Accepted Manuscript Title: Effects of Mesenchymal Stem Cell-Derived Cytokines on the Functional Properties of Endothelial Progenitor Cells Author: Witchayaporn Kamprom Pakpoom Kheolamai Yaowalak U-Pratya Aungkura Supokawej Methichit Wattanapanitch Chuti Laowtammathron Surapol Issaragrisil PII: DOI: Reference:
S0171-9335(16)30004-8 http://dx.doi.org/doi:10.1016/j.ejcb.2016.02.001 EJCB 50866
To appear in: Received date: Revised date: Accepted date:
27-8-2015 28-12-2015 3-2-2016
Please cite this article as: Kamprom, W., Kheolamai, P., U-Pratya, Y., Supokawej, A., Wattanapanitch, M., Laowtammathron, C., Issaragrisil, S.,Effects of Mesenchymal Stem Cell-Derived Cytokines on the Functional Properties of Endothelial Progenitor Cells, European Journal of Cell Biology (2016), http://dx.doi.org/10.1016/j.ejcb.2016.02.001 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
Effects of Mesenchymal Stem Cell-Derived Cytokines on the Functional Properties of Endothelial Progenitor Cells Witchayaporn Kamproma,b, Pakpoom Kheolamaib,c,d, Yaowalak U-Pratyab,e, Aungkura Supokawejf, Methichit Wattanapanitchb,g, Chuti Laowtammathronb, Surapol Issaragrisilb,e a
an
us
cr
ip t
Department of Immunology, Faculty of Medicine Siriraj hospital, Mahidol University, Bangkok, Thailand. b Siriraj Center of Excellence for Stem Cell Research, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand. c Division of Cell Biology, Faculty of Medicine, Thammasat University, Pathumthani, Thailand. d Center of Excellence in Stem Cell Research, Faculty of Medicine, Thammasat University, Pathumthani, Thailand. e Division of Hematology, Department of Medicine, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand. f Faculty of Medical Technology, Mahidol University, Bangkok, Thailand. g Department of Research and Development, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand.
*
d
M
[email protected], Division of Hematology, Department of Medicine, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok 10700, Thailand.; Tel.: +662 419 4448 50; fax: +662 411 2012
Ac ce pt e
Abstract Human mesenchymal stem cell (hMSC) is a potential source for cell therapy due to its property to promote tissue repair. Although, it has been known that hMSCs promote tissue repair via angiogenic cytokines, the interaction between hMSC-derived cytokines and the endothelial progenitor cells (EPCs), which play an important role in tissue neovascularization, is poorly characterized. We investigate the effect of cytokine released from different sources of hMSCs including bone marrow and gestational tissues on the EPC functions in vitro. The migration, extracellular matrix invasion and vessel formation of EPCs were studied in the presence or absence of cytokine released from various sources of hMSCs using transwell culture system. The migration of EPCs was highest when co-culture with secretory factors from placenta-derived hMSCs (PL-hMSCs) compared to those co-culture with other sources of hMSCs. For invasion and vessel formation, secretory factors from bone marrow-derived hMSCs (BM-hMSCs) could produce the maximal enhancement compared to other sources. We further identified the secreted cytokines and found that the migratory-enhancing cytokine from PL-hMSCs was PDGF-BB while the enhancing cytokine from BM-hMSCs on invasion was IGF-1. For vessel formation, the cytokines released from BM-hMSCs were IGF1 and SDF-1. In conclusion, hMSCs can release angiogenic cytokines which increase the migration, invasion and vessel forming capacity of EPCs. We can then use hMSCs as a source of angiogenic cytokines to induce neovascularization in injured/ischemic tissues. Keywords Mesenchymal stem cell, Neovascularization, Gestational tissues Introduction
Page 1 of 21
Ac ce pt e
d
M
an
us
cr
ip t
Human mesenchymal stem cells (hMSCs) are multipotent stem/progenitor cells which are present in bone marrow, adipose tissue, and postnatal sources including umbilical cord, placenta, the Wharton’s jelly, and amnion (Bunnell et al., 2008; In 't Anker et al., 2004; Lee et al., 2004; Wang et al., 2004). hMSCs are considered to be a potential source for cellular therapy in several disorders such as acute graft-versus-host disease (acute GvHD), myocardial infarction, and neurological diseases (Uccelli et al., 2008; Williams et al., 2013) due to immunomodulatory property and their multilineage differentiation capacity (Jiang et al., 2002; Nauta and Fibbe, 2007; Pittenger et al., 1999). The therapeutic effects of hMSCs are believed due to soluble factors or cytokine released from hMSCs rather than their direct effect (Meirelles Lda et al., 2009). hMSC-derived soluble factors can exert immunomodulatory effect, reduce tissue inflammation and prevent apoptosis of several cell types in injured tissues which might be the underlying mechanism in the improvement of tissue repair and inflammatory disease (Gnecchi et al., 2008; Wang et al., 2014). hMSCs can stimulate neovascularization in the injured ischemic tissue via pro-angiogenic cytokines which enhance endothelial cell proliferation and migration leading to neovascularization and tissue restoration (Hung et al., 2007; Kamihata et al., 2001; Li et al., 2010). It was the belief that mature endothelial cells in the local vasculature are responsible for postnatal neovascularization. However, recent studies have shown the important role of endothelial progenitor cells (EPCs) in vascular homeostasis and postnatal neovascularization in both physiological and pathological conditions (Asahara et al., 1999; Melero-Martin et al., 2007; Tepper et al., 2005). There are no reported information on the effects of cytokines released from hMSCs on the endothelial progenitor cells. In order to better understand the therapeutic effect of hMSCs on neovascularization, we investigated the effects of soluble factors released from hMSCs on the EPC functions including proliferation, migration, invasion and vessel formation capacity in vitro, and also identified those factors that play roles on various EPC functions. Soluble factors released from gestational tissues-derived hMSCs were chosen to study due to easily and non-invasive collection compared to bone marrow-derived hMSCs (BM-hMSCs).
Materials and methods Subjects This study was approved by the Siriraj Institutional Review Board, Faculty of Medicine Siriraj Hospital, Mahidol University which was in accordance with the Declaration of Helsinki, the Belmont Report, CIOMS Guidelines, and ICH-GCP. Human bone marrow samples were obtained from healthy volunteers after giving written informed consent. The gestational tissues (umbilical cord, Wharton’s jelly, placenta, and amnion) were obtained from healthy newborns after receiving written informed consent from their mothers. Isolation and culture of hMSCs Ten milliliter of heparinized bone marrow was collected for hMSC isolation. Bone marrowderived mononuclear cells were then isolated using IsoPrep® (Robbins Scientific Corporation, USA) density gradient centrifugation, washed twice with PBS (GIBCO™, Invitrogen Corporation, USA), and re-suspended in complete medium which is Dulbecco’s Modified Eagle Medium (DMEM) (GIBCO™, Invitrogen Corporation, USA) supplemented with 10% (v/v) Fetal Bovine Serum (FBS) (Lonza, USA), 100 U/ml penicillin (General Drug House CO., Ltd, Thailand), and 100 µg/ml streptomycin (General Drug House CO., Ltd, Thailand). Cell suspensions were then plated in 25 cm2 culture flask (Corning, USA) at a density of 2×105 cells/cm2. Cultures were maintained at 37°C in a humidified atmosphere
Page 2 of 21
containing 5% CO2 and the medium was replaced every 3 days throughout the entire culture period.
ip t
For hMSC isolation from gestational tissues, umbilical cord, Wharton’s jelly, placenta, and amnion were cut into small pieces and digested by incubation with 0.25% (w/v) trypsinEDTA (GIBCO™, Invitrogen Corporation, USA) for 30 minutes at 37°C. Cell suspensions were washed twice with PBS, re-suspended in complete medium and plated in 25 cm2 culture flask (Corning, USA). Cultures were maintained at 37°C in a humidified atmosphere containing 5% CO2 and the medium was replaced every 3 days throughout the entire culture period.
cr
Characterization of cultured hMSCs by flow cytometry
an
us
The 3rd-5th passage of hMSCs were characterized for hMSC surface markers by incubation with the following mouse anti-human antibodies: anti-CD45-PerCP (BD Pharmingen, USA), anti-CD34-PE (Biolegend, USA), anti-CD90-FITC (AbD Serotec, USA), anti-CD73-PE (BD Pharmingen, USA), and anti-CD105-PE (Miltenyi Biotec, Germany) for 30 minutes at 4°C in the dark. After incubation with the antibodies, cell pellets were washed twice with PBS and fixed with 1% (w/v) paraformaldehyde in PBS. Flow cytometry was performed by FACS calibur™ Flow cytometer using CellQuest™ software (Becton Dickinson, USA).
Ac ce pt e
d
M
Osteogenic and adipogenic differentiation of cultured hMSCs The 3rd-5th passage hMSCs were used to assess their adipogenic and osteogenic differentiation potentials. For adipogenic differentiation, 5×104 cells were cultured in NH AdipoDiff® Medium (Miltenyi Biotec, Germany). Medium was replaced every 3 days according to the manufacturer’s instruction. After culture for 3 weeks, cells were stained with 0.5% (w/v) Oil Red O (Sigma Aldrich, USA), in isopropanol for 20 minutes at room temperature, to determine the number of hMSC-derived adipocytes in culture. For osteogenic differentiation, 5×104 cells were cultured in NH OsteoDiff® Medium (Miltenyi Biotec, Germany). Medium was replaced every 3 days according to the manufacturer’s instruction. After culture for 3 weeks, cells were stained with 40 mM Alizarin Red S (Sigma Aldrich, USA) for 20 minutes at room temperature to determine the number of hMSC-derived osteocytes in culture. Cultures and characterization of EPCs from umbilical cord blood Twenty milliliter of heparinized umbilical cord blood was collected for EPC isolation. Umbilical cord blood-derived mononuclear cells were then isolated using IsoPrep® (Robbins Scientific Corporation, USA) density gradient centrifugation, washed twice with PBS (GIBCO™, Invitrogen Corporation, USA), re-suspended in endothelial cell growth medium [endothelial basal medium-2 (LONZA, Germany), supplemented with EGM-2 single aliquots (LONZA, Germany) containing 2% (v/v) fetal bovine serum (FBS), 5 µg/ml epidermal growth factor, 200 µg/ml hydrocortisone, 0.5 µg/ml vascular endothelial growth factor, 10 µg/ml basic fibroblast growth factor, 20 µg/ml long R3 Insulin-like growth factor 1 and 1 mg/ml ascorbic acid], and plated in an individual well of 6-well plate coated with 10 µg/ml human fibronectin (Amersham Biosciences, USA) at a density of 1×106 cells/well. After culture for 3 days, the non-adherent cells were removed and fresh medium was added. Cultures were maintained at 37°C in a humidified atmosphere containing 5% CO2 and medium was replaced every 3 days throughout the entire culture period.
Page 3 of 21
Study of hMSC-derived cytokines on the functional properties of EPCs
ip t
The cultured cells were characterized for EPC surface markers by incubation with the following mouse anti-human antibodies: anti-CD34-PE (R&D Systems, USA), antiVEGFR2-PE (R&D Systems, USA), anti-CD146-FITC (R&D Systems, USA), and anti-vWFFITC (R&D Systems, USA) for 15 minutes at 4ºC in the dark. Cell pellets were then washed twice with PBS and fixed with 1% (v/v) paraformaldehyde in PBS. Flow cytometry was performed by FACScaliburTM flow cytometer (Becton Dickinson, USA) using CellQuest® software.
Ac ce pt e
d
M
an
us
cr
Effect on proliferation 2×105 hMSCs derived from five distinct tissues were cultured in 2 ml DMEM supplemented with 2% (v/v) FBS for 48 hours to generate hMSC conditioned media for proliferation assay. Briefly, 2.5×104 EPCs were cultured in an individual well of culture E-Plate (Roche Applied Science, USA) containing 100 µl hMSC conditioned media for 7 days. The number of EPCs in each well was continuously monitored throughout the entire culture period by xCELLigence Real-Time Cell Analyzer (Roche Applied Science, USA) and reported as cell index (CI). EPCs cultured in DMEM containing 2% (v/v) FBS served as controls. Effect on migration and invasion Five distinct sources of hMSCs were co-cultured with EPCs through 8 µm transwell inserts (Corning, USA) to assess the effect of hMSC-derived cytokines on the migratory and invasive capacity of EPCs. For migration assay, hMSCs were seeded in the lower chamber containing 600 µl DMEM supplemented with 2% (v/v) FBS at the density of 2.5×104 cells/cm2, while 4×104 EPCs were seeded in the transwell inserts. After 6 hours of co-culture, numbers of EPCs that migrate to the other side of the transwell’s membrane were determined by hematoxylin staining. The procedure of invasion assay was similar to that of migration assay, except that the membrane of the transwell inserts were pre-coated with 0.4 µg/µl Matrigel (BD Biosciences, USA) to serve as an extracellular matrix barrier. The numbers of invasive EPCs which are able to degrade coated Matrigel and migrate to the other side of the transwell’s membrane were determined by hematoxylin staining. Effect on vessel-forming capacity Five distinct sources of hMSCs were co-cultured with EPCs through 0.4 µm transwell inserts (Corning, USA) to assess the effect of hMSC-derived cytokines on the vessel-forming capacity of EPCs using an in vitro vessel formation assay as previously described (Jiraritthamrong et al., 2012). Briefly, 3.5×104 EPCs labeled with 5-(6)-carboxyfluorescein diacetate succinimidyl ester (CFSE, CellTrace™; Invitrogen, USA) were mixed with an equal number of non-labeled human umbilical vein endothelial cells (HUVECs) and plated into an individual well of 6-well plate pre-coated with Matrigel™ (BD Bioscience, USA) while hMSCs were seeded in the transwell inserts at the density of 1×104 cells/cm2. After 12 hours of co-culture, the extent of incorporated EPCs in the generated capillary-like structure was determined by fluorescent microscopy (Nikon, Japan) using NIS-Elements D software (Nikon, Japan). Expression levels of angiogenic-related genes in hMSCs by quantitative real-time PCR (qRT-PCR) Total RNA were isolated from hMSCs using TRIzol® reagent (Invitrogen Corporation, USA). cDNA was synthesized from 2 µg of RNA using the SuperScript™ III Reverse Trancriptase
Page 4 of 21
ip t
(Invitrogen Corporation, USA). MicroAmp® fast optical 96-well reaction plate (Applied Biosystem, USA) was used for qRT-PCR. Each well contained 3 µl cDNA, 1 µl 10 µM forward and reverse primer mix, and 10 µl SYBR® Green PCR Mastermix (Applied Biosystem, USA). The plate was then sealed with MicroAmp® clear adhesive film (Applied Biosystem, USA) to prevent evaporation of the reactant. PCR was performed using 7500 Fast Real-time PCR system (Applied Biosystem, USA) using the following protocol: 95°C initial denaturation for 10 minutes, followed by 40 cycles of denaturation (95°C, 10 seconds), annealing (60°C, 10 seconds), and extension (72°C, 40 seconds). The relative quantity of a target gene was calculated by normalization with glyceraldehyde-3-phosphate dehydrogenase (GAPDH), using the 7500 software version 2.0.5 (Applied Biosystem, USA). The primer sequences were described in Table 1.
cr
Table 1: The sequences of primers for qRT-PCR Sequence of forward primer 5ʹ-TCCTCACACCATTGAAACCA-3ʹ 5ʹ-AGGAACCAGCCTCCTCTCTC-3ʹ 5ʹ-AACCAGACGGCTGTGATGAT-3ʹ 5ʹ-AGCGGCTGTACTGCAAAAAC-3ʹ 5ʹ-GTGGTCGTGCTGGTCCTC-3ʹ
Sequence of reverse primer 5ʹ-GATCCTGCCCTGTCTCTCTG-3ʹ 5ʹ-TTCTCCAGCAGCTGTATCTCAA-3ʹ 5ʹ-TTGTCGAGAGGGAGTGTTCC-3ʹ 5ʹ-AGCCAGGTAACGGTTAGCAC-3ʹ 5ʹ-ATCTGAAGGGCACAGTTTGG-3ʹ
IGF1
5ʹ-CCGGAGCTGTGATCTAAGGA3ʹ
5ʹ-CCTGCACTCCCTCTACTTGC-3ʹ
PlGF
5ʹ-GTTCAGCCCATCCTGTGTCT-3ʹ
5ʹ-AACGTGCTGAGAGAACGTCA3ʹ
IGF2
5ʹ-TGCTGGTGCTTCTCACCTTC-3ʹ
5ʹ-AGACGAACTGGAGGGTGTCC-3ʹ
IL6
5ʹ-CAGGAGCCCAGCTATGAACT3ʹ
5ʹ-GTGAGTGGCTGTCTGTGTGG3ʹ
IL8 TGFβ
5ʹ-AAGAAACCACCGGAAGGAAC-3ʹ 5ʹ-GAGCCTGAGGCCGACTACTA-3ʹ
5ʹ-AAATTTGGGGTGGAAAGGTT-3ʹ 5ʹ-CACGTGCTGCTCCACTTTTA-3ʹ
PDGFβ
5ʹ-CCGGAGTCGGCATGAATC-3ʹ
5ʹCGTTGGAGATCATCAAAGGAG-3ʹ
GAPDH
5ʹ-GTCAACGGATTTGGTCGTATTG-3ʹ
5ʹ-CATGGGTGGAATCATATTGGAA-3ʹ
Ac ce pt e
d
M
an
us
Gene name VEGFA ANGPT1 ANGPT2 FGF2 SDF1
Study of PDGF-BB on EPC migration
4×104 EPCs were seeded into 8 µm transwell inserts that were placed into an individual well of 24-well plates containing 600 µl DMEM supplemented with 2% (v/v) FBS and various concentrations (0, 20, 50, and 100 ng/ml) of platelet-derived growth factor-BB (PDGF-BB; R&D system, USA). After 6 hours of culture, the effect of PDGF-BB on EPC migration was determined by migration assay as previously described. To further determine the role of PDGF-β on EPC migration, 4×104 EPCs were treated with 10 µg/ml anti-PDGFR-β neutralizing antibody (R&D system, USA) for 30 minutes to inhibit the action of PDGF-BB before co-cultured with hMSCs through 8 µm transwell. After 6 hours of culture, the migration assay was performed as previously described. Study of SDF1, IGF1 and PlGF on EPC invasion 4×104 EPCs were seeded into 8 µm Matrigel-coated transwell inserts that were placed into an individual well of 24-well plates containing 600 µl DMEM with 2% (v/v) FBS which were supplemented with either 100 ng/ml stromal derived factor1 (SDF1) (R&D system, USA), 100 ng/ml insulin-like growth factor1 (IGF1; R&D system, USA), 100 ng/ml placenta growth
Page 5 of 21
factor (PlGF; R&D system, USA), or combination of those three factors (100 ng/ml each). After culture for 18 hours, the effect of those factors on capacity of EPC invasion was determined by invasion assay as previously described.
cr
Study of SDF1, IGF1 and PlGF on vessel-forming capacity of EPCs
ip t
To further determine the roles of SDF1, IGF1 and PlGF on EPC invasion, 4×104 EPCs were pre-treated with either 10 µM AMD3100 octahydrochloride (inhibitor of SDF1; Tocris Bioscience, UK), 10 µg/ml anti-hIGF1R (inhibitor of IGF1; R&D system, USA), 5 µg/ml anti-hVEGFR1 (inhibitor of PlGF; R&D Systems, USA), or the combination of all those three inhibitors for one hour before co-culture with hMSCs through 8 µm Matrigel-coated transwell. After culture for 18 hours, the invasion assay was performed as previously described.
an
us
3.5×104 CFSE-labeled EPCs were co-cultured with an equal number of non-labeled HUVECs in endothelial basal medium supplemented with either 100 ng/ml SDF1, 100 ng/ml IGF1, 100 ng/ml PlGF, or the combination of all those three factors (100 ng/ml each). The mixtures of cells were then seeded into an individual well of Matrigel-coated 24-well plate. After 12 hours of culture, an in vitro vessel formation assay was performed as previously described.
Statistical analysis
d
M
To further verify the roles of SDF1 and IGF1 on vessel-forming capacity of EPCs, 105 EPCs were treated with either 10 µM AMD3100 octahydrochloride (SDF1 inhibitor; Tocris Bioscience, UK) or 10 µg/ml anti-hIGF1R (IGF1 inhibitor; R&D system, USA) for one hour before co-culture with BM-hMSCs through transwell culture system. An in vitro vessel formation assay was performed as previously described.
Ac ce pt e
Data were presented as mean ± standard error of the mean (SEM). The paired Student’s t-test and one-way ANOVA were used to assess the significance of differences between observed data. P < 0.05 was considered to be statistically significant.
Results
Characteristics of hMSCs derived from gestational tissues and bone marrow hMSCs derived from gestational tissues, including placenta (PL-hMSCs), umbilical cord (UC-hMSCs), Wharton’s jelly (WJ-hMSCs) and amnion (AM-hMSCs) exhibited similar characteristics to those of bone marrow-derived hMSCs (BM-hMSCs). The gestational tissuederived hMSCs displayed fibroblast-like morphology (Fig. 1A-E), expressed typical hMSC surface markers (positive for CD73, CD90, CD105 and negative for hematopoietic markers CD34 and CD45; Fig. 1P). Moreover, the gestational tissue-derived hMSCs could differentiate toward adipocyte- and osteocyte-lineages as demonstrated by Oil-Red O (Fig. 1F-J) and Alizarin Red S staining (Fig. 1K-O), respectively. Expression of pro-angiogenic genes in gestational tissue and bone marrow-derived hMSCs The expression levels of pro-angiogenic genes among various sources of hMSCs were determined by quantitative real-time PCR. The expression levels of stromal cell-derived
Page 6 of 21
ip t
factor 1 (SDF1), insulin-like growth factor 1 (IGF1), and placental growth factor (PlGF) genes in the BM-hMSCs were significantly higher than those of other hMSC sources (Fig. 2). In contrast, the expression level of platelet-derived growth factor beta (PDGF-β) was significantly higher in PL-hMSCs than the rest of the hMSC sources. Apart from those 4 genes, the expression levels of several pro-angiogenic genes, including vascular endothelial growth factor A (VEGF-A), fibroblast growth factor 2 (FGF2), angiopoietin 1 (ANGPT1), angiopoietin 2 (ANGPT2), insulin-like growth factor 2 (IGF2), interleukin 6 (IL6), interleukin 8 (IL8), and transforming growth factor beta (TGF-β) were not significantly different among the various hMSC sources. Characteristics of umbilical cord blood-derived endothelial progenitor cells (EPCs)
us
cr
EPCs were cultured from mononuclear cell populations that had been separated from umbilical cord blood by density gradient centrifugation. After culture in endothelial growth medium for 7-10 days, several EPC-like colonies of cobblestone morphology were observed (Fig. 3A). These cells expressed typical EPC surface markers, including CD34, CD146, Von Willebrand factor (vWF) and vascular endothelial growth factor receptor 2 (VEGFR2) (Fig. 3B), and were able to form capillary-like structures on Matrigel™ (Fig. 3C).
an
Effects of hMSC-derived soluble factors on the functional properties of EPCs
Ac ce pt e
d
M
Effect of hMSC secreted factors on EPC proliferation. The conditioned medium derived from hMSCs from various sources were collected and used to culture EPCs. Similar to control group (cultured in DMEM supplemented with 2% [v/v] FBS), the numbers of EPCs cultured in various hMSC-conditioned media were increased initially during the first 48 hours of culture before rapidly decreasing toward the baseline level at the end of culture (Fig. 4), indicating that the soluble factors secreted from bone marrow and gestational tissue-derived hMSCs did not promote EPC proliferation. B) Effect of hMSC secreted factors on EPC migration. EPCs were co-cultured with hMSC from various sources using the transwell culture system (Fig. 5A). The numbers of EPCs that migrate toward various hMSC sources were determined after 6 hours of co-culture. Interestingly, only the soluble factors derived from PL-hMSCs significantly enhanced EPC migration in comparison to controls (306.2 ± 60% vs. 100% of control, P
S0171-9335(16)30004-8 http://dx.doi.org/doi:10.1016/j.ejcb.2016.02.001 EJCB 50866
To appear in: Received date: Revised date: Accepted date:
27-8-2015 28-12-2015 3-2-2016
Please cite this article as: Kamprom, W., Kheolamai, P., U-Pratya, Y., Supokawej, A., Wattanapanitch, M., Laowtammathron, C., Issaragrisil, S.,Effects of Mesenchymal Stem Cell-Derived Cytokines on the Functional Properties of Endothelial Progenitor Cells, European Journal of Cell Biology (2016), http://dx.doi.org/10.1016/j.ejcb.2016.02.001 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
Effects of Mesenchymal Stem Cell-Derived Cytokines on the Functional Properties of Endothelial Progenitor Cells Witchayaporn Kamproma,b, Pakpoom Kheolamaib,c,d, Yaowalak U-Pratyab,e, Aungkura Supokawejf, Methichit Wattanapanitchb,g, Chuti Laowtammathronb, Surapol Issaragrisilb,e a
an
us
cr
ip t
Department of Immunology, Faculty of Medicine Siriraj hospital, Mahidol University, Bangkok, Thailand. b Siriraj Center of Excellence for Stem Cell Research, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand. c Division of Cell Biology, Faculty of Medicine, Thammasat University, Pathumthani, Thailand. d Center of Excellence in Stem Cell Research, Faculty of Medicine, Thammasat University, Pathumthani, Thailand. e Division of Hematology, Department of Medicine, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand. f Faculty of Medical Technology, Mahidol University, Bangkok, Thailand. g Department of Research and Development, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand.
*
d
M
[email protected], Division of Hematology, Department of Medicine, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok 10700, Thailand.; Tel.: +662 419 4448 50; fax: +662 411 2012
Ac ce pt e
Abstract Human mesenchymal stem cell (hMSC) is a potential source for cell therapy due to its property to promote tissue repair. Although, it has been known that hMSCs promote tissue repair via angiogenic cytokines, the interaction between hMSC-derived cytokines and the endothelial progenitor cells (EPCs), which play an important role in tissue neovascularization, is poorly characterized. We investigate the effect of cytokine released from different sources of hMSCs including bone marrow and gestational tissues on the EPC functions in vitro. The migration, extracellular matrix invasion and vessel formation of EPCs were studied in the presence or absence of cytokine released from various sources of hMSCs using transwell culture system. The migration of EPCs was highest when co-culture with secretory factors from placenta-derived hMSCs (PL-hMSCs) compared to those co-culture with other sources of hMSCs. For invasion and vessel formation, secretory factors from bone marrow-derived hMSCs (BM-hMSCs) could produce the maximal enhancement compared to other sources. We further identified the secreted cytokines and found that the migratory-enhancing cytokine from PL-hMSCs was PDGF-BB while the enhancing cytokine from BM-hMSCs on invasion was IGF-1. For vessel formation, the cytokines released from BM-hMSCs were IGF1 and SDF-1. In conclusion, hMSCs can release angiogenic cytokines which increase the migration, invasion and vessel forming capacity of EPCs. We can then use hMSCs as a source of angiogenic cytokines to induce neovascularization in injured/ischemic tissues. Keywords Mesenchymal stem cell, Neovascularization, Gestational tissues Introduction
Page 1 of 21
Ac ce pt e
d
M
an
us
cr
ip t
Human mesenchymal stem cells (hMSCs) are multipotent stem/progenitor cells which are present in bone marrow, adipose tissue, and postnatal sources including umbilical cord, placenta, the Wharton’s jelly, and amnion (Bunnell et al., 2008; In 't Anker et al., 2004; Lee et al., 2004; Wang et al., 2004). hMSCs are considered to be a potential source for cellular therapy in several disorders such as acute graft-versus-host disease (acute GvHD), myocardial infarction, and neurological diseases (Uccelli et al., 2008; Williams et al., 2013) due to immunomodulatory property and their multilineage differentiation capacity (Jiang et al., 2002; Nauta and Fibbe, 2007; Pittenger et al., 1999). The therapeutic effects of hMSCs are believed due to soluble factors or cytokine released from hMSCs rather than their direct effect (Meirelles Lda et al., 2009). hMSC-derived soluble factors can exert immunomodulatory effect, reduce tissue inflammation and prevent apoptosis of several cell types in injured tissues which might be the underlying mechanism in the improvement of tissue repair and inflammatory disease (Gnecchi et al., 2008; Wang et al., 2014). hMSCs can stimulate neovascularization in the injured ischemic tissue via pro-angiogenic cytokines which enhance endothelial cell proliferation and migration leading to neovascularization and tissue restoration (Hung et al., 2007; Kamihata et al., 2001; Li et al., 2010). It was the belief that mature endothelial cells in the local vasculature are responsible for postnatal neovascularization. However, recent studies have shown the important role of endothelial progenitor cells (EPCs) in vascular homeostasis and postnatal neovascularization in both physiological and pathological conditions (Asahara et al., 1999; Melero-Martin et al., 2007; Tepper et al., 2005). There are no reported information on the effects of cytokines released from hMSCs on the endothelial progenitor cells. In order to better understand the therapeutic effect of hMSCs on neovascularization, we investigated the effects of soluble factors released from hMSCs on the EPC functions including proliferation, migration, invasion and vessel formation capacity in vitro, and also identified those factors that play roles on various EPC functions. Soluble factors released from gestational tissues-derived hMSCs were chosen to study due to easily and non-invasive collection compared to bone marrow-derived hMSCs (BM-hMSCs).
Materials and methods Subjects This study was approved by the Siriraj Institutional Review Board, Faculty of Medicine Siriraj Hospital, Mahidol University which was in accordance with the Declaration of Helsinki, the Belmont Report, CIOMS Guidelines, and ICH-GCP. Human bone marrow samples were obtained from healthy volunteers after giving written informed consent. The gestational tissues (umbilical cord, Wharton’s jelly, placenta, and amnion) were obtained from healthy newborns after receiving written informed consent from their mothers. Isolation and culture of hMSCs Ten milliliter of heparinized bone marrow was collected for hMSC isolation. Bone marrowderived mononuclear cells were then isolated using IsoPrep® (Robbins Scientific Corporation, USA) density gradient centrifugation, washed twice with PBS (GIBCO™, Invitrogen Corporation, USA), and re-suspended in complete medium which is Dulbecco’s Modified Eagle Medium (DMEM) (GIBCO™, Invitrogen Corporation, USA) supplemented with 10% (v/v) Fetal Bovine Serum (FBS) (Lonza, USA), 100 U/ml penicillin (General Drug House CO., Ltd, Thailand), and 100 µg/ml streptomycin (General Drug House CO., Ltd, Thailand). Cell suspensions were then plated in 25 cm2 culture flask (Corning, USA) at a density of 2×105 cells/cm2. Cultures were maintained at 37°C in a humidified atmosphere
Page 2 of 21
containing 5% CO2 and the medium was replaced every 3 days throughout the entire culture period.
ip t
For hMSC isolation from gestational tissues, umbilical cord, Wharton’s jelly, placenta, and amnion were cut into small pieces and digested by incubation with 0.25% (w/v) trypsinEDTA (GIBCO™, Invitrogen Corporation, USA) for 30 minutes at 37°C. Cell suspensions were washed twice with PBS, re-suspended in complete medium and plated in 25 cm2 culture flask (Corning, USA). Cultures were maintained at 37°C in a humidified atmosphere containing 5% CO2 and the medium was replaced every 3 days throughout the entire culture period.
cr
Characterization of cultured hMSCs by flow cytometry
an
us
The 3rd-5th passage of hMSCs were characterized for hMSC surface markers by incubation with the following mouse anti-human antibodies: anti-CD45-PerCP (BD Pharmingen, USA), anti-CD34-PE (Biolegend, USA), anti-CD90-FITC (AbD Serotec, USA), anti-CD73-PE (BD Pharmingen, USA), and anti-CD105-PE (Miltenyi Biotec, Germany) for 30 minutes at 4°C in the dark. After incubation with the antibodies, cell pellets were washed twice with PBS and fixed with 1% (w/v) paraformaldehyde in PBS. Flow cytometry was performed by FACS calibur™ Flow cytometer using CellQuest™ software (Becton Dickinson, USA).
Ac ce pt e
d
M
Osteogenic and adipogenic differentiation of cultured hMSCs The 3rd-5th passage hMSCs were used to assess their adipogenic and osteogenic differentiation potentials. For adipogenic differentiation, 5×104 cells were cultured in NH AdipoDiff® Medium (Miltenyi Biotec, Germany). Medium was replaced every 3 days according to the manufacturer’s instruction. After culture for 3 weeks, cells were stained with 0.5% (w/v) Oil Red O (Sigma Aldrich, USA), in isopropanol for 20 minutes at room temperature, to determine the number of hMSC-derived adipocytes in culture. For osteogenic differentiation, 5×104 cells were cultured in NH OsteoDiff® Medium (Miltenyi Biotec, Germany). Medium was replaced every 3 days according to the manufacturer’s instruction. After culture for 3 weeks, cells were stained with 40 mM Alizarin Red S (Sigma Aldrich, USA) for 20 minutes at room temperature to determine the number of hMSC-derived osteocytes in culture. Cultures and characterization of EPCs from umbilical cord blood Twenty milliliter of heparinized umbilical cord blood was collected for EPC isolation. Umbilical cord blood-derived mononuclear cells were then isolated using IsoPrep® (Robbins Scientific Corporation, USA) density gradient centrifugation, washed twice with PBS (GIBCO™, Invitrogen Corporation, USA), re-suspended in endothelial cell growth medium [endothelial basal medium-2 (LONZA, Germany), supplemented with EGM-2 single aliquots (LONZA, Germany) containing 2% (v/v) fetal bovine serum (FBS), 5 µg/ml epidermal growth factor, 200 µg/ml hydrocortisone, 0.5 µg/ml vascular endothelial growth factor, 10 µg/ml basic fibroblast growth factor, 20 µg/ml long R3 Insulin-like growth factor 1 and 1 mg/ml ascorbic acid], and plated in an individual well of 6-well plate coated with 10 µg/ml human fibronectin (Amersham Biosciences, USA) at a density of 1×106 cells/well. After culture for 3 days, the non-adherent cells were removed and fresh medium was added. Cultures were maintained at 37°C in a humidified atmosphere containing 5% CO2 and medium was replaced every 3 days throughout the entire culture period.
Page 3 of 21
Study of hMSC-derived cytokines on the functional properties of EPCs
ip t
The cultured cells were characterized for EPC surface markers by incubation with the following mouse anti-human antibodies: anti-CD34-PE (R&D Systems, USA), antiVEGFR2-PE (R&D Systems, USA), anti-CD146-FITC (R&D Systems, USA), and anti-vWFFITC (R&D Systems, USA) for 15 minutes at 4ºC in the dark. Cell pellets were then washed twice with PBS and fixed with 1% (v/v) paraformaldehyde in PBS. Flow cytometry was performed by FACScaliburTM flow cytometer (Becton Dickinson, USA) using CellQuest® software.
Ac ce pt e
d
M
an
us
cr
Effect on proliferation 2×105 hMSCs derived from five distinct tissues were cultured in 2 ml DMEM supplemented with 2% (v/v) FBS for 48 hours to generate hMSC conditioned media for proliferation assay. Briefly, 2.5×104 EPCs were cultured in an individual well of culture E-Plate (Roche Applied Science, USA) containing 100 µl hMSC conditioned media for 7 days. The number of EPCs in each well was continuously monitored throughout the entire culture period by xCELLigence Real-Time Cell Analyzer (Roche Applied Science, USA) and reported as cell index (CI). EPCs cultured in DMEM containing 2% (v/v) FBS served as controls. Effect on migration and invasion Five distinct sources of hMSCs were co-cultured with EPCs through 8 µm transwell inserts (Corning, USA) to assess the effect of hMSC-derived cytokines on the migratory and invasive capacity of EPCs. For migration assay, hMSCs were seeded in the lower chamber containing 600 µl DMEM supplemented with 2% (v/v) FBS at the density of 2.5×104 cells/cm2, while 4×104 EPCs were seeded in the transwell inserts. After 6 hours of co-culture, numbers of EPCs that migrate to the other side of the transwell’s membrane were determined by hematoxylin staining. The procedure of invasion assay was similar to that of migration assay, except that the membrane of the transwell inserts were pre-coated with 0.4 µg/µl Matrigel (BD Biosciences, USA) to serve as an extracellular matrix barrier. The numbers of invasive EPCs which are able to degrade coated Matrigel and migrate to the other side of the transwell’s membrane were determined by hematoxylin staining. Effect on vessel-forming capacity Five distinct sources of hMSCs were co-cultured with EPCs through 0.4 µm transwell inserts (Corning, USA) to assess the effect of hMSC-derived cytokines on the vessel-forming capacity of EPCs using an in vitro vessel formation assay as previously described (Jiraritthamrong et al., 2012). Briefly, 3.5×104 EPCs labeled with 5-(6)-carboxyfluorescein diacetate succinimidyl ester (CFSE, CellTrace™; Invitrogen, USA) were mixed with an equal number of non-labeled human umbilical vein endothelial cells (HUVECs) and plated into an individual well of 6-well plate pre-coated with Matrigel™ (BD Bioscience, USA) while hMSCs were seeded in the transwell inserts at the density of 1×104 cells/cm2. After 12 hours of co-culture, the extent of incorporated EPCs in the generated capillary-like structure was determined by fluorescent microscopy (Nikon, Japan) using NIS-Elements D software (Nikon, Japan). Expression levels of angiogenic-related genes in hMSCs by quantitative real-time PCR (qRT-PCR) Total RNA were isolated from hMSCs using TRIzol® reagent (Invitrogen Corporation, USA). cDNA was synthesized from 2 µg of RNA using the SuperScript™ III Reverse Trancriptase
Page 4 of 21
ip t
(Invitrogen Corporation, USA). MicroAmp® fast optical 96-well reaction plate (Applied Biosystem, USA) was used for qRT-PCR. Each well contained 3 µl cDNA, 1 µl 10 µM forward and reverse primer mix, and 10 µl SYBR® Green PCR Mastermix (Applied Biosystem, USA). The plate was then sealed with MicroAmp® clear adhesive film (Applied Biosystem, USA) to prevent evaporation of the reactant. PCR was performed using 7500 Fast Real-time PCR system (Applied Biosystem, USA) using the following protocol: 95°C initial denaturation for 10 minutes, followed by 40 cycles of denaturation (95°C, 10 seconds), annealing (60°C, 10 seconds), and extension (72°C, 40 seconds). The relative quantity of a target gene was calculated by normalization with glyceraldehyde-3-phosphate dehydrogenase (GAPDH), using the 7500 software version 2.0.5 (Applied Biosystem, USA). The primer sequences were described in Table 1.
cr
Table 1: The sequences of primers for qRT-PCR Sequence of forward primer 5ʹ-TCCTCACACCATTGAAACCA-3ʹ 5ʹ-AGGAACCAGCCTCCTCTCTC-3ʹ 5ʹ-AACCAGACGGCTGTGATGAT-3ʹ 5ʹ-AGCGGCTGTACTGCAAAAAC-3ʹ 5ʹ-GTGGTCGTGCTGGTCCTC-3ʹ
Sequence of reverse primer 5ʹ-GATCCTGCCCTGTCTCTCTG-3ʹ 5ʹ-TTCTCCAGCAGCTGTATCTCAA-3ʹ 5ʹ-TTGTCGAGAGGGAGTGTTCC-3ʹ 5ʹ-AGCCAGGTAACGGTTAGCAC-3ʹ 5ʹ-ATCTGAAGGGCACAGTTTGG-3ʹ
IGF1
5ʹ-CCGGAGCTGTGATCTAAGGA3ʹ
5ʹ-CCTGCACTCCCTCTACTTGC-3ʹ
PlGF
5ʹ-GTTCAGCCCATCCTGTGTCT-3ʹ
5ʹ-AACGTGCTGAGAGAACGTCA3ʹ
IGF2
5ʹ-TGCTGGTGCTTCTCACCTTC-3ʹ
5ʹ-AGACGAACTGGAGGGTGTCC-3ʹ
IL6
5ʹ-CAGGAGCCCAGCTATGAACT3ʹ
5ʹ-GTGAGTGGCTGTCTGTGTGG3ʹ
IL8 TGFβ
5ʹ-AAGAAACCACCGGAAGGAAC-3ʹ 5ʹ-GAGCCTGAGGCCGACTACTA-3ʹ
5ʹ-AAATTTGGGGTGGAAAGGTT-3ʹ 5ʹ-CACGTGCTGCTCCACTTTTA-3ʹ
PDGFβ
5ʹ-CCGGAGTCGGCATGAATC-3ʹ
5ʹCGTTGGAGATCATCAAAGGAG-3ʹ
GAPDH
5ʹ-GTCAACGGATTTGGTCGTATTG-3ʹ
5ʹ-CATGGGTGGAATCATATTGGAA-3ʹ
Ac ce pt e
d
M
an
us
Gene name VEGFA ANGPT1 ANGPT2 FGF2 SDF1
Study of PDGF-BB on EPC migration
4×104 EPCs were seeded into 8 µm transwell inserts that were placed into an individual well of 24-well plates containing 600 µl DMEM supplemented with 2% (v/v) FBS and various concentrations (0, 20, 50, and 100 ng/ml) of platelet-derived growth factor-BB (PDGF-BB; R&D system, USA). After 6 hours of culture, the effect of PDGF-BB on EPC migration was determined by migration assay as previously described. To further determine the role of PDGF-β on EPC migration, 4×104 EPCs were treated with 10 µg/ml anti-PDGFR-β neutralizing antibody (R&D system, USA) for 30 minutes to inhibit the action of PDGF-BB before co-cultured with hMSCs through 8 µm transwell. After 6 hours of culture, the migration assay was performed as previously described. Study of SDF1, IGF1 and PlGF on EPC invasion 4×104 EPCs were seeded into 8 µm Matrigel-coated transwell inserts that were placed into an individual well of 24-well plates containing 600 µl DMEM with 2% (v/v) FBS which were supplemented with either 100 ng/ml stromal derived factor1 (SDF1) (R&D system, USA), 100 ng/ml insulin-like growth factor1 (IGF1; R&D system, USA), 100 ng/ml placenta growth
Page 5 of 21
factor (PlGF; R&D system, USA), or combination of those three factors (100 ng/ml each). After culture for 18 hours, the effect of those factors on capacity of EPC invasion was determined by invasion assay as previously described.
cr
Study of SDF1, IGF1 and PlGF on vessel-forming capacity of EPCs
ip t
To further determine the roles of SDF1, IGF1 and PlGF on EPC invasion, 4×104 EPCs were pre-treated with either 10 µM AMD3100 octahydrochloride (inhibitor of SDF1; Tocris Bioscience, UK), 10 µg/ml anti-hIGF1R (inhibitor of IGF1; R&D system, USA), 5 µg/ml anti-hVEGFR1 (inhibitor of PlGF; R&D Systems, USA), or the combination of all those three inhibitors for one hour before co-culture with hMSCs through 8 µm Matrigel-coated transwell. After culture for 18 hours, the invasion assay was performed as previously described.
an
us
3.5×104 CFSE-labeled EPCs were co-cultured with an equal number of non-labeled HUVECs in endothelial basal medium supplemented with either 100 ng/ml SDF1, 100 ng/ml IGF1, 100 ng/ml PlGF, or the combination of all those three factors (100 ng/ml each). The mixtures of cells were then seeded into an individual well of Matrigel-coated 24-well plate. After 12 hours of culture, an in vitro vessel formation assay was performed as previously described.
Statistical analysis
d
M
To further verify the roles of SDF1 and IGF1 on vessel-forming capacity of EPCs, 105 EPCs were treated with either 10 µM AMD3100 octahydrochloride (SDF1 inhibitor; Tocris Bioscience, UK) or 10 µg/ml anti-hIGF1R (IGF1 inhibitor; R&D system, USA) for one hour before co-culture with BM-hMSCs through transwell culture system. An in vitro vessel formation assay was performed as previously described.
Ac ce pt e
Data were presented as mean ± standard error of the mean (SEM). The paired Student’s t-test and one-way ANOVA were used to assess the significance of differences between observed data. P < 0.05 was considered to be statistically significant.
Results
Characteristics of hMSCs derived from gestational tissues and bone marrow hMSCs derived from gestational tissues, including placenta (PL-hMSCs), umbilical cord (UC-hMSCs), Wharton’s jelly (WJ-hMSCs) and amnion (AM-hMSCs) exhibited similar characteristics to those of bone marrow-derived hMSCs (BM-hMSCs). The gestational tissuederived hMSCs displayed fibroblast-like morphology (Fig. 1A-E), expressed typical hMSC surface markers (positive for CD73, CD90, CD105 and negative for hematopoietic markers CD34 and CD45; Fig. 1P). Moreover, the gestational tissue-derived hMSCs could differentiate toward adipocyte- and osteocyte-lineages as demonstrated by Oil-Red O (Fig. 1F-J) and Alizarin Red S staining (Fig. 1K-O), respectively. Expression of pro-angiogenic genes in gestational tissue and bone marrow-derived hMSCs The expression levels of pro-angiogenic genes among various sources of hMSCs were determined by quantitative real-time PCR. The expression levels of stromal cell-derived
Page 6 of 21
ip t
factor 1 (SDF1), insulin-like growth factor 1 (IGF1), and placental growth factor (PlGF) genes in the BM-hMSCs were significantly higher than those of other hMSC sources (Fig. 2). In contrast, the expression level of platelet-derived growth factor beta (PDGF-β) was significantly higher in PL-hMSCs than the rest of the hMSC sources. Apart from those 4 genes, the expression levels of several pro-angiogenic genes, including vascular endothelial growth factor A (VEGF-A), fibroblast growth factor 2 (FGF2), angiopoietin 1 (ANGPT1), angiopoietin 2 (ANGPT2), insulin-like growth factor 2 (IGF2), interleukin 6 (IL6), interleukin 8 (IL8), and transforming growth factor beta (TGF-β) were not significantly different among the various hMSC sources. Characteristics of umbilical cord blood-derived endothelial progenitor cells (EPCs)
us
cr
EPCs were cultured from mononuclear cell populations that had been separated from umbilical cord blood by density gradient centrifugation. After culture in endothelial growth medium for 7-10 days, several EPC-like colonies of cobblestone morphology were observed (Fig. 3A). These cells expressed typical EPC surface markers, including CD34, CD146, Von Willebrand factor (vWF) and vascular endothelial growth factor receptor 2 (VEGFR2) (Fig. 3B), and were able to form capillary-like structures on Matrigel™ (Fig. 3C).
an
Effects of hMSC-derived soluble factors on the functional properties of EPCs
Ac ce pt e
d
M
Effect of hMSC secreted factors on EPC proliferation. The conditioned medium derived from hMSCs from various sources were collected and used to culture EPCs. Similar to control group (cultured in DMEM supplemented with 2% [v/v] FBS), the numbers of EPCs cultured in various hMSC-conditioned media were increased initially during the first 48 hours of culture before rapidly decreasing toward the baseline level at the end of culture (Fig. 4), indicating that the soluble factors secreted from bone marrow and gestational tissue-derived hMSCs did not promote EPC proliferation. B) Effect of hMSC secreted factors on EPC migration. EPCs were co-cultured with hMSC from various sources using the transwell culture system (Fig. 5A). The numbers of EPCs that migrate toward various hMSC sources were determined after 6 hours of co-culture. Interestingly, only the soluble factors derived from PL-hMSCs significantly enhanced EPC migration in comparison to controls (306.2 ± 60% vs. 100% of control, P
