after 15 days of culture, validated in multiple hPSC lines. We also report on a variation of this protocol for use with cell lines not currently adapted to single-cell passaging, the success of which has been verified in 42 hPSC lines. Cardiomyocytes generated using these protocols express lineage-specific markers and show expected electrophysiological functionalities. Our protocol presents a simple, efficient and robust platform for the large-scale production of human cardiomyocytes.

Here, we present a robust, fast and scalable cardiomyocyte differentiation protocol for human pluripotent stem cells (hPSCs). Cardiomyocytes derived using this large-scale method can provide sufficient cell numbers for their effective use in human cardiovascular disease modeling, high-throughput drug screening, and potentially clinical applications.

Maximizing the benefit of human pluripotent stem cells (hPSCs) for research, disease modeling, pharmaceutical and clinical applications requires robust ...

Generation of Induced Pluripotent Stem Cells from Human Melanoma Tumor-infiltrating Lymphocytes

Adoptive transfer of ex vivo expanded autologous tumor-infiltrating lymphocytes (TILs) can mediate durable and complete responses in significant subsets of patients with metastatic melanoma. Major obstacles of this approach are the reduced viability of transferred T cells, caused by telomere shortening, and the limited number of TILs obtained from patients. Less-differentiated T cells with long telomeres would be an ideal T cell subset for adoptive T cell therapy;however, generating large numbers of these less-differentiated T cells is problematic. This limitation of adoptive T cell therapy can be theoretically overcome by using induced pluripotent stem cells (iPSCs) that self-renew, maintain pluripotency, have elongated telomeres, and provide an unlimited source of autologous T cells for immunotherapy. Here, we present a protocol to generate iPSCs using Sendai virus vectors for the transduction of reprogramming factors into TILs. This protocol generates fully reprogrammed, vector-free clones. These TIL-derived iPSCs might be able to generate less-differentiated patient- and tumor-specific T cells for adoptive T cell therapy.

The goal of this protocol is to show the protocol for reprogramming melanoma tumor-infiltrating lymphocytes into induced pluripotent stem cells.

Adoptive transfer of ex vivo expanded autologous tumor-infiltrating lymphocytes (TILs) can mediate durable and complete responses in significant subsets ...

Rapid Neuronal Differentiation of Induced Pluripotent Stem Cells for Measuring Network Activity on Micro-electrode Arrays

Neurons derived from human induced Pluripotent Stem Cells (hiPSCs) provide a promising new tool for studying neurological disorders. In the past decade, many protocols for differentiating hiPSCs into neurons have been developed. However, these protocols are often slow with high variability, low reproducibility, and low efficiency. In addition, the neurons obtained with these protocols are often immature and lack adequate functional activity both at the single-cell and network levels unless the neurons are cultured for several months. Partially due to these limitations, the functional properties of hiPSC-derived neuronal networks are still not well characterized. Here, we adapt a recently published protocol that describes production of human neurons from hiPSCs by forced expression of the transcription factor neurogenin-212. This protocol is rapid (yielding mature neurons within 3 weeks) and efficient, with nearly 100% conversion efficiency of transduced cells (&#62;95% of DAPI-positive cells are MAP2 positive). Furthermore, the protocol yields a homogeneous population of excitatory neurons that would allow the investigation of cell-type specific contributions to neurological disorders. We modified the original protocol by generating stably transduced hiPSC cells, giving us explicit control over the total number of neurons. These cells are then used to generate hiPSC-derived neuronal networks on micro-electrode arrays. In this way, the spontaneous electrophysiological activity of hiPSC-derived neuronal networks can be measured and characterized, while retaining interexperimental consistency in terms of cell density. The presented protocol is broadly applicable, especially for mechanistic and pharmacological studies on human neuronal networks.

We modify and implement a previously published protocol describing the rapid, reproducible, and efficient differentiation of human&#160;induced&#160;Pluripotent Stem Cells (hiPSCs) into excitatory cortical neurons12. Specifically, our modification allows for control of neuronal cell density and use on micro-electrode arrays to measure electrophysiological properties at the network level.

Neurons derived from human induced Pluripotent Stem Cells (hiPSCs) provide a promising new tool for studying neurological disorders. In the past decade, ...

Directed Differentiation of Primitive and Definitive Hematopoietic Progenitors from Human Pluripotent Stem Cells

One of the major goals for regenerative medicine is the generation and maintenance of hematopoietic stem cells (HSCs) derived from human pluripotent stem cells (hPSCs). Until recently, efforts to differentiate hPSCs into HSCs have predominantly generated hematopoietic progenitors that lack HSC potential, and instead resemble yolk sac hematopoiesis.&#160;These resulting hematopoietic progenitors may have limited utility for in vitro disease modeling of various adult hematopoietic disorders, particularly those of the lymphoid lineages. However, we have recently described methods to generate erythro-myelo-lymphoid multilineage definitive hematopoietic progenitors from hPSCs using a stage-specific directed differentiation protocol, which we outline here. Through enzymatic dissociation of hPSCs on basement membrane matrix-coated plasticware, embryoid bodies (EBs) are formed. EBs are differentiated to mesoderm by recombinant BMP4, which is subsequently specified to the definitive hematopoietic program by the GSK3&#946; inhibitor, CHIR99021. Alternatively, primitive hematopoiesis is specified by the PORCN inhibitor, IWP2. Hematopoiesis is further driven through the addition of recombinant VEGF and supportive hematopoietic cytokines. The resulting hematopoietic progenitors generated using this method have the potential to be used for disease and developmental modeling, in vitro.

Here, we present human pluripotent stem cell (hPSC) culture protocols, used to differentiate hPSCs into CD34+ hematopoietic progenitors. This method uses stage-specific manipulation of canonical WNT signaling to specify cells exclusively to either the definitive or primitive hematopoietic program.

One of the major goals for regenerative medicine is the generation and maintenance of hematopoietic stem cells (HSCs) derived from human pluripotent stem ...

In Vitro Differentiation of Human Pluripotent Stem Cells into Trophoblastic Cells

The placenta is the first organ to develop during embryogenesis and is required for the survival of the developing embryo. The placenta is comprised of various trophoblastic cells that differentiate from the extra-embryonic trophectoderm cells of the preimplantation blastocyst. As such, our understanding of the early differentiation events of the human placenta is limited because of ethical and legal restrictions on the isolation and manipulation of human embryogenesis. Human pluripotent stem cells (hPSCs) are a robust model system for investigating human development and can also be differentiated in vitro into trophoblastic cells that express markers of the various trophoblast cell types. Here, we present a detailed protocol for differentiating hPSCs into trophoblastic cells using bone morphogenic protein 4 and inhibitors of the Activin/Nodal signaling pathways. This protocol generates various trophoblast cell types that can be transfected with siRNAs for investigating loss-of-function phenotypes or can be infected with pathogens. Additionally, hPSCs can be genetically modified and then differentiated into trophoblast progenitors for gain-of-function analyses. This in vitro differentiation method for generating human trophoblasts starting from hPSCs overcomes the ethical and legal restrictions of working with early human embryos, and this system can be used for a variety of applications, including drug discovery and stem cell research.

In Vitro Differentiation of Human Pluripotent Stem Cells into Trophoblastic Cells

Here, we present a protocol to efficiently generate human trophoblastic cells from human pluripotent stem cells using bone morphogenic protein 4 and inhibitors of the Activin/Nodal pathways. This method is suitable for the efficient differentiation of human pluripotent stem cells and can generate large quantities of cells for genetic manipulation.

The placenta is the first organ to develop during embryogenesis and is required for the survival of the developing embryo. The placenta is comprised of ...

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 ...

Rapid, Directed Differentiation of Retinal Pigment Epithelial Cells from Human Embryonic or Induced Pluripotent Stem Cells

We describe a robust method to direct the differentiation of pluripotent stem cells into retinal pigment epithelial cells (RPE). The purpose of providing a detailed and thorough protocol is to clearly demonstrate each step and to make this readily available to researchers in the field. This protocol results in a homogenous layer of RPE with minimal or no manual dissection needed. The method presented here has been shown to be effective for induced pluripotent stem cells (iPSC) and human embryonic stem cells. Additionally, we describe methods for cryopreservation of intermediate cell banks that allow long-term storage. RPE generated using this protocol might be useful for iPSC disease-in-a-dish modeling or clinical application.

This protocol describes how to produce retinal pigment epithelial cells (RPE) from pluripotent stem cells. The method uses a combination of growth factors and small molecules to direct the differentiation of stem cells into immature RPE in fourteen days and mature, functional RPE after three months.

We describe a robust method to direct the differentiation of pluripotent stem cells into retinal pigment epithelial cells (RPE). The purpose of providing ...

Efficient and Scalable Directed Differentiation of Clinically Compatible Corneal Limbal Epithelial Stem Cells from Human Pluripotent Stem Cells

Corneal limbal epithelial stem cells (LESCs) are responsible for continuously renewing the corneal epithelium, and thus maintaining corneal homeostasis and visual clarity. Human pluripotent stem cell (hPSC)-derived LESCs provide a promising cell source for corneal cell replacement therapy. Undefined, xenogeneic culture and differentiation conditions cause variation in research results and impede the clinical translation of hPSC-derived therapeutics. This protocol provides a reproducible and efficient method for hPSC-LESC differentiation under xeno- and feeder cell-free conditions. Firstly, monolayer culture of undifferentiated hPSC on recombinant laminin-521 (LN-521) and defined hPSC medium serves as a foundation for robust production of high-quality starting material for differentiations. Secondly, a rapid and simple hPSC-LESC differentiation method yields LESC populations in only 24 days. This method includes a four-day surface ectodermal induction in suspension with small molecules, followed by adherent culture phase on LN-521/collagen IV combination matrix in defined corneal epithelial differentiation medium. Cryostoring and extended differentiation further purifies the cell population and enables banking of the cells in large quantities for cell therapy products. The resulting high-quality hPSC-LESCs provide a potential novel treatment strategy for corneal surface reconstruction to treat limbal stem cell deficiency (LSCD).

This protocol introduces a simple two-step method for differentiating corneal limbal epithelial stem cells from human pluripotent stem cells under xeno- and feeder cell-free culture conditions. The cell culture methods presented here enable cost-efficient, large-scale production of clinical quality cells applicable to corneal cell therapy use.

Corneal limbal epithelial stem cells (LESCs) are responsible for continuously renewing the corneal epithelium, and thus maintaining corneal homeostasis ...

Speed

Subtitles

Differentiation and Characterization of Osteoclasts from Human Induced Pluripotent Stem Cells