R. Z. is supported by UTHealth Innovation for Cancer Prevention Research Training Program Pre-Doctoral Fellowship (Cancer Prevention and Research Institute of Texas grant RP160015). J.T. is supported by the Ke Lin Program of the First Affiliated Hospital of Sun Yat-sen University. D.-F.L. is the CPRIT scholar in Cancer Research and supported by NIH Pathway to Independence Award R00 CA181496 and CPRIT Award RR160019.

This work was approved by The University of Texas Health Science Center at Houston (UTHealth) Animal Welfare Committee. The experiments are performed in strict accordance with the standards established by the UTHealth Center for Laboratory Animal Medicine &#38; Care (CLAMC) which is accredited by American Association for Laboratory Animal Care (AAALAC International).The human subjects in this study fall under Scenario A (&#34;No Human Subjects Research&#34;) as defined by the NIH SF424 documentation. Therefore, it does not need any approval by UTHealth human research ethics committee.
1. Generation of LFS iPSCs (Figure 1A)
<ol>
	<li>On day -2, plate 2 x 104 LFS patient fibroblasts into one well of a 6-well plate, and culture the cells with fibroblast medium (Table 1) at 37 &#176;C in a humidified 5% CO2 incubator. Ensure that the fibroblasts reach 50 - 60% confluence on the day of transduction (Day 0).</li>
	<li>Perform Sendai virus transduction on Day 0. To do this, replace the spent medium with pre-warmed fibroblast medium. Thaw the commercial Sendai virus reprogramming kit (see Table of Materials) on ice. Infect LFS fibroblasts with mixed Sendai viruses according to the manufacturer&#39;s instructions.</li>
	<li>Maintenance of the transduced cells in fibroblast medium (Day 1 - 6):
	<ol>
		<li>On the day 1, 24 h after transduction, remove the fibroblast medium containing remaining Sendai virus and replace it with 2 mL fresh fibroblast medium. Culture the transduced fibroblasts at 37 &#176;C in a humidified 5% CO2 incubator.</li>
		<li>Change the medium with the fibroblast medium every other day from the Day 2 - 6.</li>
	</ol>
	</li>
	<li>Maintenance of the transduced cells in the feeder culture condition (Day 7 - 28):
	<ol>
		<li>Prepare irradiated CF1 mouse embryonic fibroblast (MEF) culture dishes on Day 6. To do this coat six 100 mm tissue culture dishes by adding 5 mL of 0.1% gelatin to each plate at room temperature for 30 min. Aspirate the 0.1% gelatin after coating.</li>
		<li>Remove MEFs from liquid nitrogen container. Thaw MEFs using the commercially available thawing system following the manufacturer&#39;s instructions. Plate the cells on the gelatin-coated plates with MEF/MSC culture medium (Table 1) at a seeding density of around 6.7 x 105 cells per 100 mm tissue culture dish.</li>
		<li>Inoculate the transduced cells on MEF plates (Day 7): Trypsinize the transduced cells using 1 mL of 0.25% Trypsin-EDTA per 100 mm dish for 2 - 4 min at room temperature. Neutralize trypsin with 9 mL of fibroblast medium. Transfer cells to conical tubes and centrifuge cells at 230 x g at 37 &#176;C for 4 min. Plate the transduced fibroblasts on the six MEF dishes prepared on Day 6 and culture them in 8 mL of fibroblast medium per dish. 
		NOTE: The confluency of MEFs is around 20% before seeding transduced cell on MEF plates.</li>
		<li>On the day 8, discard the fibroblast medium and replace with hESC medium (Table 1).</li>
		<li>Wait for the emergence of iPS clones from Days 9 - 28 (Figure 1C). Change the spent hESC medium every other day and monitor the emergence of iPS clones under the microscope. Wait for iPSCs clones big enough to be observed by the naked eye and then proceed to expand iPS clones in feeder-free culture condition.</li>
	</ol>
	</li>
	<li>Expansion of the emerged iPS clones in feeder-free culture condition (Day 29 - 79~99):
	<ol>
		<li>To prepare the matrix-coated plate, coat a 48-well plate with 120 &#181;L of the basement membrane matrix coating solution (Table 1) per well. Make sure the basement membrane matrix coating solution covers the well and coats the culture dish surface. Leave it at room temperature for 1 h. Aspirate the coating solution immediately before use and add 250 &#181;L of hESC medium per well with 2 &#181;M ROCK inhibitor Thiazovivin.</li>
		<li>On the Day 28, aspirate the spent hESC medium and wash iPS clones with 1x DPBS. Pick up visible iPS clones with a 1 mL pipette tip and transfer into a well of a treated 48-well plate (step 1.4.1). Seed one colony per well of 48-well plate.</li>
		<li>Centrifuge the plate at 230 x g for 4 min to facilitate the cell attachment. Culture iPS clones at 37 &#176;C in a humidified 5% CO2 incubator.</li>
		<li>The following day (Day 29), remove the spent hESC medium and add 250 &#181;L of the commercially available iPSC medium in each well.</li>
		<li>Maintain iPS clones at 37 &#176;C in a humidified 5% CO2 incubator, and change the iPSC medium every other day.</li>
		<li>When iPS clones reach 85% confluence, subculture cells at a 1:10 ratio by culture area. Add 60 &#181;L of the commercial passaging solution to detach the cells and incubate at 37 &#176;C for 3 min.</li>
		<li>Seed half of the detached iPSCs on a basement membrane matrix-coated 12-well plate in 1 mL of hESC medium with 2 &#181;M ROCK inhibitor Thiazovivin. Culture iPS clones in a 12-well plate with 1 mL of the commercially available iPSC medium per well and subculture iPSCs at a 1:10 ratio until they reach passage 10. 
		NOTE: iPSCs are sub cultured every 5 - 7 days at a 1:10 ratio. It takes up to 10 weeks for iPS clones to reach passage 10. iPSC characterizations including cell morphology, alkaline phosphatase (AP) activity, and the expression of pluripotency factors and hESC markers, can be examined after confirming the removal of Sendai virus in iPSCs (usually demonstrated after around 10 passages) (Figure 1B-E). Antibody and primer information for iPSC characterization are included in Table 2 and 4.</li>
	</ol>
	</li>
</ol>
2. Differentiation of LFS iPSCs to Mesenchymal Stem Cells (MSCs) (Figure 2A)
<ol>
	<li>Culture iPSC clones on MEF plates for at least 2 weeks (Day ~ -14 - 0): Prepare the MEF dishes (100 mm tissue culture dishes) as previously described (1.3.1 - 1.3.2). Maintain iPSCs in hESC medium and change medium every other day. Whenever iPSCs reach 90% confluence, subculture iPSCs by 1:10 dilution.</li>
	<li>Removal of MEFs from iPSCs (Day 0):
	<ol>
		<li>After maintaining iPSCs on MEFs for 2 weeks, culture iPSCs in 100 mm tissue culture dishes until cells reach 80 - 90% confluence. Remove the spent hESC medium and wash iPSCs with 5 mL of 1x DPBS. Add 1 mL of the cell detachment solution to detach the cells and incubate at room temperature for 3 min.</li>
		<li>Add 9 mL of MEF/MSC culture medium to the plate to neutralize the cell detachment solution activity and transfer the detached cells to a new 100 mm tissue culture dish without any coating.</li>
		<li>Incubate the cells at room temperature for 30 min to allow the MEFs to attach to the plate.</li>
		<li>Carefully collect the supernatant which contains unattached iPSCs, and centrifuge at 230 x g at 4 &#176;C for 4 min. 
		NOTE: Carelessly flushing the plate bottom leads to detachment of MEFs.</li>
	</ol>
	</li>
	<li>Differentiation of iPSCs toward MSCs (Day 0 - 28):
	<ol>
		<li>Coat three wells of a 6-well plate with 1 mL of 0.1% gelatin per well at room temperature for 30 min.</li>
		<li>Remove the supernatant of the collected iPSCs from 2.2.4.</li>
		<li>Resuspend iPSCs in 6 mL of MSC differentiation medium (Table 1) and seed cells in the three coated wells with 2 mL per well (Day 0).</li>
		<li>Change the MSC differentiation medium every two days to remove unattached and/or dead cells (Day 1 - 28).</li>
	</ol>
	</li>
	<li>Maturation of iPSC-derived MSCs (Day 28 - 45):
	<ol>
		<li>Remove the spent MSC differentiation medium and wash cells with 1 mL of 1x DPBS.</li>
		<li>Trypsinize MSCs with 0.5 mL of 0.25% Trypsin-EDTA per well at room temperature for 3 min.</li>
		<li>Neutralize trypsin by 1.5 mL of the MEF/MSC culture medium and collect MSCs from three wells into one 15 mL conical tube. Centrifuge at 230 x g at 4 &#176;C for 4 min.</li>
		<li>Remove the supernatant and resuspend MSCs in 3 mL of MEF/MSC culture medium and plate on one 60 mm 0.1% gelatin-coated plate (Day 28).</li>
		<li>Culture MSCs in MEF/MSC culture medium and change medium every two days. When the cells reach 100% confluence, subculture MSCs at a 1:3 ratio using 0.25% Trypsin-EDTA. 
		NOTE: The MSCs display a fibroblast-like morphology (Figure 2B). When MSCs grow in a high density, elongated MSCs can form a swirl-like pattern (Figure 2B).</li>
		<li>Perform the immunofluorescent staining to evaluate iPSC-derived MSCs by examining MSC surface markers CD44, CD73, CD105, and CD166 (Figure 2C). 
		NOTE: The dilution of antibodies are shown in Table 2. Immunofluorescent staining of live cells is performed according to manufacturer&#39;s instructions.</li>
		<li>After confirmation, expand MSCs at a 1:3 ratio in 100 mm tissue culture dishes. Freeze down vials (3 vials per 100 mm tissue culture dish) using the freezing medium containing 10% DMSO and 90% FBS. 
		NOTE: To enrich the MSC population, MSCs can be sorted using CD105+, CD24- criteria by flow cytometry9,17. iPSC-derived MSCs differentiated using the PDGF-AB-based method commonly can be maintained for 2 - 3&#160;months in culture without loss of MSC identity9. Freeze MSCs from an early passage number after confirming MSC characteristics.</li>
	</ol>
	</li>
</ol>
3. Differentiation of LFS MSCs to Osteoblasts (Figure 3A)
<ol>
	<li>Preparation of MSCs for the osteogenic differentiation (Day -1)
	<ol>
		<li>To ensure an adequate quantity of cells to complete the osteogenic differentiation protocol, begin with MSCs cultured in 100 mm tissue culture dishes to 95% confluence. Remove the spent culture medium and wash MSCs with 5 mL of 1x DPBS.</li>
		<li>Trypsinize MSCs with 1 mL of 0.25% trypsin-EDTA per 100 mm dish at room temperature for 3 min. To ensure the cells are not over trypsinized, stop trypsinization when 50% of the cell detachment is observed under the microscope.</li>
		<li>Neutralize trypsin by 9 mL of MEF/MSC culture medium and collect MSCs from 3 wells into one 15 mL conical tube. Centrifuge at 230 x g at 4 &#176;C for 4 min.</li>
		<li>Aspirate the supernatant, resuspend MSCs in 5 mL of MSC medium, and count the number of cells by a hemocytometer.</li>
		<li>Plate proper number of MSCs on a culture plate. 
		NOTE: The detail of cell numbers of 12-well plate, 6 well-plate, and 100 mm tissue culture dishes are summarized in Table 3.</li>
	</ol>
	</li>
	<li>Induction of osteogenic differentiation by osteoblast differentiation medium (ODM) (Table 1) (Day 0 - 24):
	<ol>
		<li>Aspirate MEF/MSC culture medium and replace with the proper volume of ODM (1 mL, 2 mL, and 8 mL for each well of 12-well plate, each well of 6-well plate, and 100 mm tissue culture dish, respectively) to differentiate MSCs to osteoblasts (Day 0).</li>
		<li>Aspirate the spent ODM and replace with the proper volume of ODM every two days during the osteogenic differentiation process.</li>
		<li>Perform the alkaline phosphatase (AP) and the Alizarin Red S (ARS) staining to detect bone-associated alkaline phosphatase produced by preosteoblasts and mineral deposition produced by mature osteoblasts, respectively, on Day 3, 6, 9, 12, 15, 18, 21, and 24 (Figure 3B). 
		NOTE: AP staining and ARS staining are performed using commercially available kit following manufacturer protocol.</li>
		<li>Examine the expression levels of preosteoblast and mature osteoblast markers (ALPL and COL1A1 for preosteoblasts; PTH1R and BGLAP for mature osteoblasts) by qRT-PCR (Figure 3C). 
		NOTE: Positive AP staining is expected after Day 9 of osteogenic differentiation and positive ARS staining can be expected after Day 21 of osteogenic differentiation. The MSCs growing in MEF/MSC culture medium can be served as a negative control of AP staining and ARS staining.</li>
	</ol>
	</li>
</ol>
4. The Xenograft Model to Study In Vivo Tumorigenesis of LFS MSC-derived Osteoblasts (Figure 4A)
<ol>
	<li>Differentiation of LFS MSCs to osteoblasts:
	<ol>
		<li>Seed 3 - 4.5 x 105 MSCs in 100 mm tissue culture dish. Prepare five dishes for one subcutaneous injection.</li>
		<li>Culture MSCs in ODM to differentiate MSCs to osteoblasts as described in 3.2 until Day 14.</li>
	</ol>
	</li>
	<li>Preparation of differentiated osteoblasts for subcutaneous injection:
	<ol>
		<li>To prepare osteoblast detachment solution (Table 1), dissolve 1 g type II collagenase in 1,000 mL of &#945;MEM medium to make the collagenase II solution (1 mg/mL). Keep the collagenase II solution at -20 &#176;C. Mix 0.25% Trypsin-EDTA and the collagenase II solution at a 1:1 ratio to make the osteoblasts detachment solution.</li>
		<li>Remove the spent ODM and wash differentiated osteoblasts with 5 mL of 1x DPBS per 100 mm dish.</li>
		<li>Add 1.5 mL of osteoblast detachment solution per 100 mm dish and incubate dishes at 37 &#176;C for 30 min. Examine and ensure the osteoblast detachment by microscopy.</li>
		<li>Neutralize the osteoblasts detachment solution by ODM and collect the detached osteoblasts in a 50 mL conical tube. Centrifuge at 230 x g at 4 &#176;C for 5 min.</li>
		<li>Remove the supernatant and resuspend cells in 25 mL of ODM medium. Pass the cells through a 70 &#181;m cell strainer to remove aggregated cells. Count the cell numbers using a hemocytometer.</li>
		<li>Prepare 1 x 107 osteoblasts per subcutaneous injection, and centrifuge cells at 230 x g at 4 &#176;C for 4 min.</li>
		<li>Resuspend the 1 x 107 osteoblasts in 50 &#181;L ice-cold 1x DPBS. Mix differentiated osteoblasts with 50 &#181;L of phenol-red free basement membrane matrix (osteoblasts suspension and basement membrane matrix were mixed at a 1:1 ratio) and maintain osteoblasts on ice before injection.</li>
	</ol>
	</li>
	<li>In vivo xenograft tumor model of LFS MSC-derived osteoblasts:
	<ol>
		<li>Anesthetize 8-week-old female immunocompromised NU/NU mice in a chamber supplying 5% (v/v) inhaled isoflurane in 1 L/min of oxygen (provided by CLAMC). After the animal becomes recumbent, switch the anesthetic delivery system to the nose cone, supplying 2% (v/v) inhaled isoflurane in 200mL/min of oxygen (provided by CLAMC). Monitor the depth of anesthesia by checking the corneal and pedal reflexes to make the mice are sufficiently anesthetized and are not suffering from discomfort during the procedure.</li>
		<li>Disinfect the area to be injected with alternating swabs of chlorhexidine and 70% isopropanol. Use 26 G needle to perform subcutaneous injection of basement membrane matrix-mixed LFS osteoblasts into one side of hind legs of 8-week-old immunocompromised NU/NU mice (Figure 4A).</li>
		<li>Monitor mice by performing weekly palpation at injection sites for growth of subcutaneous nodular densities. Tumor formation can be observed around 6-10 weeks after injection (Figure 4B).</li>
		<li>Euthanize the mice by carbon dioxide asphyxiation method followed by cervical dislocation. Dissect the developed tumors. Determine the tumor grade and the differentiation index, preosteoblasts, and mature osteoblasts by various staining methods (e.g. Hematoxylin and Eosin (H&#38;E) staining, Picro Sirius Red staining, and von Kossa staining respectively).</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

To achieve higher MSC differentiation efficiency, several aspects are critical. One is the culture condition of iPSCs before initiating MSC differentiation. The protocol presented in the manuscript is based on previous studies 9,17. iPSCs need to be cultured on MEFs for at least 2 weeks. Maintaining iPSCs in good conditions on MEFs are critical for cells to attach on the gelatin-coated plate for MSC differentiation. Another important aspect is the density of iPSC...

Between 2006 and 2007, several breakthrough findings from the laboratories of Drs. Shinya Yamanaka and James A. Thomson led to the development of induced pluripotent stem cells (iPSCs)1,2,3. By reprogramming somatic cells with defined transcriptional factors to form iPSCs, researchers were able to generate cells with key characteristics namely, pluripotency, and self-renewal, which was previously thought to only exist in human embryonic stem cells (hESCs). iPSCs could be generated from any individual or patient and did not have to be derived from embryos, vastly expanding the repertoire of available diseases and backgrounds for the study. Since then, patient-derived iPSCs have been used to recapitulate the phenotype of various human diseases, from Alzheimer&#39;s disease4 and amyotrophic lateral sclerosis5 to long QT syndrome6,7,8.
These advances in iPSC research also have opened new avenues for the cancer research. Several groups recently have used patient iPSCs to model cancer development under a susceptible genetic background9,10,11, with successful application demonstrated to date in osteosarcoma9, leukemia10,11,12, and colorectal cancer13. Although iPSC-derived cancer models are still in their infancy, they have demonstrated great potential in phenocopying disease-associated malignancies, elucidating pathological mechanisms, and identifying therapeutic compounds14.
Li-Fraumeni syndrome (LFS) is an autosomal dominant hereditary cancer disorder caused by TP53 germline mutation15. Patients with LFS are predisposed to a various type of malignancies including osteosarcoma, making LFS iPSCs and their derived cells particularly well-suited to studying this malignancy16. An iPSC-based osteosarcoma model was first established in 2015 using LFS patient-derived iPSCs9 subsequently differentiated into mesenchymal stem cells (MSCs) and then to osteoblasts, the originating cells of osteosarcoma. These LFS osteoblasts recapitulate osteosarcoma-associated osteogenic differentiation defects and oncogenic properties, demonstrating the model potential as a &#34;bone tumor in a dish&#34; platform. Interestingly, genome-wide transcriptome analyses reveal aspects of an osteosarcoma gene signature in LFS osteoblasts and that features of this LFS gene expression profile are correlated with poor prognosis in osteosarcoma9, indicating the potential of LFS iPSCs disease models to reveal features of clinical relevance.
This manuscript provides a detailed description of how to use LFS patient-derived iPSCs to model osteosarcoma. It details the generation of LFS iPSCs, differentiation of iPSCs to MSCs and then to osteoblasts, and use of an in vivo xenograft model using LFS osteoblasts. The LFS disease model comprises several advantages, most notably the ability to generate unlimited cells at all stages of osteosarcoma development for mechanistic studies, biomarker identification, and drug screening9,14,16.
In summary, the LFS iPSC-based osteosarcoma model offers an attractive complementary system for advancing osteosarcoma research. This platform also provides a proof-of-concept for cancer modeling using patient-derived iPSCs. This strategy described below can be readily extended to model malignancies associated with other genetic disorders with cancer predispositions.

<table><tbody><tr><td>&lt;strong&gt;Plastic ware&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>100 mm Dish</td><td>Corning</td><td>430107</td><td></td></tr><tr><td>60 mm Dish</td><td>Corning</td><td>430166</td><td></td></tr><tr><td>6-well Plate</td><td>Falcon</td><td>353046</td><td></td></tr><tr><td>12-well Plate</td><td>Falcon</td><td>353043</td><td></td></tr><tr><td>48-well Plate</td><td>Falcon</td><td>353078</td><td></td></tr><tr><td>1 mL Pipet Tip</td><td>USA Scientific</td><td>1111-2721</td><td></td></tr><tr><td>200 &amp;micro;L Pipet Tip</td><td>USA Scientific</td><td>1111-0706</td><td></td></tr><tr><td>10 &amp;micro;L Pipet Tip</td><td>USA Scientific</td><td>1111-3700</td><td></td></tr><tr><td>5 mL Serological Pipette</td><td>SARSTEDT</td><td>86.1253.001</td><td></td></tr><tr><td>10 mL Serological Pipette</td><td>SARSTEDT</td><td>86.1254.001</td><td></td></tr><tr><td>25 mL Serological Pipette</td><td>SARSTEDT</td><td>86.1685.001</td><td></td></tr><tr><td>50 mL Tube, PP</td><td>SARSTEDT</td><td>62.547.100</td><td></td></tr><tr><td>15 mL Tube, PP</td><td>SARSTEDT</td><td>62.554.100</td><td></td></tr><tr><td>&lt;strong&gt;Culture materials and Reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>CytoTune- iPS 2.0 Sendai Reprogramming Kit</td><td>Invitrogen</td><td>A16517</td><td>Commercial Sendai virus reprogramming kit</td></tr><tr><td>Corning hESC-Qualified Matrix</td><td>Corning</td><td>354277</td><td>Basement membrane matrix</td></tr><tr><td>CF1 MEFs, irradiated</td><td>ThermoFisher</td><td>A34180</td><td></td></tr><tr><td>DMEM</td><td>Sigma-Aldrich</td><td>D5671</td><td></td></tr><tr><td>DMEM/F12</td><td>Corning</td><td>10-090-CV</td><td></td></tr><tr><td>&amp;alpha;MEM</td><td>Corning</td><td>10-022-CV</td><td></td></tr><tr><td>StemMACS iPS-Brew XF</td><td>Miltenyi Biotec</td><td>130-104-368</td><td>Commercial iPSC medium</td></tr><tr><td>KnockOut DMEM/F-12</td><td>ThermoFisher</td><td>12660012</td><td></td></tr><tr><td>FBS Opti-Gold</td><td>GenDEPOT</td><td>F0900-050</td><td></td></tr><tr><td>KnockOut Serum Replacement</td><td>ThermoFisher</td><td>A3181502</td><td></td></tr><tr><td>Penicillin-Streptomycin</td><td>Sigma-Aldrich</td><td>P4333</td><td></td></tr><tr><td>MEM Nonessential Amino Acids</td><td>Corning</td><td>25-025-CI</td><td></td></tr><tr><td>L-Glutamine Solution</td><td>Sigma-Aldrich</td><td>G7513</td><td></td></tr><tr><td>2-Mercaptoethanol</td><td>Sigma-Aldrich</td><td>M3148</td><td></td></tr><tr><td>Human FGF-basic (bFGF)</td><td>PEPROTECH</td><td>100-18B</td><td></td></tr><tr><td>Recombinant Human PDGF-AB</td><td>PEPROTECH</td><td>100-00AB</td><td></td></tr><tr><td>&amp;beta;-Glycerophosphate</td><td>Sigma-Aldrich</td><td>G9422</td><td></td></tr><tr><td>Dexamethasone</td><td>Sigma-Aldrich</td><td>A4902</td><td></td></tr><tr><td>Ascorbic Acid</td><td>Sigma-Aldrich</td><td>A5960</td><td></td></tr><tr><td>Dulbecco&#039;s Phosphate-Buffered Saline, 1x 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Biosciences</td><td>560123</td><td></td></tr><tr><td>Alexa Fluor 488 Donkey Anti-Goat IgG (H+L)</td><td>Jackson ImmunoResearch</td><td>705-545-003</td><td></td></tr><tr><td>Alexa Fluor 488 Goat Anti-Rabbit IgG (H+L)</td><td>Jackson ImmunoResearch</td><td>111-545-144</td><td></td></tr><tr><td>PE Mouse Anti-Human CD105</td><td>eBioscience</td><td>12-1057-42</td><td></td></tr><tr><td>FITC Mouse Anti-Human CD44</td><td>DB Biosciences</td><td>555478</td><td></td></tr><tr><td>PE Mouse Anti-Human CD73</td><td>DB Biosciences</td><td>550257</td><td></td></tr><tr><td>PE Mouse Anti-Human CD166</td><td>DB Biosciences</td><td>560903</td><td></td></tr><tr><td>FITC Mouse Anti-Human CD24</td><td>DB Biosciences</td><td>555427</td><td></td></tr><tr><td>Donkey Serum</td><td>Jackson ImmunoResearch</td><td>017-000-121</td><td></td></tr><tr><td>Goat Serum</td><td>Jackson ImmunoResearch</td><td>005-000-121</td><td></td></tr><tr><td>Alkaline Phosphatase Staining Kit II</td><td>Stemgent</td><td>00-0055</td><td></td></tr><tr><td>Alizarin Red S</td><td>Sigma-Aldrich</td><td>A5533</td><td></td></tr><tr><td>TRIzol Reagent</td><td>ThermoFisher</td><td>15596018</td><td></td></tr><tr><td>Chloroform</td><td>ThermoFisher</td><td>C298-500</td><td></td></tr><tr><td>2-Propanol</td><td>ThermoFisher</td><td>A416-4</td><td></td></tr><tr><td>Ethanol, Absolute, Molecular Biology Grade</td><td>ThermoFisher</td><td>BP28184</td><td></td></tr><tr><td>DNase I, RNase-free (1 U/&amp;micro;L)</td><td>ThermoFisher</td><td>EN0521</td><td></td></tr><tr><td>iScript cDNA Synthesis Kit</td><td>BioRad</td><td>1708891BUN</td><td></td></tr><tr><td>iQ SYBR Green Supermix</td><td>BioRad</td><td>1708884</td><td></td></tr><tr><td>Matrigel Matrix High Concentration (HC), Phenol-Red Free</td><td>Corning</td><td>354262</td><td></td></tr><tr><td>1 mL Slip Tip Syringe, 26 Gauge x 5/8 Inch</td><td>DB Biosciences</td><td>309597</td><td></td></tr></tbody></table>

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.

This protocol presents the procedures including LFS iPSC generation, MSC differentiation, osteoblast differentiation, and in vivo tumorigenesis assay using LFS MSC-derived osteoblasts.
Scheme for the generation of LFS iPSCs from fibroblasts by using a commercially available Sendai virus reprogramming kit is shown in Figure 1A. Sendai virus-based delivery of the Yamanaka four factors is a no...

Watch this Scientific Journal Video about Modeling Osteosarcoma Using Li-Fraumeni Syndrome Patient-derived Induced Pluripotent Stem Cells at JoVE.com

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

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 ...

modeling-osteosarcoma-using-li-fraumeni-syndrome-patient-derived

Research

JoVE Journal

Cancer Research

8.8K Views.  The University of Texas Health Science Center at Houston. Hier presenteren we een protocol voor het genereren van geïnduceerde pluripotente stamcellen (iPSCs) van Li-Fraumeni syndroom (LFS) patiënt afgeleide fibroblasten, differentiatie van iPSCs via mesenchymale stamcellen (MSCs) botcellen te modelleren in vivo tumorigenesis met behulp van LFS patiënt afkomstige botcellen.

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

Establishment of Cancer Stem Cell Cultures from Human Conventional Osteosarcoma

The current improvements in therapy against osteosarcoma (OS) have prolonged the lives of cancer patients, but the survival rate of five years remains poor when metastasis has occurred. The Cancer Stem Cell (CSC) theory holds that there is a subset of tumor cells within the tumor that have stem-like characteristics, including the capacity to maintain the tumor and to resist multidrug chemotherapy. Therefore, a better understanding of OS biology and pathogenesis is needed in order to advance the development of targeted therapies to eradicate this particular subset and to reduce morbidity and mortality among patients. Isolating CSCs, establishing cell cultures of CSCs, and studying their biology are important steps to improving our understanding of OS biology and pathogenesis. The establishment of human-derived OS-CSCs from biopsies of OS has been made possible using several methods, including the capacity to create 3-dimensional stem cell cultures under nonadherent conditions. Under these conditions, CSCs are able to create spherical floating colonies formed by daughter stem cells; these colonies are termed &#34;cellular spheres&#34;. Here, we describe a method to establish CSC cultures from primary cell cultures of conventional OS obtained from OS biopsies. We clearly describe the several passages required to isolate and characterize CSCs.

The presence of cancer stem cells (CSCs) in bone sarcomas has recently been linked to their pathogenesis. In this article, we present the isolation of CSCs from primary cell cultures obtained from human biopsies of conventional osteosarcoma (OS) using the ability of CSCs to grow under nonadherent conditions.

The current improvements in therapy against osteosarcoma (OS) have prolonged the lives of cancer patients, but the survival rate of five years remains ...

Establishment of a Primary Culture of Patient-derived Soft Tissue Sarcoma

Soft tissue sarcomas (STS) represent a spectrum of heterogeneous malignancies with a difficult diagnosis, classification, and management. To date, more than 50 histological subtypes of these rare solid tumors have been identified. Thus, due to their extraordinary diversity and low incidence, our understanding of the biology of these tumors is still limited. Patient-derived cultures represent the ideal platform to study STS pathophysiology and pharmacology. We thus developed a human preclinical model of STS starting from tumor specimens harvested from patients undergoing surgical resection. Patient-derived STS cell cultures were obtained from the surgical specimens by collagenase digestion and isolated by filtration. Cells were counted, seeded, and left for 14 days in standard monolayer cultures and then processed by downstream analysis. Before performing molecular or pharmaceutical analyses, the establishment of STS primary cultures was confirmed through the evaluation of cytomorphologic features and, when available, immunohistochemical markers. This method represents a useful tool 1) to study the natural history of these poorly explored malignancies and 2) to test the effects of different drugs in an effort to learn more about their mechanisms of action.

The following protocol focuses on the establishment of a primary culture of patient-derived soft tissue sarcoma (STS). This model could help us to better understand the molecular background and pharmacological profile of these rare malignancies and could represent a starting point for further research aimed at improving STS management.

Soft tissue sarcomas (STS) represent a spectrum of heterogeneous malignancies with a difficult diagnosis, classification, and management. To date, more ...

A Preclinical Mouse Model of Osteosarcoma to Define the Extracellular Vesicle-mediated Communication Between Tumor and Mesenchymal Stem Cells

Within the tumor microenvironment, resident or recruited mesenchymal stem cells (MSCs) contribute to malignant progression in multiple cancer types. Under the influence of specific environmental signals, these adult stem cells can release paracrine mediators leading to accelerated tumor growth and metastasis. Defining the crosstalk between tumor and MSCs is of primary importance to understand the mechanisms underlying cancer progression and identify novel targets for therapeutic intervention. 
Cancer cells produce high amounts of extracellular vesicles (EVs), which can profoundly affect the behavior of target cells in the tumor microenvironment or at distant sites. Tumor EVs enclose functional biomolecules, including inflammatory RNAs and (onco)proteins, that can educate stromal cells to enhance the metastatic behavior of cancer cells or to participate in the pre-metastatic niche formation. In this article, we describe the development of a preclinical cancer mouse model that enables specific evaluation of the EV-mediated crosstalk between tumor and mesenchymal stem cells. First, we describe the purification and characterization of tumor-secreted EVs and the assessment of the EV internalization by MSCs. We then make use of a multiplex bead-based immunoassay to evaluate the alteration of the MSC cytokine expression profile induced by cancer EVs. Finally, we illustrate the generation of a bioluminescent orthotopic xenograft mouse model of osteosarcoma that recapitulates the tumor-MSC interaction, and show the contribution of EV-educated MSCs to tumor growth and metastasis formation. 
Our model provides the opportunity to define how cancer EVs shape a tumor-supporting environment, and to evaluate whether blockade of the EV-mediated communication between tumor and MSCs prevents cancer progression.

Direct injection of cancer-derived extracellular vesicles (EVs) leads to reprogramming of bone marrow supporting tumor progression; however, which cells mediate this effect is unclear. Herein, we describe a step-by-step protocol to investigate EV-mediated tumor-mesenchymal stem cell (MSC) interactions in vivo, revealing a crucial role for EV-educated MSCs in metastasis.

Within the tumor microenvironment, resident or recruited mesenchymal stem cells (MSCs) contribute to malignant progression in multiple cancer types. Under ...

Isolation and Characterization of Tumor-initiating Cells from Sarcoma Patient-derived Xenografts

The existence and importance of tumor-initiating cells (TICs) have been supported by increasing evidence during the past decade. These TICs have been shown to be responsible for tumor initiation, metastasis, and drug resistance. Therefore, it is important to develop specific TIC-targeting therapy in addition to current chemotherapy strategies, which mostly focus on the bulk of non-TICs. In order to further understand the mechanism behind the malignancy of TICs, we describe a method to isolate and to characterize TICs in human sarcomas. Herein, we show a detailed protocol to generate patient-derived xenografts (PDXs) of human sarcomas and to isolate TICs by fluorescence-activated cell sorting (FACS) using human leukocyte antigen class I (HLA-1) as a negative marker. Also, we describe how to functionally characterize these TICs, including a sphere formation assay and a tumor formation assay, and to induce differentiation along mesenchymal pathways. The isolation and characterization of PDX TICs provide clues for the discovery of potential targeting therapy reagents. Moreover, increasing evidence suggests that this protocol may be further extended to isolate and characterize TICs from other types of human cancers.

We describe a detailed protocol for the isolation of tumor-initiating cells from human sarcoma patient-derived xenografts by fluorescence-activated cell sorting, using human leukocyte antigen-1 (HLA-1) as a negative marker, and for the further validation and characterization of these HLA-1-negative tumor-initiating cells.

The existence and importance of tumor-initiating cells (TICs) have been supported by increasing evidence during the past decade. These TICs have been ...

Three-Dimensional Bone Extracellular Matrix Model for Osteosarcoma

Osteosarcoma (OS) is the most common and a highly aggressive primary bone tumor. It is characterized with anatomic and histologic variations along with diagnostic or prognostic difficulties. OS comprises genotypically and phenotypically heterogeneous cancer cells. Bone microenvironment elements are proved to account for tumor heterogeneity and disease progression. Bone extracellular matrix (BEM) retains the microstructural matrices and biochemical components of native extracellular matrix. This tissue-specific niche provides a favorable and long-term scaffold for OS cell seeding and proliferation. This article provides a protocol for the preparation of BEM model and its further experimental application. OS cells can grow and differentiate into multiple phenotypes consistent with the histopathological complexity of OS clinical specimens. The model also allows visualization of diverse morphologies and their association with genetic alterations and underlying regulatory mechanisms. As homologous to human OS, this BEM-OS model can be developed and applied to the pathology and clinical research of OS.

The bone extracellular matrix (BEM) model for osteosarcoma (OS) is well established and shown here. It can be used as a suitable scaffold for mimicking primary tumor growth in vitro and providing an ideal model for studying the histologic and cytogenic heterogeneity of OS.

Osteosarcoma (OS) is the most common and a highly aggressive primary bone tumor. It is characterized with anatomic and histologic variations along with ...

Tumorsphere Derivation and Treatment from Primary Tumor Cells Isolated from Mouse Rhabdomyosarcomas

Rhabdomyosarcoma (RMS) is the most common soft tissue sarcoma in children. Although significant efforts have enabled the identification of common mutations associated with RMS and allowed discrimination of different RMS subtypes, major challenges still exist for the development of novel treatments to further improve prognosis. Although identified by the expression of myogenic markers, there is still significant controversy over whether RMS has myogenic or non-myogenic origins, as the cell of origin is still poorly understood. In the present study, a reliable method is provided for the tumorsphere assay for mouse RMS. The assay is based on functional properties of tumor cells and allows the identification of rare populations in the tumor with tumorigenic functions. Also described are procedures for testing recombinant proteins, integrating transfection protocols with the tumorsphere assay, and evaluating candidate genes involved in tumor development and growth. Described further is a procedure for allograft transplantation of tumorspheres into recipient mice to validate tumorigenic function in vivo. Overall, the described method allows reliable identification and testing of rare RMS tumorigenic populations that can be applied to RMS arising in different contexts. Finally, the protocol can be utilized as a platform for drug screening and future development of therapeutics.

This protocol describes a reproducible method for isolation of mouse rhabdomyosarcoma primary cells, tumorsphere formation and treatment, and allograft transplantation starting from tumorspheres cultures.

Rhabdomyosarcoma (RMS) is the most common soft tissue sarcoma in children. Although significant efforts have enabled the identification of common ...

Genetics

Modeling Primary Bone Tumors and Bone Metastasis with Solid Tumor Graft Implantation into Bone

Primary bone tumors or bone metastasis from solid tumors result in painful osteolytic, osteoblastic, or mixed osteolytic/osteoblastic lesions. These lesions compromise bone structure, increase the risk of pathologic fracture, and leave patients with limited treatment options. Primary bone tumors metastasize to distant organs, with some types capable of spreading to other skeletal sites. However, recent evidence suggests that with many solid tumors, cancer cells that have spread to bone may be the primary source of cells that ultimately metastasize to other organ systems. Most syngeneic or xenograft mouse models of primary bone tumors involve intra-osseous (orthotopic) injection of tumor cell suspensions. Some animal models of skeletal metastasis from solid tumors also depend on direct bone injection, while others attempt to recapitulate additional steps of the bone metastatic cascade by injecting cells intravascularly or into the organ of the primary tumor. However, none of these models develop bone metastasis reliably or with an incidence of 100%. In addition, direct intra-osseous injection of tumor cells has been shown to be associated with potential tumor embolization of the lung. These embolic tumor cells engraft but do not recapitulate the metastatic cascade. We reported a mouse model of osteosarcoma in which fresh or cryopreserved tumor fragments (consisting of tumor cells plus stroma) are implanted directly into the proximal tibia using a minimally invasive surgical technique. These animals developed reproducible engraftment, growth, and, over time, osteolysis and lung metastasis. This technique has the versatility to be used to model solid tumor bone metastasis and can readily employ grafts consisting of one or multiple cell types, genetically-modified cells, patient-derived xenografts, and/or labeled cells that can be tracked by optical or advanced imaging. Here, we demonstrate this technique, modeling primary bone tumors and bone metastasis using solid tumor graft implantation into bone.

Bone metastasis models do not develop metastasis uniformly or with a 100% incidence. Direct intra-osseous tumor cell injection can result in embolization of the lung. We present our technique modeling primary bone tumors and bone metastasis using solid tumor graft implantation into bone, leading to reproducible engraftment and growth.

Primary bone tumors or bone metastasis from solid tumors result in painful osteolytic, osteoblastic, or mixed osteolytic/osteoblastic lesions. These ...

A Syngeneic Orthotopic Osteosarcoma Sprague Dawley Rat Model with Amputation to Control Metastasis Rate

The most recent advance in the treatment of osteosarcoma (OS) occurred in the 1980s when multi-agent chemotherapy was shown to improve overall survival compared to surgery alone. To address this problem, the aim of the study is to refine a lesser-known model of OS in rats with a comprehensive histologic, imaging, biologic, implantation, and amputation surgical approach that prolongs survival. We used an immunocompetent, outbred Sprague-Dawley (SD), syngeneic rat model with implanted UMR106 OS cell line (originating from a SD rat) with orthotopic tibial tumor implants into 3-week-old male and female rats to model pediatric OS. We found that rats develop reproducible primary and metastatic pulmonary tumors, and that limb amputations at 3 weeks post implantation significantly reduce the incidence of pulmonary metastasis and prevent unexpected deaths. Histologically, the primary and metastatic OSs in rats were very similar to human OS. Using immunohistochemistry methods, the study shows that rat OS are infiltrated with macrophages and T cells. A protein expression survey of OS cells reveals that these tumors express ErbB family kinases. Since these kinases are also highly expressed in most human OSs, this rat model could be used to test ErbB pathway inhibitors for therapy.

Here, a syngeneic orthotopic implantation followed by an amputation procedure of the osteosarcoma with spontaneous pulmonary metastasis that can be used for preclinical investigation of metastasis biology and development of novel therapeutics is described.

The most recent advance in the treatment of osteosarcoma (OS) occurred in the 1980s when multi-agent chemotherapy was shown to improve overall survival ...

Intratibial Osteosarcoma Cell Injection to Generate Orthotopic Osteosarcoma and Lung Metastasis Mouse Models

Osteosarcoma is the most common primary bone cancer in children and adolescents, with lungs as the most common metastatic site. The five-year survival rate of osteosarcoma patients with pulmonary metastasis is less than 30%.Therefore, the utilization of mouse models mimicking the osteosarcoma development in humans is of great significance for understanding the fundamental mechanism of osteosarcoma carcinogenesis and pulmonary metastasis to develop novel therapeutics. Here, detailed procedures are reported to generate the primary osteosarcoma and pulmonary metastasis mouse models via&#160;intratibia injection of osteosarcoma cells. Combined with the bioluminescence or X-ray live imaging system, these living mouse models are utilized to monitor and quantify osteosarcoma growth and metastasis. To establish this model, a basement membrane matrix containing osteosarcoma cells was loaded in a micro-volume syringe and injected into one tibia of each athymic mouse after being anesthetized. The mice were sacrificed when the primary osteosarcoma reached the size limitation in the IACUC-approved protocol. The legs bearing osteosarcoma and the lungs with metastasis lesions were separated. These models are characterized by a short incubation period, rapid growth, severe lesions, and sensitivity in monitoring the development of primary and pulmonary metastatic lesions. Therefore, these are ideal models for exploring the functions and mechanisms of specific factors in osteosarcoma carcinogenesis and pulmonary metastasis, the tumor microenvironment, and evaluating the therapeutic efficacy in vivo.

The present protocol describes intratibia osteosarcoma cell injection to generate mouse models bearing orthotopic osteosarcoma and pulmonary metastasis lesions.

Osteosarcoma is the most common primary bone cancer in children and adolescents, with lungs as the most common metastatic site. The five-year survival ...

Reprogramming PDAC and Pancreatic Cells into Induced Pluripotent Stem Cells

Reprogramming Pancreatic Ductal Adenocarcinoma to Pluripotency

The generation of induced pluripotent stem cells (iPSCs) using transcription factors has been achieved from almost any differentiated cell type and has proved highly valuable for research and clinical applications. Interestingly, iPSC reprogramming of cancer cells, such as pancreatic ductal adenocarcinoma (PDAC), has been shown to revert the invasive PDAC phenotype and override the cancer epigenome. The differentiation of PDAC-derived iPSCs can recapitulate PDAC progression from its early pancreatic intraepithelial neoplasia (PanIN) precursor, revealing the molecular and cellular changes that occur early during PDAC progression. Therefore, PDAC-derived iPSCs can be used to model the earliest stages of PDAC for the discovery of early-detection diagnostic markers. This is particularly important for PDAC patients, who are typically diagnosed at the late metastatic stages due to a lack of reliable biomarkers for the earlier PanIN stages. However, reprogramming cancer cell lines, including PDAC, into pluripotency remains challenging, labor-intensive, and highly variable between different lines. Here, we describe a more consistent protocol for generating iPSCs from various human PDAC cell lines using bicistronic lentiviral vectors. The resulting iPSC lines are stable, showing no dependence on the exogenous expression of reprogramming factors or inducible drugs. Overall, this protocol facilitates the generation of a wide range of PDAC-derived iPSCs, which is essential for discovering early biomarkers that are more specific and representative of PDAC cases.

Author Spotlight: Reprogramming Cancer Cells to iPSCs to Study Disease Progression and Treatment Targets

The present protocol describes the reprogramming of Pancreatic Ductal Adenocarcinoma (PDAC) and normal pancreatic ductal epithelial cells into induced pluripotent stem cells (iPSCs). We provide an optimized and detailed, step-by-step procedure, from preparing lentivirus to establishing stable iPSC lines.

The generation of induced pluripotent stem cells (iPSCs) using transcription factors has been achieved from almost any differentiated cell type and has ...

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

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