Differentiation and Characterization of Osteoclasts from Human Induced Pluripotent Stem Cells

Alexander Bl&#252;mke; Jessica Simon; Elizabeth Leber; Marta Scatena; Cecilia M. Giachelli

doi:10.3791/66527

JoVE Journal > Developmental Biology

Biología del desarrollo

Differentiation and Characterization of Osteoclasts from Human Induced Pluripotent Stem Cells

Published: March 22, 2024

doi:

10.3791/66527

Alexander Blümke^1,2, Jessica Simon¹, Elizabeth Leber¹, Marta Scatena¹, Cecilia M. Giachelli¹

¹Department of Bioengineering, Department of Medicine,University of Washington, ²Department of Orthopedics and Trauma Surgery, Medical Faculty Mannheim,Heidelberg University

Summary

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.

Abstract

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.

Introduction

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-κB 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 (IκB-α), a binding protein that binds NF-kB dimers¹⁶^,¹⁷. Hence, IκB-α 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 Rössler 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 µg/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 Rössler 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.

Protocol

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 °C before use. All centrifugation steps are performed at 37 °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

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 µg/mL. Place the well plate at 4 °C overnight.
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.
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.
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 µM of Rho kinase (ROCK) inhibitor Y-27632) using P1000 with wide bore tips.
Aspirate the basal membrane extract from the well coated the previous day (step 1.1) and transfer 1 mL of hiPSC-SFM with 10 µM Y-27632 into the well. Add resuspended iPSCs to the well to reach a final volume of 2 mL per well.
Swirl the plate to evenly distribute iPSC aggregates before placing in an incubator at 37 °C and 5% CO₂.
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.

2. Passaging iPSCs

One day prior to passaging iPSCs, coat wells of a 6-well plate with a basal membrane extract at a concentration of 200-300 µg/mL. Place the well plate at 4 °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.
Begin passaging iPSCs by removing differentiated regions or regions with many dead iPSC aggregates under the stereomicroscope by using 10 µL or 20 µL 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.
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's bottom.
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).
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.
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.
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.
Rinse the well with DMEM/F-12 and transfer the medium to the 15 mL tube with the iPSC aggregates.
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.
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.
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.
Check the well for floating aggregates and aggregate size under the stereomicroscope. Ideally, the aggregate size should be between 50-200 µm.
Swirl the well plate to distribute the cells evenly across the plate after the aggregates have been transferred and incubate at 37 °C at 5% CO₂until 70-80% confluency.

3. Freezing back iPSCs

To freeze iPSC cells back, passage cells as described above (see protocol step "2. Passaging of iPSCs"). 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.
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.
Transfer cells into prelabeled cryotubes. Close tubes and transfer tubes into a prechilled cryopreservation container at 4 °C. Store cells at -80 °C for 24-48 h.
Transfer cryovials into liquid nitrogen for long-term storage.
NOTE: A schematic summary of the OC differentiation process is illustrated in Figure 1.

4. Embryoid body induction

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.
Aspirate the spent medium from the iPSC cultures and rinse the iPSC colonies with D-PBS.
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 °C for 5-8 min.
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 µM Y-27632.
Gently detach cells by rinsing the well with the Stage 1 medium. Pool dissociated iPSCs into a 15 mL conical tube.
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.
After transferring all cells to the tube, centrifuge at 200 × 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.
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 µL of the Stage 1 differentiation medium.
NOTE: Under the microscope, cells will appear as a single-cell suspension with cells dispersed throughout the well.
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 °C incubator for 24 h.
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 µL of the spent Stage 1 differentiation medium into a Petri dish.
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.
Add 50 µL of fresh Stage 1 differentiation medium to each well of a round bottom ultra-low attachment 96-well plate using a multichannel pipette.

5. Hematopoietic differentiation

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 µg/mL. Place the well plate at 4 °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.
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 µM 2-mercaptoethanol, 25 ng/mL human interleukin 3 (hIL-3), and 100 ng/mL human macrophage colony-stimulating factor (hM-CSF) is added.
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.
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.
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.
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.
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.
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.

6. M-CSF maturation and OC differentiation

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.
To perform functional assays or imaging, detach the M-CSF matured cells 3 days after seeding by washing with pre-warmed PBS to 37 °C using a P1000 with a wide bore tip. Repeated washing steps might be needed to fully detach cells.
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.
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.).
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.

Representative Results

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).

Figure 1: Schematic illustration of the osteoclast differentiation process from human iPSCs. Illustration drawn by Hannah Blümke using Affinity Designer 2.1.1. The illustration utilizes previously used drawings³³. Please click here to view a larger version of this figure.

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 µm, B, C, H, I = 50 µm, E = 1 mm. Please click here to view a larger version of this figure.

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. Please click here to view a larger version of this figure.

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 µm, C, D = 50 µm, E, F = 1 mm. Images have been edited from previously published data³³. Please click here to view a larger version of this figure.

Discussion

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 "3. Freezing back iPSCs"), one well at the point of passaging was frozen back into one cryovial. When thawing (see protocol step "1. Thawing and propagation of human iPSCs"), 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% ± 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 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.

Divulgaciones

The authors have nothing to disclose.

Acknowledgements

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 Blümke 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.

Materials

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

Referencias

Bar-Shavit, Z. The osteoclast: A multinucleated, hematopoietic-origin, bone-resorbing osteoimmune cell. Journal of Cellular Biochemistry. 102 (5), 1130-1139 (2007).
Boyce, B. F. Advances in the regulation of osteoclasts and osteoclast functions. Journal of Dental Research. 92 (10), 860-867 (2013).
Buranaphatthana, W., et al. Engineered osteoclasts resorb necrotic alveolar bone in anti-RANKL antibody-treated mice. Bone. 153, 116144 (2021).
Wang, L., et al. Isolation, purification, and differentiation of osteoclast precursors from rat bone marrow. Journal of Visualized Experiments. 2019 (147), e58895 (2019).
Mercatali, L., et al. Development of a Human Preclinical Model of Osteoclastogenesis from Peripheral Blood Monocytes Co-cultured with Breast Cancer Cell Lines. Journal of Visualized Experiments. (127), e56311 (2017).
Marino, S., Bishop, R. T., de Ridder, D., Delgado-Calle, J., Reagan, M. R. 2D and 3D in vitro co-culture for cancer and bone cell interaction studies. Methods in Molecular Biology. 1914, 71-98 (2019).
Rementer, C. W., et al. An inducible, ligand-independent receptor activator of NF-κB gene to control osteoclast differentiation from monocytic precursors. PloS One. 8 (12), e0084465 (2013).
Rementer, C., et al. Engineered myeloid precursors differentiate into osteoclasts and resorb heterotopic ossification in mice. Research Square Priprint. , 2156913 (2022).
Bjelić, D., Finšgar, M. Bioactive coatings with anti-osteoclast therapeutic agents for bone implants: Enhanced compliance and prolonged implant life. Pharmacological Research. 176, 106060 (2022).
Minkin, C., Marinho, V. C. Role of the osteoclast at the bone-implant interface. Advances in dental research. 13, 49-56 (1999).
Kodama, J., Kaito, T. Osteoclast multinucleation: Review of current literature. International Journal of Molecular Sciences. 21 (16), 1-35 (2020).
Mun, S. H., Park, P. S. U., Park-Min, K. H. The M-CSF receptor in osteoclasts and beyond. Experimental & Molecular Medicine. 52 (8), 1239-1254 (2020).
Plotkin, L. I., Bivi, N. Local regulation of bone cell function. Basic and Applied Bone Biology. , 47-73 (2014).
Hodge, J. M., Kirkland, M. A., Nicholson, G. C. Multiple roles of M-CSF in human osteoclastogenesis. Journal of Cellular Biochemistry. 102 (3), 759-768 (2007).
Ono, T., Nakashima, T. Recent advances in osteoclast biology. Histochemistry and Cell Biology. 149 (4), 325-341 (2018).
Boyce, B. F., Xiu, Y., Li, J., Xing, L., Yao, Z. NF-κB-mediated regulation of osteoclastogenesis. Endocrinology and Metabolism. 30 (1), 35 (2015).
Novack, D. V. Role of NF-κB in the skeleton. Cell Research. 21 (1), 169-182 (2010).
Kim, K., Lee, S. H., Jung, H. K., Choi, Y., Kim, N. NFATc1 induces osteoclast fusion via up-regulation of Atp6v0d2 and the dendritic cell-specific transmembrane protein (DC-STAMP). Molecular Endocrinology. 22 (1), 176-185 (2008).
Wu, H., Xu, G., Li, Y. P. Atp6v0d2 is an essential component of the osteoclast-specific proton pump that mediates extracellular acidification in bone resorption. Journal of Bone and Mineral Research. 24 (5), 871 (2009).
Vignery, A. Macrophage fusion: The making of osteoclasts and giant cells. The Journal of Experimental Medicine. 202 (3), 337 (2005).
Chiu, Y. H., Ritchlin, C. T. DC-STAMP: A key regulator in osteoclast differentiation. Journal of Cellular Physiology. 231 (11), 2402 (2016).
Riedlova, P., Sood, S., Goodyear, C. S., Ansalone, C. Differentiation of functional osteoclasts from human peripheral blood CD14+ monocytes. Journal of Visualized Experiments. (191), e64698 (2023).
Gothot, A., Pyatt, R., McMahel, J., Rice, S., Srour, E. F. Functional heterogeneity of human CD34+ cells isolated in subcompartments of the G0 /G1 phase of the cell cycle. Blood. 90 (11), 4384-4393 (1997).
Dinella, J., Koster, M. I., Koch, P. J. Use of induced pluripotent stem cells in dermatological research. The Journal of Investigative Dermatology. 134 (8), e23 (2014).
Grigoriadis, A. E., et al. Directed differentiation of hematopoietic precursors and functional osteoclasts from human ES and iPS cells. Blood. 115 (14), 2769-2776 (2010).
Rössler, U., et al. Efficient generation of osteoclasts from human induced pluripotent stem cells and functional investigations of lethal CLCN7-related osteopetrosis. Journal of Bone and Mineral Research: The Official Journal of the American Society for Bone and Mineral Research. 36 (8), 1621-1635 (2021).
Chen, I. -. P. Differentiation of human induced pluripotent stem cells (hiPSCs) into osteoclasts. Bio-Protocol. 10 (24), e3854-e3854 (2020).
Buchrieser, J., James, W., Moore, M. D. Human induced pluripotent stem cell-derived macrophages share ontogeny with MYB-independent tissue-resident macrophages. Stem Cell Reports. 8 (2), 334-345 (2017).
Abe, K., et al. Endoderm-specific gene expression in embryonic stem cells differentiated to embryoid bodies. Experimental Cell Research. 229 (1), 27-34 (1996).
Perlingeiro, R. C. R., Kyba, M., Daley, G. Q. Clonal analysis of differentiating embryonic stem cells reveals a hematopoietic progenitor with primitive erythroid and adult lymphoid-myeloid potential. Development. 128 (22), 4597-4604 (2001).
Pettinato, G., Wen, X., Zhang, N. Engineering strategies for the formation of embryoid bodies from human pluripotent stem cells. Stem Cells and Development. 24 (14), 1595 (2015).
Ruiz, J. P., et al. Robust generation of erythroid and multilineage hematopoietic progenitors from human iPSCs using a scalable monolayer culture system. Stem Cell Research. 41, 101600 (2019).
Blümke, A., et al. Comparison of osteoclast differentiation protocols from human induced pluripotent stem cells of different tissue origins. Stem Cell Research & Therapy. 14 (1), 319 (2023).
Klepikova, A., et al. iPSC-Derived Macrophages: The differentiation protocol affects cell immune characteristics and differentiation trajectories. International Journal of Molecular Sciences. 23 (24), 16087 (2022).
Neely, M. D., Tidball, A. M., Aboud, A. A., Ess, K. C., Bowman, A. B. Induced pluripotent stem cells (iPSCs): An emerging model system for the study of human neurotoxicology. Neuromethods. 56, 27-61 (2011).
Doulatov, S., et al. Induction of multipotential hematopoietic progenitors from human pluripotent stem cells via re-specification of lineage-restricted precursors. Cell Stem Cell. 13 (4), 459-470 (2013).
Van Winkle, A. P., Gates, I. D., Kallos, M. S. Mass transfer limitations in embryoid bodies during human embryonic stem cell differentiation. Cells, Tissues, Organs. 196 (1), 34-47 (2012).
Kim, J. M., et al. Assessment of differentiation aspects by the morphological classification of embryoid bodies derived from human embryonic stem cells. Stem Cells and Development. 20 (11), 1925-1935 (2011).
Mohr, J. C., et al. The microwell control of embryoid body size in order to regulate cardiac differentiation of human embryonic stem cells. Biomaterials. 31 (7), 1885-1893 (2010).
Lopez-Yrigoyen, M., et al. Production and characterization of human macrophages from pluripotent stem cells. Journal of Visualized Experiments. (158), e61038 (2020).
Cao, X., et al. Differentiation and functional comparison of monocytes and macrophages from hiPSCs with peripheral blood derivatives. Stem Cell Reports. 12 (6), 1282 (2019).
Omata, Y., et al. Interspecies single-cell RNA-seq analysis reveals the novel trajectory of osteoclast differentiation and therapeutic targets. JBMR Plus. 6 (7), e10631 (2022).
Hodge, J. M., Kirkland, M. A., Nicholson, G. C. Multiple roles of M-CSF in human osteoclastogenesis. Journal of Cellular Biochemistry. 102 (3), 759-768 (2007).
Kim, N., Takami, M., Rho, J., Josien, R., Choi, Y. A novel member of the leukocyte receptor complex regulates osteoclast differentiation. The Journal of Experimental Medicine. 195 (2), 201 (2002).
Nedeva, I. R., Vitale, M., Elson, A., Hoyland, J. A., Bella, J. Role of OSCAR signaling in psteoclastogenesis and bone disease. Frontiers in Cell and Developmental Biology. 9, 641162 (2021).
Bernhardt, A., Schamel, M., Gbureck, U., Gelinsky, M. Osteoclastic differentiation and resorption is modulated by bioactive metal ions Co2+, Cu2+ and Cr3+ incorporated into calcium phosphate bone cements. PLoS One. 12 (8), e0182109 (2017).
Akasaka, T., et al. Different micro/nano-scale patterns of surface materials influence osteoclastogenesis and actin structure. Nano Research. 15 (5), 4201-4211 (2022).
Hobolt-Pedersen, A. S., Delaissé, J. M., Søe, K. Osteoclast fusion is based on heterogeneity between fusion partners. Calcified Tissue International. 95 (1), 73 (2014).

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Blümke, A., Simon, J., Leber, E., Scatena, M., Giachelli, C. M. Differentiation and Characterization of Osteoclasts from Human Induced Pluripotent Stem Cells. J. Vis. Exp. (205), e66527, doi:10.3791/66527 (2024).