Primary Culture of Dental Pulp Stem Cells

Summary

This article provides a stepwise guide to establish a primary culture of dental pulps stem cells using the explant culture method and characterization of these cells based on ICSCRT guidelines. The cells isolated by this protocol can be considered as mesenchymal stem cells for further applications.

Abstract

The human dental pulp represents a promising multipotent stem cell reservoir with pre-eminent regenerative competence that can be harvested from an extracted tooth. The neural crest-derived ecto-mesenchymal origin of dental pulp stem cells (DPSCs) bestows a high degree of plasticity that owes to its multifaceted benefits in tissue repair and regeneration. There are various practical ways of harvesting, maintaining, and proliferating adult stem cells being investigated for their use in regenerative medicine. In this work, we demonstrate the establishment of a primary mesenchymal stem cell culture from dental tissue by the explant culture method. The isolated cells were spindle-shaped and adhered to the plastic surface of the culture plate. The phenotypic characterization of these stem cells showed positive expression of the international society of cell therapy (ISCT)-recommended cell surface markers for MSC, such as CD90, CD73, and CD105. Further, negligible expression of hematopoietic (CD45) and endothelial markers (CD34), and less than 2% expression of HLA-DR markers, confirmed the homogeneity and purity of the DPSC cultures. We further illustrated their multipotency based on differentiation to adipogenic, osteogenic, and chondrogenic lineages. We also induced these cells to differentiate into hepatic-like and neuronal-like cells by adding corresponding stimulation media. This optimized protocol will aid in the cultivation of a highly expandable population of mesenchymal stem cells to be utilized in the laboratory or for preclinical studies. Similar protocols can be incorporated into clinical setups for practicing DPSC-based treatments.

Introduction

Adult stem cells have transpired into a powerful therapeutic tool for cell-directed treatments and therapies due to their plasticity, paracrine mechanisms, and immunomodulatory properties^1,2,3. The encouraging data from stem cell-based preclinical studies have inspired researchers to work for the bench to-bedside translation. The type of stem cells used for stem cell therapy plays a significant role in successful outcomes. In preclinical and clinical studies, the most widely reported source for mesenchymal stem cells (MSCs) remain bone marrow^4,5. However, major drawbacks to using bone marrow-derived stem cells (BMSCs) include their rare population, highly invasive procedures for isolation, and their limited ability to expand. Therefore, alternative sources of MSCs are being explored. In this regard, dental tissues, with their ease of accessibility, enormous plasticity, high regenerative potential, and high proliferative ability, have now been deemed as a rich and potential alternative source of stem cells^6,7,8,9,10.

Dental pulp stem cells (DPSCs) were the first type of dental stem cells to be isolated and characterized by Gronthos in 2000¹¹. DPSCs have grabbed the attention for tissue engineering applications because of their high proliferation rate, significant differentiation potential, ease of accessibility with effortless culturing, and, most importantly, their ability to be obtained from a discarded tooth without any ethical concern¹². The limitations posed by other stem cell sources, such as BMSCs and adipose-derived stem cells (ADSCs), in their isolation and inadequate self-renewal capacities are circumvented by DPSCs¹³. Human DPSCs can be obtained from human primary teeth, permanent teeth, wisdom teeth, exfoliated deciduous teeth (SHEDs), and apical papillae. Moreover, DPSCs can also be isolated from supernumerary teeth, which are generally discarded¹⁴. DPSCs express neural crest-associated markers and have the potential to differentiate into neuronal cells both in vitro and in vivo¹⁵. In addition to their neurogenic potential, DPSCs can differentiate into other cell lineages, such as osteogenic, chondrogenic, adipogenic, hepatic, and myogenic, when given specific differentiation conditions¹³. Thus, these multipotent cells hold great potential for cell-based therapy and can be employed for the regeneration of various tissues. Studies have also reported the potential role of DPSCs in the reconstruction of the cornea¹⁶, repair of myocardial infarction¹⁷, and their potential therapeutic role in diseases like limb ischemia¹⁸, Alzheimer's ¹⁹, Parkinson's ²⁰, and aging²¹. Therefore, dental tissue-derived stem cells can be used not only for dental regeneration, but also for the repair and regeneration of non-dental organs like eyes¹⁶, hearts¹⁷, livers²², bones²³ etc.

There are two particular methods for the isolation of an MSC population from pulp tissue - enzymatic digestion and explant culture^24,25. Successful establishment of primary cultures without any significant difference in the quantity and properties of DPSCs have been reported by both these methods²⁶. In this study, we have focused on the isolation of DPSCs by the explant method²⁷, since this method generates DPSCs without contamination of hematopoietic and endothelial cells, as compared to enzymatic digestion which can result in fibroblast contamination²⁸.

Protocol

All the procedures described in the study have been approved by the Institute Ethics Committee (IEC# 9195/PG-12 ITRG/2571-72) of PGIMER, Chandigarh. All the cell culture related experiments need to be performed in a Class II biological safety cabinet (BSC) following aseptic technique. Dental pulp was obtained from healthy teeth of three (F/14, M/14, and M/20) patients undergoing third molar extractions for orthodontic reasons. Before the sample collection, written informed consent was obtained from the patient/guardian in accordance with the guidelines provided by the ethics committee of PGIMER, Chandigarh.

1. Establishment of primary culture of dental pulp stem cells (DPSCs) from human dental tissue

NOTE: Any decayed teeth should not be used.

1. Dental pulp sample collection and transportation to tissue culture lab
   1. Perform a tooth extraction procedure under non-traumatic and aseptic conditions. Rinse the extracted tooth thoroughly with a sterile saline solution to remove remnants of blood.
   2. Cut the extracted tooth longitudinally into two pieces using a carbide fissure bur with constant irrigation with sterile normal saline to expose the pulp.
   3. Transport the extracted tooth along with the intact pulp in sterile α-minimum essential media (α-MEM) maintained on ice to the cell and tissue culture laboratory.
      ​NOTE: Ideally, process all samples received within 2 h for cell culture. However, a recent report has suggested that processing the tooth sample even after 24 h did not affect the viability and proliferation of DPSCs, however, it did reduce their functionality²⁹.

2. Removal of pulp tissue from the tooth and cell culture of DPSCs (time: 60-120 min)

NOTE: All the steps after tooth transportation have been performed in the cell and tissue culture lab inside a biosafety cabinet level 2.

1. Place the tooth sample in sterile culture Petri dishes and thoroughly rinse (three or four times) with 1x sterile phosphate-buffered saline (PBS) solution.
2. Carefully remove the pulp tissue from the pulp cavity with the help of a sharp excavator and forceps.
3. Once the pulp is removed, clean it thoroughly with sterile PBS to remove any remnants of blood.
4. Cut the pulp tissue into small pieces of approximately 2-3 mm with the help of a surgical blade in a Petri dish.
   1. Divide the surface area of the dish into four parts by putting 1-2 mL of sterile PBS in each area. Put the pulp tissue sample in one area with PBS and cut the tissue into the smallest possible pieces with a surgical blade (the smaller the piece, the easier it will be for stem cells to migrate out of the tissue). After this, transfer each piece by lifting it into another area with PBS, and so on till the fourth area, so that the tissue pieces are properly washed and free of any contaminants.
5. Meanwhile, pour approximately 0.8-1 mL of α-MEM containing non-essential amino acids with 10% fetal bovine serum (FBS) and an antibiotic cocktail (i.e., penicillin and streptomycin) in the wells of a 6-well plate. Distribute the media uniformly by moving the plate, so as to spread it evenly on the well surface.
6. Seed the small pieces of pulp tissue or explants with the help of forceps into a single well (four to six pieces per well) of the 6-well plate and keep the plate undisturbed for 5 min in a biosafety hood.
   NOTE: This step allows/aids in the adherence of tissue/explants to the treated surfaces of culture plates.
7. Maintain the culture plate in a humidified atmosphere with 37 °C temperature and 5% CO₂ level_.
8. The next day, carefully aspirate the media from the wells without disturbing the explants and add fresh media to it. Change the media every other day at this step to prevent drying. Make sure the incubator has sufficient water in the tray so that extra evaporation does not cause the media in the plate wells to dry up.
   ​NOTE: This is a very crucial step, and disturbance can cause the explants to dislodge from their position. If the explant pieces are dislodged from their position during media transfer, try to reattach them by removing the media completely and letting them adhere to the well surface. The media should be carefully added along the wall of the well without disturbing the explants, otherwise they might detach again. Observe the cell culture plates regularly under an inverted microscope (4x or 10x magnification). Identify DPSCs by their ability to adhere to the plastic surface and their spindle-shaped morphology.
9. Once sufficient cells migrate from the explants, remove the tissues and allow the cells to proliferate and reach the optimum 70%-75% cell confluence before subculturing. The volume of α-MEM containing 10% FBS can be increased to 2 mL at this step so that the cells can divide faster. Change the media every third day at this stage.

3. DPSC expansion

1. Wash the confluent T75 culture flask (confluency: 70%-80%) of DPSCs twice with PBS (10 mL).
2. Add 2.5 mL of 0.25% trypsin-ethylenediaminetetraacetic acid (EDTA) to the side of the T75 flask and gently swirl it to ensure that the trypsin-EDTA covers the entire cell sheet or monolayer of DPSCs.
3. Keep the flask inside the hood at 37 °C for 3-5 min. After 3 min, gently tap the flask to ensure the dissociation of cells.
4. Add 5 mL of complete media (α-MEM with 10% FBS) or 250 µL of FBS (10% of the trypsin volume used) as a neutralizing agent to inactivate the trypsin activity. Transfer the whole cell suspension of dissociated DPSCs in a fresh 15 mL conical centrifuge tube.
5. Centrifuge the dissociated cells at 500 x g for 2 min at room temperature to get the cell pellet.
6. Aspirate off all supernatant using the pipette, add 5-10 mL of PBS to the flask to collect remnant cells, wash down the cell in PBS with gentle pipetting, and centrifuge the cells again at 500 x g for 2 min. Repeat if needed.
7. Resuspend the cell pellet in 1 mL of complete media (α-MEM with 10% FBS) after carefully removing the supernatant and performing gentle pipetting to obtain a single-cell suspension.
8. Split the cells and seed them (equal volume) into two T75 culture flasks, each containing 10 mL of full complete media. Keep the flask inside the hood at 37 °C.
9. The next day, aspirate off the media and replace it with the fresh complete media.
   NOTE: Prolong trypsin treatment can compromise cell integrity, viability, and cell function by cleaving the cell surface protein and extracellular matrix protein.

4. Identification of stem cell phenotypic markers (time: 90 - 120 min)

NOTE: For the characterization of cells harvested from the pulp tissue, use cells between the third and fifth passage.

1. Use two 70%-80% (~3-4 x 10⁶ cells; count the cells using an image-based cytometer or hemocytometer) confluent T75 culture flasks of DPSCs and wash the cells twice with 10 mL of PBS.
2. Add 2.5 mL of 0.25% trypsin-EDTA and keep the flask inside the hood at 37 °C for 3-5 min. Add 5 mL of complete media or 250 µL of FBS (10% of the trypsin volume used) as a neutralizing agent to inactivate the trypsin activity.
3. Collect the cells using a pipette in a 15 mL conical centrifuge tube. Centrifuge the dissociated cells at 500 x g for 2 min at room temperature to get the cell pellet.
4. Aspirate off all supernatant using the pipette, add 5 mL of PBS to the flask to collect remnant cells, wash down the cell in PBS with gentle pipetting, and centrifuge the cells again at 500 x g for 2 min.
5. Resuspend the pellet in 1 mL of PBS after carefully removing the supernatant and perform gentle pipetting to obtain a single-cell suspension.
6. Count the cells using an image-based cytometer or hemocytometer, depending on the availability, and distribute them in microcentrifuge tubes at 1 x 10⁴ cells/tube. Incubate the cells in 0.5%-1% bovine serum albumin (BSA) for 0.5-1 h.
7. To each tube, add 1-2 µL of fluorescein isothiocyanate (FITC)- and phycoerythrin (PE)-conjugated antibodies for cluster of differentiation (CD) markers (i.e., CD90 [0.4 µg], CD73 [0.8 µg], CD105 [0.4 µg], CD34 [0.8 µg], CD45 [0.4 µg], and HLA-DR [0.08 µg])^6,7,8,9 and incubate for 0.5 h in the dark at room temperature.
8. After 0.5 h, pellet down the cells by centrifuging at 500 x g for 5 min. Aspirate the supernatant, wash down the pellet of cells with PBS, and centrifuge again.
9. After centrifugation, resuspend the pellet in PBS (200 µL/tube), transfer the cell suspension to flow cytometry tubes, and acquire them using a flow cytometer. Any commercial software can be used to analyze the percentage of positive cells further for various antibodies.
   ​NOTE: Steps 4.7 to 4.9 are to be performed in the dark at room temperature. Proper isotype controls and unstained samples should be used to omit the background fluorescence of the cells.

5. DPSC differentiation into multiple lineages

NOTE: Use 75%-80% confluent cultures of DPSCs at the third to fifth passage for the assessment of multipotency. A control cell group containing α-MEM should be used for all types of differentiations described below.

1. Osteogenic differentiation of DPSCs, alizarin red staining, and quantification
   1. Make a single-cell suspension using 0.25% trypsin-EDTA, as described above in step 4.2, and resuspend the cell pellet in complete α-MEM.
   2. Count the cells and seed them in 24-well tissue culture-treated plates at a 2 x 10⁴ cells/well cell density containing 0.4-0.5 mL of complete media for 48 h at 37 °C and 5% CO₂.
   3. After 48 h, remove the media from the wells and add 0.4-0.5 mL of osteogenic inductive media (i.e., α-MEM with osteogenic inductive factors such as beta glycerophosphate [5 mM], monopotassium phosphate [1.8 mM], and dexamethasone disodium phosphate [0.01 mM])⁸. Prepare control wells (uninduced cells) containing complete α-MEM only without any osteogenic inductive factors (control media).
      NOTE: The Addition of 50 µM ascorbate is also reported to enhance osteogenic differentiation ^30,31. After every third day, replace the induction media and control media with fresh media and culture for 21 days (3 weeks). At the end of induction period, remove the media from the culture plates, add 0.5 mL of PBS, and wash two times.
   4. After PBS washing, fix the cells in 0.5 mL of 10% neutral buffered formalin (NBF) solution for 20-30 min at room temperature. Perform fixation in a fume/chemical hood.
2. Alizarin red staining
   1. Prepare the alizarin red S stain (2%) by dissolving 1 g of alizarin red S in 50 mL of double distilled water (ddH₂O) and adjusting the pH to 4.0-4.2 with the help of hydrochloric acid (HCL) or ammonium hydroxide (NH₄OH). Filter the stain with filter paper. Store the stain in the dark (4 °C).
   2. Wash the cells twice with double distilled water (ddH₂O), add alizarin red S stain, and incubate at room temperature for 30-45 min.
   3. Remove the stain from all the wells and wash the cells four or five times with ddH₂O to remove any unbound stain. Capture the images using a bright field microscope.
3. Alizarin red stain extraction for quantitative analysis
   1. Add cetyl pyridinium chloride (CPC) buffer (pH 7.0) to the stained wells (0.4 mL/well) and incubate the cells for 2 h at 37 °C. Additionally, the cells can also be incubated with 10% acetic acid for the proper dissolution of alizarin red crystals³². Keep the plates on a plate rocker or shaker.
   2. After the incubation, collect the supernatant (extracted dye), transfer it to a 96-well plate, and read the plate or samples at 405 nm. To obtain a blank value, add CPC (same volume) buffer only and read^32,33 .

6. Adipogenic differentiation of DPSCs, oil red O staining, and quantification

NOTE: The initial steps of seeding are the same as above (i.e., osteogenic differentiation step 5.1).

1. After 48 h, remove the media from the wells and add 0.4-0.5 mL of adipogenic inductive media (i.e., α-MEM with adipogenic inductive factors such as isobutyl-methylxanthine [IBMX; 0.5 mM], dexamethasone [1 µM], indomethacin [200 µM], and insulin [10 µM])⁸. Prepare control wells containing uninduced cells (i.e., cultured in α-MEM only without inductive factors {control media]).
2. Replenish the media every third day and culture the cells for 3 weeks. After 3 weeks, aspirate the media from the culture plates and add 0.5 mL of PBS to wash the cells twice.
3. After PBS washing, fix the cells in 10% NBF solution for 20-30 min at room temperature.
4. Wash the cells twice with double distilled water (ddH₂O), add oil red O stain, and incubate at room temperature for 30-40 min.
5. Remove the stain from all the wells and rinse the wells containing cells repeatedly with ddH₂O until clear (two to four times). Observe under the microscope and capture images.
6. Oil red O extraction for quantitative analysis
   1. Add 0.4 mL of 100% 2-propanol to the stained wells and incubate at room temperature by properly covering the plate for 30 min. Keep the plate on a plate rocker or shaker.
   2. After the incubation, collect the supernatant, transfer it to 96-well plate, read the plate or samples at 510 nm. Use 100% 2-propanol for blanking.

7. Chondrogenic differentiation of DPSCs, Alcian blue staining, and quantification

NOTE: Chondrogenic differentiation was induced in the monolayer culture of DPSCs. The initial steps of seeding are the same as above (i.e., osteogenic differentiation step 5.1).

1. After 48 h, remove the media from the wells and add 0.4-0.5 mL of chondrogenic induction media (i.e., α-MEM with inductive factors specific for chondrogenic differentiation, ascorbic acid [0.2 M], sodium pyruvate [1 mM], transforming growth factor-beta 3 [TGF-β₃; 10 µM], dexamethasone [1 mM], and 1x ITS premix [insulin, transferrin, and selenous acid)¹⁰. Prepare control wells containing uninduced cells (i.e., cultured in α-MEM only without inductive factors [control media]).
2. Replenish the media every third day and culture the cells for 3 weeks. After the end of the induction period, aspirate the media from the culture plates and add 0.5 mL of PBS to wash the cells twice.
3. After PBS washing, fix the cells in 10% NBF solution for 15-20 min at room temperature. Wash the cells two or three times with double distilled water (ddH₂O), add Alcian blue stain, and incubate at room temperature for 45 min.
4. Remove the stain from all the wells, followed by rinsing with 0.1 N HCL two or three times. Wash the cells four or give times with ddH₂O.
5. Observe the cells for proteoglycans (blue color) under the microscope and capture the images.
6. For Alcian blue quantification, extract the dye using 6 M guanidine-HCl and read the samples at an optical density (OD) of 595 nm. The Alcian blue kit used in this study stains acidic proteoglycans in cartilage and specially binds sulfated glycosaminoglycans (GAGs) present in the matrix of chondrocytes¹⁰.

8. Differentiation of DPSCs toward hepatic-like lineage and characterization

1. Make a single-cell suspension of cells by trypsin treatment, as described above, and resuspend the cell pellet in complete α-MEM.
2. Count the cells and seed them in 24-well-treated culture plates at 2 x 10⁴ cells/well containing 0.4-0.5 mL of complete media for 48 h.
3. After 48 h, remove the media from the wells and incubate the cells with hepatic induction medium for 14 days. The induction media consists of low glucose Dulbecco's modified eagle medium (DMEM) containing dexamethasone (0.5 µM), epidermal growth factor (EGF; 2 ng/mL), ITS+ premix (50 mg/mL), and hepatocyte growth factor (HGF; 20 ng/mL) ⁶.
4. After 14 days, replace the media with maturation medium for another 14 days (α-MEM supplemented with Oncostatin M [OSM; 20 ng/mL], ITS premix [50 mg/mL], and dexamethasone [0.5 µM]) ⁶. Replenish the media with fresh media after every third day.
5. Perform immunohistochemistry analysis and a low-density lipoprotein (LDL) assay for confirming hepatic-like differentiation.
   1. Uptake of LDL and immunofluorescent staining for the LDL receptor
      NOTE: Hepatocyte-like cells post-differentiation were characterized by the LDL uptake assay kit as per the kit instructions. LDL-550 fluorochrome was used to assess the LDL uptake in differentiated cells under the live state and the LDL receptor antibody was used for the detection of LDL receptors in differentiated cells post-fixation.
      1. After differentiation, stain the live cells with LDL-550 (1:100) for 3-4 h. After 3-4 h, replace the staining solution with culture media and observe the LDL uptake under a fluorescent microscope capable of propidium iodide (PI) excitation and emission wavelengths at 537 and 617 nm, respectively.
      2. Fix the cells in the fixative solution for 10 min. After cell fixation, block the cells in the blocking solution for 30 min.
      3. Incubate the cells with a primary LDL receptor antibody for 1 h, wash with 1x PBS three times, and then incubate with 488 secondary antibody (1:100) for 1 h in dark.
      4. Wash the cells with PBS three times to remove excess secondary antibody. Add 4′,6-diamidino-2-phenylindole (DAPI) as a nuclear stain.
      5. Observe the cells under a fluorescence microscope. Take images using a fluorescent microscope capable of measuring wavelengths in excitation/emission range of DAPI, PI, and FITC.
         ​NOTE: Single-cell intensity parameters can be recorded to quantify the LDL receptor antibody expression and LDL uptake. Proper secondary antibody controls should be used for immunofluorescence staining to omit background fluorescence resulting from the secondary antibody only.

9. Neural differentiation of DPSCs toward neuronal-like lineage and characterization

1. After the plating of the cells, as described above, culture the cells in α-MEM for 48 h.
2. Aspirate the media and replace it with neurobasal media containing growth factors such as 0.5% G5 supplement, 0.2% N2 supplement, 20 ng/mL epidermal growth factor, 20 ng/mL basic fibroblast growth factor, and B27 supplement (1%). Culture the cells in this media for 25-41 days⁷.
3. Change the media every second or third day, depending on its metabolism by the cells. To attain efficient neuronal maturation and more microtubule-associated protein-2 (MAP-2+) neurons, remove EGF and fibroblast growth factor 2 (FGF2) after the neuronal induction stage (14 days).
   NOTE: Over time and after repeated media change, differentiated neurons might show detachment from the plate wells due to low adherence, resulting in neuron loss in long-term culture. To avoid this, coat the plate wells with laminin or fibronectin (optional).
4. Perform immunohistochemistry analysis for confirming neuronal-like differentiation.
   1. Immunostaining of differentiated DPSCs
      NOTE: To confirm the neuronal-like differentiation of DPSCs, immunocytochemistry was performed in the differentiated cells using MAP-2 and neurofilament (NFM) antibodies, which are neurogenic lineage-specific proteins⁷.
      1. Remove the culture media after differentiation is complete. Add PBS, gently swirl the plate, and remove the PBS with the help of a pipette. Repeat the step.
      2. Fix the cells using 10% NBF or 4% paraformaldehyde for 20-30 min at room temperature.
      3. Remove the fixative and wash the cells with 1x PBS two or three times. Add permeabilization solution (Triton-X-100; 0.1%) and incubate the cells for 5 min.
      4. Perform PBS wash two or three times and block the cells with 0.5%-1% blocking agent (BSA) for 1 h at 37 °C or overnight at 4 °C. Remove the blocking agent and wash the cells with 1x PBS.
      5. Incubate the cells overnight with primary antibody (prepared in 0.1% BSA, NFM [1:50], and MAP-2 [1:200]).
      6. Remove the primary antibody and wash the cells with 1x PBS three or four times to remove unbound antibodies.
      7. Add secondary antibody (rabbit anti-mouse 1:2,000) and incubate the cells at room temperature for 1.5-2 h in the dark. Wash the cells with PBS (two or three times).
      8. Incubate the cells in DAPI for 10 min and wash again with PBS. Keep the residual volume to avoid drying of the sample. Observe under the fluorescence microscope.
         NOTE: Additionally, karyotype stability¹⁰ and senescence characterization can also be performed to get a better idea of the purity and functionality of the cells.

Results

Here, we describe how researchers can establish a pure culture of DPSCs via the explant method^6,7,8,9,10 and induce them toward multiple lineages to establish the purity of culture for downstream applications.

We established a primary culture of DPSCs from the small tissue of pulp extracted from the third molar tooth of pat...

Discussion

Stem cells have pinned the hopes of curing numerous diseases, owing to their plasticity, robustness, immunomodulatory properties, paracrine mechanisms, and homing efficiencies. Dental pulp tissue is considered the most potent and valuable source of stem cells, with eminent plasticity and a regenerative capability. Here, we demonstrate the isolation of DPSCs, utilizing the widely adopted explant culture method, in which the cells migrate from pieces of pulp tissue or explants to grow into a homogenous cell culture that mo...

Materials

Name	Company	Catalog Number	Comments
6 well cell culture plate	Costar	3516	For cell culture
Alcian blue stain	EZstain chondrocyte staining kit, HiMedia	CCK029
alizarin red S stain	Sigma-Aldrich	TMS-008	Osteogenic stain
Antibiotic cocktail	Himedia	A002-5X50ML	To prevent culture contamination
Ascorbic Acid	Himedia	TC094-25G	Chondrogenic induction
B27 supplement	Gibco	17504044	For neural induction
bFGF ( basic Fibroblast Growth Factor)	Gibco	PHG0024	For neural induction
CD 105	BD-Pharmingen	560839
CD 35	Biolegend	343604
CD 45	Biolegend	304006
CD 73	Biolegend	344016
CD 90	Biolegend	328107	Characterization
cetyl pyridinium chloride (CPC)	Sigma-Aldrich	1104006	For Alizarin Red extraction
Dexamethasone 21-phosphate disodium	Sigma-Aldrich	D1159-100MG
Dulbecco's Phosphate Buffered Saline	Himedia	TS1006-5L	For washing purpose
EGF (Epidermal Growth Factor)	Gibco	PHG0311	For hepatic and neural  induction
EVOS LED microscope	Invitrogen		For  fluorescence imaging
EZ stain Chondrocyte staining kit	Himedia	CCK029-1KT	Chondro stain Kit
FACS Canto flow cytometer	BD Biosciences		For cell characterization
Fetal Bovine Serum	Gibco	16000044	For primary culture
Fetal Bovine Serum	Sigma-Aldrich	F2442	For cell culture
G5 supplement	Gibco	17503012	For neural induction
HGF( Hepatocyte Growth Factor)	Sigma-Aldrich	H1404	For hepatic Induction
HLA-DR	Biolegend	307605
Human TGF-β3	Peprotech	#100-36E-10U
Insulin-Transferrin-Selenous acid premix	Sigma-Aldrich	I3146	For hepatic Induction
ITS premix	Corning	354350
LDL Uptake Assay kit	Abcam	ab133127	For hepatic characterization
Low glucose DMEM	Gibco	11885-084	For hepatic induction
MAP2 antibody	Sigma-Aldrich	M4403	For neural characterization
N2 supplement	Gibco	17502048	For neural induction
Neural Basal Media	Gibco	21103049	For neural induction
NFM antibody	Sigma-Aldrich	N4142	For neural characterization
Nikon Elipse TS100 microscope	Nikon		For  fluorescence imaging
Oil Red O	Sigma-Aldrich	01391-250Ml	Adipogenic stain
Oncostatin M	R&D Systems	295-OM-010/CF	For hepatic Induction
Petridish	Tarson	460090-90MM	For tissue cutting
Potassium phosphate monobasic	Sigma-Aldrich	15655-100G	Osteogenic induction
Propan-2-ol	Thermo Fisher	Q13827	For Oil Red O extraction
Sodium pyruvate solution	Sigma life sciences	S8636-100ML
Trypsin-EDTA	Sigma-Aldrich	T4049	For cell passaging
Whatman filter paper	merck	WHA1001325	filter paper
α- Minimum Essential Media (α-MEM)	Sigma-Aldrich	M0643-10X 1L	Media for primary culture
β-glycerophosphate disodium salt hydrate	Sigma-Aldrich	G9422-50G