Mesenchymal stem cell secreted platelet derived growth factor exerts a pro-migratory effect on resident Cardiac Atrial appendage Stem Cells

Journal of Molecular and Cellular Cardiology 66 (2014) 177–188

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Journal of Molecular and Cellular Cardiology journal homepage: www.elsevier.com/locate/yjmcc

Original article

Mesenchymal stem cell secreted platelet derived growth factor exerts a pro-migratory effect on resident Cardiac Atrial appendage Stem Cells Severina Windmolders a,b, Astrid De Boeck c, Remco Koninckx a,b, Annick Daniëls a, Olivier De Wever c, Marc Bracke c, Marc Hendrikx b,d, Karen Hensen a,b, Jean-Luc Rummens a,b,⁎ a

Laboratory of Experimental Hematology, Jessa Hospital, Campus Virga Jesse, Stadsomvaart 11, 3500 Hasselt, Belgium Faculty of Medicine and Life Sciences, Hasselt University, Martelarenlaan 42, 3500 Hasselt, Belgium Department of Radiation Oncology and Experimental Cancer Research, Ghent University Hospital, De Pintelaan 185, 9000 Ghent, Belgium d Department of Cardiothoracic Surgery, Jessa Hospital, Campus Virga Jesse, Stadsomvaart 11, 3500 Hasselt, Belgium b c

a r t i c l e

i n f o

Article history: Received 20 July 2013 Received in revised form 12 November 2013 Accepted 28 November 2013 Available online 8 December 2013 Keywords: Myocardial infarction Mesenchymal stem cells Conditioned medium Cardiac stem cells Migration Platelet derived growth factor

a b s t r a c t Mesenchymal stem cells (MSCs) modulate cardiac healing after myocardial injury through the release of paracrine factors, but the exact mechanisms are still unknown. One possible mechanism is through mobilization of endogenous cardiac stem cells (CSCs). This study aimed to test the pro-migratory effect of MSC conditioned medium (MSC-CM) on endogenous CSCs from human cardiac tissue. By using a three-dimensional collagen assay, we found that MSC-CM improved migration of cells from human cardiac tissue. Cell counts, perimeter and area measurements were utilized to quantify migration effects. To examine whether resident stem cells were among the migrating cells, specific stem cell properties were investigated. The migrating cells displayed strong similarities with resident Cardiac Atrial appendage Stem Cells (CASCs), including a clonogenic potential of ~21.5% and expression of pluripotency associated genes like Oct-4, Nanog, c-Myc and Klf-4. Similar to CASCs, migrating cells demonstrated high aldehyde dehydrogenase activity and were able to differentiate towards cardiomyocytes. Receptor tyrosine kinase analysis and collagen assays performed with recombinant platelet derived growth factor (PDGF)-AA and Imatinib Mesylate, a PDGF receptor inhibitor, suggested a role for the PDGFAA/PDGF receptor α axis in enhancing the migration process of CASCs. In conclusion, our findings demonstrate that factors present in MSC-CM improve migration of resident stem cells from human cardiac tissue. These data open doors towards future therapies in which MSC secreted factors, like PDGF-AA, can be utilized to enhance the recruitment of CASCs towards the site of myocardial injury. © 2013 Elsevier Ltd. All rights reserved.

1. Introduction Abbreviations: 3D, Three-dimensional; ALDH, Aldehyde dehydrogenase; AT, Annealing temperature; AxlR, Axl receptor; BM-SCs, Bone marrow stem cells; CASCs, Cardiac Atrial appendage Stem Cells; CFU-Fs, Colony-forming-unit fibroblasts; CSCs, Cardiac stem cells; cTn, Cardiac troponin; DEAB, Diethylamino-benzaldehyde; EGFR, Epidermal growth factor receptor; ELISA, Enzyme-linked immunosorbent assay; FAK, Focal adhesion kinase; FCS, Fetal calf serum; FGFR, Fibroblast growth factor receptor; FMO, Fluorescence minus one; GFP, Green fluorescent protein; IHD, Ischemic heart disease; InsR, Insulin receptor; LG-DMEM, Low-glucose Dulbecco's Modified Eagle Medium; MI, Myocardial infarction; MMPs, Matrix metalloproteinases; MSCs, Mesenchymal stem cells; MSC-CM, Mesenchymal stem cell conditioned medium; NRCMs, Neonatal rat cardiomyocytes; PDGF, Platelet derived growth factor; PDGFR, Platelet derived growth factor receptor; PLC-γ, Phospholipase C-γ; PI3K, Phospho-inositide-3 kinase; P/S, Penicillin/ streptomycin; RALDH2, Retinaldehyde dehydrogenase 2; RTK, Receptor tyrosine kinase; RT-PCR, Reverse transcriptase PCR; Tβ4, Thymosin β4; Tbx18, T-Box 18; TGF-β, Transforming growth factor-β; VEGF, Vascular endothelial growth factor; VEGFR, Vascular endothelial growth factor receptor; Wt1, Wilm's tumor 1. ⁎ Corresponding author at: Laboratory of Experimental Hematology, Jessa Hospital, Campus Virga Jesse, Stadsomvaart 11, 3500 Hasselt, Belgium. Tel.: +32 11309740; fax: +32 11309750. E-mail addresses: [email protected] (S. Windmolders), [email protected] (A. De Boeck), [email protected] (R. Koninckx), [email protected] (A. Daniëls), [email protected] (O. De Wever), [email protected] (M. Bracke), [email protected] (M. Hendrikx), [email protected] (K. Hensen), [email protected] (J.-L. Rummens). 0022-2828/$ – see front matter © 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.yjmcc.2013.11.016

In the last decade, stem cell therapy has emerged as an innovative approach to restore cardiac function after myocardial infarction (MI) either directly by regeneration of functional myocardium [1] or indirectly by paracrine actions stimulating cardiac tissue healing [2]. Previously, researchers reported the existence of cardiac stem cells (CSCs) residing in the adult mammalian heart [3,4]. While phase 1 clinical studies [5,6] are completed only recently, CSC transplantations performed in the past already showed an improved cardiac function in animal models through regeneration of the damaged myocardium [7]. In the past, most experimental and clinical studies concerning ischemic heart disease (IHD) were performed with bone marrow stem cells (BM-SCs) [8,9]. Although it was demonstrated that BM-SC implantation can reduce ventricular remodeling and improve left ventricular function after MI, the underlying mechanism is still under debate [10]. Recent data propose that among BM-SCs, mesenchymal stem cells (MSCs) are especially capable of mediating cardiac repair through the release of a broad spectrum of cytokines, growth factors and chemokines into the damaged tissue area [11]. Strong evidence comes from studies that have utilized the conditioned medium derived from MSCs (MSC-CM).

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Indeed, using in vitro assays and small animal models of MI, researchers have found that the administration of concentrated MSC-CM significantly improves myocardial regeneration and ventricular function [12]. Proposed mechanisms of action include the enhancement/ modulation of cytoprotection [13], neovascularization [14], contractility [15], fibrotic remodeling [16] and inflammatory processes [17]. One proposed mechanism of paracrine influence, which has to date received little attention, is the mobilization of CSCs towards the injured site. To this end, the most important therapeutic goal is to stimulate CSCs to form cardiomyocytes and vascular cells to repopulate and regenerate the injured tissue. In this study, we postulate that migration of endogenous stem cells from cardiac tissue can be enhanced by specific factors secreted by MSCs. To address our hypothesis, we directly assessed the effect of MSC-CM on human cardiac tissue fragments cultured inside a threedimensional (3D) collagen matrix. We showed that MSCs secrete factors that improve migration of resident Cardiac Atrial appendage Stem Cells (CASCs), that are characterized by high levels of aldehyde dehydrogenase (ALDH) expression [4]. Further results indicated that the platelet derived growth factor receptor α (PDGFRα) plays an important role in the migration process. These findings open new perspectives to stimulate cardiac tissue healing via activation of endogenous repair mechanisms. 2. Materials and methods All procedures were carried out in accordance with the principles set forth in the Helsinki Declaration. Approval by the institutional review board and informed consent from each patient were obtained. All animal studies were approved by the Hasselt University Institutional Animal Care and Use Committee. A detailed version of Materials and methods is available in the Suppl. Material. 2.1. Preparation of MSC-CM Human bone marrow MSCs were obtained as previously described [18]. Once MSCs of at least passage 4 (P4) reached 70–75% confluence, medium was replaced with low-glucose Dulbecco's Modified Eagle Medium (LG-DMEM; Invitrogen) containing 10% fetal calf serum (FCS: Hyclone) and 2% penicillin/streptomycin (P/S; Lonza) until MSC-CM was prepared. MSC-CM was prepared from 85 to 90% confluent T175-cm2 flasks containing 1.5 × 106 MSCs as described [19]. MSC-CM at 10× concentration was utilized, unless stated otherwise. 2.2. Collagen migration assay Migration assays were adapted from experiments performed by De Wever et al. with minor modifications [20]. Under a macroscope with a calibrated ocular grid (Wild M5, Wild Heerbrugg), atrial appendages were cut into ±400 μm3 fragments and suspended in collagen type I solution (1 mg/ml; Becton&Dickinson). Depending on the type of experiment, either 5 × 105 MSCs were plated on the bottom of the well or 1 ml of the appropriate medium was administered on top of the matrix. Where only medium was administered, 1 ml of serum-free LG-DMEM served as negative control medium. Tested media consisted of 1 ml of prepared MSC-CM or 1 ml of recombinant platelet derived growth factor (PDGF)-AA (R&D Systems) dissolved in serum-free LG-DMEM at a concentration of 0.1 ng/ml, 1 ng/ml or 100 ng/ml. Additionally, inhibition experiments were performed in which 1 μM of the multi-target inhibitor Imatinib Mesylate (Selleckchem) was added to control, MSC-CM or PDGF-AA conditions, as indicated. Alternatively, to block receptor binding by PDGF-AA, a neutralizing PDGFRα antibody (R&D systems) was added to control and MSC-CM at a final concentration of 20 ng/ml. Cellular migration was regularly scored during a total period of at least 10 days. Per test condition, minimal 20 randomly selected

tissue fragments were analyzed. The number of migrating cells was counted using an Axiovert 200 M microscope (Zeiss). Perimeter and area measurements were performed on the evasion zone from the tissue fragments with Axiovision 4.8 software (Zeiss) and utilized as measures of migration. Tissue viability was assessed at the end of the followup using an Annexin V staining kit (Becton&Dickinson) according to the manufacturer's instructions. 2.3. Isolation and expansion of migrating cells To isolate migrating cells, cardiac tissue pieces were first removed from the collagen matrix. The remaining collagen was dissolved by collagenase type II treatment (600 U/ml; Invitrogen) at room temperature for 10 min. Cells were seeded in a 96-well plate and expanded in X-Vivo 15 medium containing 20% FCS and 2% P/S. After P1, serum level was reduced to 10%. 2.4. Cell cycle analysis of migrating cells For analysis of cell cycle distribution, the BD Cycletest™ Plus DNA Reagent Kit (Becton& Dickinson) was utilized, in accordance with the manufacturer's instructions. To obtain a mitotic arrest at the metaphase, 10 ng/ml KaryoMAX® Colcemid™ Solution (Invitrogen) was added for 48 h. Cellular DNA content was monitored on a FACSCanto® (Becton&Dickinson). Cells in S + G 2 M phase were judged to be actively proliferating cells. Mitotic indices were calculated with ModFit LT software 3.0 (Verity Software House). 2.5. Clonogenic assay Prior to assess the clonogenic character, migrating cells were labeled with green fluorescent protein (GFP). Subsequently, a total of 864 GFP+ cells (P5-P7) were flow sorted with a FACSAria® at a density of 1 cell/ well in X-Vivo 15 medium containing 20% FCS. Single-cell deposition was confirmed by fluorescence microscopy and wells containing more than one cell were excluded. Colonies were scored after 10 days. 2.6. Expression of pluripotency associated genes Total RNA was isolated from expanded migrating cells (P3–P5) using the RNeasy Mini kit (Qiagen). cDNA was synthesized using Superscript III and random hexamers (Invitrogen). Reverse transcriptase PCR (RTPCR) using Taq polymerase (Roche) was performed for 35 cycles consisting of 40 s at 95 °C, 50 s at annealing temperature (AT) and 1 min at 72 °C with a final extension step of 10 min at 72 °C. β-Actin was used as control. Primer sequences with corresponding AT and expected fragment size are listed in Suppl. Table 1. 2.7. Analysis of ALDH activity ALDH expression of migrating cells was analyzed with the Aldefluor™ kit (Aldagen, Inc). Cells were seeded in 24-well plates at 1 × 104 cells/well and incubated in 500 μl Aldefluor assay buffer containing 2.5 μl of activated Aldefluor® (Aldagen, Inc). To confirm specificity for ALDH, 20 μl of diethylamino-benzaldehyde (DEAB) was administered to block ALDH activity. After incubation at 37 °C for 30 min, cells were washed and kept in Aldefluor assay buffer for microscopical visualization for the green fluorescent reaction product. Exposure times were kept constant during each acquisition. 2.8. Flow cytometrical analysis The antigen expression profile of migrating cells and MSCs was determined by flow cytometry. 5 × 104 cells/tube were incubated for 20 min in the dark with human monoclonal antibodies as recommended by the manufacturer.

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2.9. Functional differentiation assays

2.15. Statistical analysis

Differentiation of migrating cells into adipocytes was conducted with the Human Mesenchymal Stem Cell Functional Identification Kit (R&D Systems), according to the manufacturer's recommendations. Adipogenic differentiation was evaluated using Oil Red O staining. In order to stimulate cardiomyogenic differentiation, co-culture systems were set up between GFP+ migrating cells and primary neonatal rat cardiomyocytes (NRCMs) as previously described [18]. After one week, cardiac differentiation was evaluated by immunofluorescence for expression of cardiac troponin (cTn)-T and -I. CASCs were utilized as positive control.

Comparisons between groups were performed using a two-way repeated measures ANOVA followed by a Mann–Whitney test after d'Agostino and Pearson testing for normal distribution. All illustrated data are representative of cardiac specimens obtained from at least four patients and are displayed as mean ± SEM. Recurring conditions in separate figures illustrate data gathered from replicate experiments. Statistical tests were two-sided. For transwell migration assays, d'Agostino and Pearson testing for normal distribution was performed followed by a Mann–Whitney test. Data are representative for four different patients and displayed as mean ± SEM. *p b 0.05, **p b 0.01 and ***p b 0.001.

2.10. Isolation of CASCs CASCs were obtained from atrial appendages removed during cardiac surgery, as previously described by Koninckx et al. [4]. 2.11. Transwell migration assays To examine chemotaxis, 5 × 105 CASCs (n = 4) were seeded in serum-free LG-DMEM onto the upper chamber of 8-μm Thincert™ 24well inserts (Greiner Bio-One). The bottom chambers were filled with serum-free LG-DMEM (control medium), LG-DMEM with 10% FCS (positive control) or MSC-CM. After 48 h, migrated CASCs were stained with 0.1% crystal violet and images were taken in 2 randomly selected fields on an Axiovert 200 M microscope (Zeiss). Quantification was performed by Axiovision 4.8 software (Zeiss). Values were expressed as mean area percentage and normalized to the positive control. 2.12. Receptor tyrosine kinase (RTK) array and western blotting A human phospho-RTK array kit (ARY001, R&D systems) was used to simultaneously detect the relative tyrosine phosphorylation levels of 42 different RTKs. Assays were performed according to the manufacturer's protocol. Expanded CASCs and migrating cells were subjected to control medium or MSC-CM at 37 °C/5% CO2 for 90 min. PDGFRα expression was evaluated by western blotting. Lysates were prepared from CASCs directly isolated from human atrial appendages, cultured CASCs and CASCs incubated for 90 min in MSC-CM. MSCs and Human Umbilical Vein Endothelial Cell line (HUVEC) served as positive [21] and negative controls, respectively. To evaluate PDGFRα phosphorylation, lysates were prepared from expanded CASCs subjected to serum-free LG-DMEM control medium or MSC-CM at 37 °C/5%CO2 for 90 min. The following primary antibodies were used: rabbit monoclonal anti-PDGFRα (1:1000; Cell Signaling), rabbit monoclonal anti-pPDGFRα (tyr849) (1:1000; Cell Signaling) and mouse monoclonal anti-α-Tubulin (1:5000; Abcam). 2.13. Immunofluorescence Immunofluorescence was performed on directly isolated CASCs with abovementioned rabbit monoclonal anti-PDGFRα antibody (1:50; Cell Signaling), followed by a secondary sheep anti-rabbit rhodaminelabeled antibody. Stainings with only the secondary antibody served as negative control. 2.14. Enzyme-linked immunosorbent assay (ELISA) The concentration of PDGF-AA in MSC-CM was quantified with a Human/Mouse PDGF-AA Quantikine ELISA Kit (R&D Systems). ELISA assays were performed in two independent assays using MSC-CM prepared from different MSC populations. Data are displayed as mean ± SEM.

3. Results 3.1. MSC-CM enhances cellular migration from human cardiac tissue Co-culture experiments between MSCs and human cardiac tissue fragments in type I collagen gel, demonstrated that MSCs stimulated migration of cells from the cardiac tissue fragments throughout the collagen matrix (Suppl. Fig. 1A). To investigate whether this functional effect was caused by paracrine mediators released by the MSCs, we performed collagen migration assays in which MSCs were replaced with MSC-CM. We observed that treatment of cardiac fragments with MSC-CM promoted cellular migration in a similar way (Fig. 1A). Morphologically, migrating cells displayed an invasive phenotype with the formation of cellular extensions or filopodia in control (Suppl. Fig. 1B upper panel) as well as MSC-CM (Suppl. Fig. 1B lower panel). To perform a more quantitative analysis, perimeter and area measurements were utilized to measure the cellular migration distance. Cardiac fragments cultured in the presence of MSC-CM generated migrating cells that traveled over larger perimeter (p b 0.001, Fig. 1B) and area (p b 0.001, Suppl. Fig. 2A) surfaces, in comparison with controls. The effect initiated at day 6 and lasted up to day 10. Furthermore, at day 6 and day 10, the number of migrating cells was counted in relation to the distance traveled from the center of the cardiac fragments. According to preliminary screening experiments in which the maximum migration distance was assessed in control and MSC-CM, migrating cells were categorized into 3 distinct zones: zone A (b450 μm), zone B (450–750 μm) and zone C (N750 μm) (Suppl. Fig. 3). After 6 days, the amount of migrating cells was 2.0-fold (p b 0.01) and 4.9-fold (p b 0.001) higher for MSC-CM in zones A and B, respectively (Fig. 1C). Zone C contained no cells in control conditions. At day 10, 1.7-fold (p b 0.001) and 2.2fold (p b 0.001) differences were observed in zones A and B, respectively. At this time point, cells in the control condition also migrated into zone C, but the number of cells in MSC-CM was 8.3-fold higher (p b 0.001; Fig. 1D). When the total number of migrating cells was counted at day 6 (p b 0.001) and day 10 (p b 0.001), without taking the different zones into account, migration effects were much more pronounced in MSC-CM as well (Suppl. Fig. 2B). Next, to assess whether MSC-CM influenced migration of cells from cardiac fragments in a dose dependent way, different concentrations of MSC-CM were evaluated. Perimeter (p b 0.01; Fig. 2A) and area measurements (p b 0.05; Suppl. Fig. 4A) illustrated increased migration distances in all MSC-CM conditions compared to control, except for perimeter measurements of 10× MSC-CM at day 6. In general, the effect started at day 6 and lasted during the entire follow-up. When the cell number was counted in relation to the distance traveled, we found that, at day 6, the cell number in zones A and B was significantly higher in all conditions compared to controls (p b 0.05; Fig. 2B). In zone C, no cells were detected. At day 10, increased migration was clearly observed in all zones in all MSC-CM conditions (p b 0.01; Fig. 2C), except for 10× MSC-CM in zone C. Considering the total number of cells regardless of the different zones, migration was significantly elevated in all MSCCM conditions at day 6 as well as day 10 (p b 0.01; Suppl. Fig. 4B). In

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Fig. 1. MSC-CM promotes migration of cells from human cardiac tissue fragments. A) Representative phase contrast images of cardiac fragments cultured in collagen type I matrix in control medium (upper panel) or MSC-CM (lower panel) at indicated time intervals. Scale bar, 500 μm. B) Graph representing perimeter measurements illustrative of the migration distances traveled by cells in control versus MSC-CM. Cell number in function of the migration distance as categorized into zone A (b450 μm), B (450 μm–750 μm) or C (N750 μm) in control versus MSC-CM at day 6 (C) and day 10 (D). *p b 0.05, **p b 0.01, ***p b 0.001 compared to control.

general, the migration responses followed a dose dependent trend, with a statistically significant difference between area surfaces of 10 × vs. 30× MSC-CM after 6 days (p b 0.05; Suppl. Fig. 4A). 3.2. Migrating cells display strong similarities with resident CASCs In a next phase, we wanted to identify the cells that migrated from the cardiac tissue fragments. Cell characteristics and the antigenic expression profile were investigated. Cells were ex vivo expanded after isolation from the collagen matrix. The proliferative capacity of these expanded cells indicated a normal DNA content and a mean mitotic index of 20.58 ± 9.59% (n = 6). Further investigation learned that migrating cells mainly resembled progenitor cells originating from human cardiac tissue. Expanded migrating cells (P3–5) expressed several pluripotency associated genes like Oct-4, Nanog, Dppa-3, Tbx-3, c-Myc and Klf-4. However, no expression of Sox-2, Tert, nor Gdf-3 could be detected (Fig. 3A). Moreover, when the clonogenic potential was tested by single cell sorting of expanded GFP+ migrating cells (P5–P7), 187 clones were counted after 10 days. This corresponded to a clonogenic percentage of 21.6 ± 4.3% (n = 3) (Fig. 3B). The antigenic expression profile illustrated that migrating cells expressed numerous markers that are involved in cell–cell and cell–matrix adhesion, as well as cellular migration. These markers

included CD49c, CD73, CD44, CD29, CD105, CD13 and CD90 (Suppl. Fig. 5A). On the other hand, cells lacked expression of the hematopoietic stem cell marker CD34, the pan leukocyte marker CD45, the MSC/stromal cell markers CD271 and PDGFRβ (CD140b) as well as the receptor for stem cell factor c-kit or CD117 (Suppl. Fig. 5B). This antigenic phenotype differs from HSCs (lack of CD34), MSCs (lack of CD140b) as well as c-kit+ CSCs (lack of c-kit), but seems to be more related to the recently described CASCs [4] as well as epicardial colony-forming-unit fibroblasts (CFU-Fs) [22]. To further demonstrate this, ALDH activity was assessed in migrating cells. Since recent studies demonstrated that the preservation of ALDH activity during expansion is a unique feature seen in CASCs but not in MSCs [4,23,24], CASCs and MSCs were used as positive and negative controls, respectively. High ALDH activity could clearly be detected in migrating cells (Fig. 4A) compared to measurements with DEAB (negative control) (Fig. 4D). This was similar to the elevated ALDH activity seen in CASCs (Fig. 4B). Expanded MSCs did not display ALDH activity (Fig. 4C). To further confirm that the migrating cells were indeed resident CASCs and not mobilized bone marrow MSCs, functional differentiation assays were performed. First, in contrast to the MSCs (Suppl. Fig. 6B), migrating cells were unable to differentiate into adipocytes as illustrated by the lack of lipid droplets inside the cells' cytoplasm until at least 3 weeks after induction of differentiation (Suppl. Fig. 6A). Furthermore, in contrast to MSCs, CASCs are able to

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Fig. 2. MSC-CM enhances cellular migration following a dose dependent trend. A) Graph depicting perimeter measurements illustrative of the cells' migration distances in either control, 10× MSC-CM (standard), 20× MSC-CM and 30× concentrated MSC-CM. Conditions as indicated. Number of cells in function of their migration into zone A, B or C at day 6 (B) and day 10 (C). No statistical differences were detected between MSC-CM concentrates.

differentiate down the cardiac lineage, so we verified the differentiation capacity of migrating cells towards cardiomyocytes [4,18]. GFP+ migrating cells and CASCs (positive control) were cultured under serum deprived conditions in the presence of NRCMs. After 1 week of coculture, both migrating cells as well as CASCs clearly resembled a cardiac phenotype as indicated by pronounced expression of cardiac specific markers cTnT (Fig. 5A) and cTnI (Fig. 5B), compared to negative control staining (Suppl. Fig. 7). When migrating cells were placed in standard or monoculture conditions, no cTnT nor cTnI could be detected (data not shown). Based on these results, we subsequently investigated whether MSC-CM was able to directly influence chemotaxis of CASCs. Therefore, transwell assays were performed in which CASCs were allowed to migrate under the influence of serum-free LG-DMEM (control medium), LG-DMEM with 10% FCS (positive control) or MSC-CM (Fig. 6A). Results from these assays indicated that MSC-CM effectively stimulated CASC chemotaxis, as displayed by a 5.8-fold higher chemotaxis towards factors present in MSC-CM compared to the control (57.54 ± 10.41% in MSC-CM vs. 9.98 ± 2.02% in control; p b 0.05; Fig. 6B). Moreover, MSC-CM induced migration was only 1.7-fold less and not significantly different from the chemotactic response induced by the positive control. These results clearly indicated that MSC-CM exerts a prochemotactic effect on CASCs in vitro. 3.3. A potential role for the PDGF-AA/PDGFRα axis in CASC migration To investigate which signaling pathways were involved in the observed migration response, a human p-RTK array was utilized to detect the phosphorylation levels of 42 different RTKs (Suppl. Table 2). Expanded CASCs and migrating cells treated with MSC-CM revealed a substantial increase in the activity of the PDGFRα (ranging from 27.8 to 44.1-fold, Suppl. Fig. 8), which was the only receptor that was consistently and strongly phosphorylated in 3 independent experiments. In addition, the RTK array showed modest increases in the activation of the epidermal growth factor receptor (EGFR, 4.5-fold), the PDGFRβ (8.4-fold), the insulin receptor (InsR, 2.6-fold), the Axl receptor (AxlR,

2.4-fold) and the vascular endothelial growth factor receptor 1 (VEGFR1, 4.9-fold), but also a moderate decrease in the activation of fibroblast growth factor receptor 3 (FGFR3, 7.7-fold) (data not shown). Nonetheless, these alterations in receptor activation status were detected only once and were not reproducible. RTK array analysis with 30× MSC-CM, the highest MSC-CM concentrate used in this study, did not deliver extra results besides another robust activation of the PDGFRα (14.1-fold). Since only PDGFRα activation was reproducible, we confirmed these results before further examination. By western blot analysis, the expression of the PDGFRα was demonstrated not only on CASCs directly isolated from human atrial appendages, but also on CASCs in culture and CASCs subjected to MSC-CM (Fig. 7A). For the latter, expression of the PDGFRα was also confirmed by immunofluorescence (Fig. 7B). Subsequently, by western blotting, we also confirmed the phosphorylation of the PDGFRα on CASCs after MSC-CM treatment (Fig. 7C). The PDGFRα is known to interact with all three PDGF-ligands, PDGFAA, -AB and -BB [25]. However, based on the results from several studies [26,27], showing that bone marrow MSCs only secrete PDGF-AA, but not -AB and -BB ligand, we further investigated the role of the PDGF-AA/ PDGFRα signaling pathway in the migration process. By performing an ELISA assay, the PDGF-AA concentration in MSC-CM was found to be 37 ± 6 pg/ml (n = 7). To investigate whether PDGF-AA was capable of enhancing recruitment of CASCs in a similar way, migration assays were set up in which MSC-CM was replaced by different concentrations of PDGF-AA. As shown by perimeter measurements, 100 ng/ml PDGFAA significantly enhanced migration of cells from cardiac tissue in a similar way as MSC-CM (p b 0.05, Fig. 8A), albeit with a slight temporal delay. PDGF-AA at lower concentrations of 0.1 ng/ml and 1 ng/ml also significantly promoted migration after 13 days (p b 0.05). Area measurements showed similar results (Suppl. Fig. 9A). Because the migration response with PDGF-AA was slightly delayed, observations were quantified at days 8 and 13, instead of days 6 and 10, respectively. Zonal (p b 0.05; Figs. 8B and C) and total cell counts (p b 0.001; Suppl. Fig. 9B) illustrated that the amount of cells in 100 ng/ml PDGF-

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Fig. 3. Migrating cells display typical stem cell features. A) Representative RT-PCR showing the expression of pluripotency associated genes Oct-4, Nanog, Dppa-3, Tbx-3, c-Myc and Klf-4 by ex vivo expanded migrating cells (MC). H2O was used as non-template control (NTC). Marker (M). Different parts of the same gel are made explicit by dividing lines. β-Actin (β-act) was used as control. B) Fluorescent images showing clone formation by migrating cells at indicated time intervals. Scale bar upper panel, 200 μm; scale bar lower left figure, 500 μm, inset scale bar, 200 μm.

AA was markedly increased, similar to that of MSC-CM. In the same way, significant effects were also detected for 0.1 and 1 ng/ml PDGF-AA, especially after 13 days (p b 0.05; Fig. 8B and Suppl. Fig. 9B). In addition, analyzing the PDGF-AA dose responsive effect, we generally found significant differences between 0.1 and 100 ng/ml PDGF-AA for all measurements, especially at day 8. Also, between 1 and 100 ng/ml PDGFAA, we found significant differences, but only when comparing the number of migrating cells (connecting bars; Fig. 8 and Suppl. Fig. 9). Furthermore, cells migrating in response to PDGF-AA illustrated a similar phenotype as cells migrating in response to MSC-CM (data not shown) and stained positive for ALDH as well (Fig. 8D). The observed response was predominantly migration dependent since the addition of mitomycin C, a proliferation inhibitor, did not considerably alter the migration pattern in control, MSC-CM nor 100 ng/ml PDGF-AA conditions (data not shown). Additional Ki67 stainings performed on migrating cells suggested that the observed responses were mainly migration and not proliferation dependent (data not shown). To examine whether the observed migration response was specifically mediated by PDGF, the effect of the PDGFR inhibitor Imatinib Mesylate was investigated. Migration assays were set up in which Imatinib was administered to control, MSC-CM as well as 100 ng/ml PDGF-AA (Suppl. Figs. 10A–F). We opted for a concentration of 1 μM Imatinib as it did not influence the basal migration response observed in the control condition (Fig. 9 and Suppl. Fig. 11). Based on perimeter (p b 0.05, Fig. 9A) and area (p b 0.05, Suppl. Fig. 11A) measurements, Imatinib significantly compromised the cells' migration distance in PDGF-AA

and MSC-CM. Also, the number of cells in function of their migration distance, was largely decreased at day 8 (p b 0.05, Fig. 9B) and day 13 (p b 0.01, Fig. 9C) when Imatinib was added to PDGF-AA and MSC-CM conditions. Furthermore, without taking the distance into account, Imatinib decreased the total number of cells migrating under the influence of PDGF-AA and MSC-CM (Suppl. Fig. 11B). Importantly, PDGF-AA and MSC-CM induced migration was significantly increased compared to control in all graphs illustrated (Fig. 9 and Suppl. Fig. 11). So, in general, Imatinib significantly attenuated PDGF-AA as well as MSC-CM induced migration, albeit the impact of Imatinib on MSC-CM induced migration was less prominent. The inhibitory effect of Imatinib was predominantly caused by attenuating the cells' migration capacity and not by induction of apoptosis or inhibition of proliferation, as evaluated by Annexin V and Ki67 stainings on CASCs in control, MSC-CM and 100 ng/ml PDGF-AA (data not shown). As Imatinib is a multi-target tyrosine kinase inhibitor that is also known to influence c-kit and Abl, we performed confirmation experiments in which we tested the effect of a specific neutralizing PDGFRα antibody on MSC-CM induced migration. At day 8, perimeter measurements illustrated a reduction of 19.86% with Imatinib vs. 21.62% with PDGFRα antibody and 18.70% with Imatinib vs. 11.19% with PDGFRα antibody at day 13. Moreover, cell counts illustrated a decreased migration of 51.22% with Imatinib and 41.45% with PDGFRα antibody after 8 days, and 50.94% with Imatinib vs. 33.57% with antibody after 13 days, illustrating that the reduction in migration by Imatinib is mediated by the PDGFRα.

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Fig. 4. Migrating cells demonstrate elevated ALDH activity. Photomicrographs representing migrating cells (MC) (A), CASCs (B) and MSCs (C) in culture after incubation with activated Aldefluor®. DEAB (inhibitor of ALDH activity) (D) served as negative control. Phase contrast (left) and ALDH (middle) images were taken and merged (right). Scale bar, 200 μm.

4. Discussion In this study, we showed that (1) MSC-CM increased the migration of cardiac derived cells from human heart tissue fragments, (2) the migrating cell population shared strong similarities with resident stem cells, more specifically with CASCs, and (3) PDGF-AA was found to be a promising mediator to enhance the migration process. To determine the direct effect of soluble factors released by MSCs on human cardiac tissue fragments in vitro, we constructed a 3D collagen type I gel to culture cardiac fragments in the presence of MSC-CM. With approximately 85% of total collagen, collagen type I is the most abundant collagen type present in the normal as well as in the infarcted heart. Thus, by culturing cardiac fragments in collagen type I, we were able to mimic the in vivo cardiac molecular constitution as well as some aspects of the local cardiac milieu, even after MI. Therefore, this kind of model was ideally suited to directly assess the influence of MSC-CM on cardiac tissue cells in situ, while retaining a tissue viability of more than 60% for all conditions until the end of the follow-up. Applying this model, we illustrated that MSC secreted agents promoted the migration of CASCs in a shorter time span, over greater distances and at significant higher quantities compared to the innate migration response observed in control conditions. Moreover, by assessing the influence of different concentrations of PDGF-AA, we found a clear dose dependent effect with higher PDGF-AA concentrates inducing higher migrating responses. In order to identify the migrating cell population, these cells were isolated, ex vivo expanded and extensively characterized. A thorough flow cytometrical analysis revealed that these migrating cells expressed multiple cell–cell and cell–matrix interacting proteins,

such as CD49c (integrin α3), CD29 (integrin β1) and CD44 (H-CAM) [28,29]. The expression of these markers is illustrative of their migrating activities, although it could not be ruled out that they could be induced by ex vivo expansion of the cells. Despite the fact that stem cell surface markers like c-kit and CD34 were not detected by means of flow cytometry, migrating cells did possess typical stem cell characteristics. These included a clonogenicity of ~21% and a uniform expression of pluripotency associated genes like Oct-4, Nanog, Dppa-3, Tbx-3, c-Myc and Klf-4. These data are in accordance with our findings in CASCs illustrating a clonogenic potential of ~17% and expression of identical stem cell genes. Based on the absence of the MSC marker CD140b [30], the lack of adipogenic differentiation capacity and the preservation of ALDH expression, we concluded that the migrating cells were unlikely to be MSCs, but rather have a cardiac origin [31]. Although it is well described that ALDH activity can be utilized to isolate progenitor cells like MSCs from the human bone marrow, it is often not monitored during expansion in culture. Yet, recent studies illustrated that MSCs no longer display ALDH protein expression when cultured ex vivo [23,24]. Therefore, an elevated ALDH activity combined with the lack of c-kit expression provided additional evidence that these cells resembled our recently identified CASCs. Indeed, CASCs can be readily isolated from patients with IHD and display a significant myocardial differentiation potential which renders them a favorable candidate for cardiac repair. Similar to ex vivo expanded CASCs, migrating cells lacked the CD34 stem cell marker, but showed superior differentiation towards the cardiac lineage in comparison to c-kit+ CSCs, as shown by expression of cTnT and cTnI after co-culture with NRCMs [4].

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Fig. 5. Cardiomyogenic differentiation potential of migrating cells. Representative immunofluorescent stainings showing cTnT (red, A) and cTnI staining (red, B) on GFP+ migrating cells (MCs) (green, left panel) and GFP+ CASCs (positive control, green, right panel) after one week in co-culture with NRCMs. Nuclei were stained with dapi (blue). Scale bar, 20 μm.

The ALDH detection kit utilized in this study is also reported to identify cells expressing retinaldehyde dehydrogenase 2 (RALDH2) [32]. After injury or thymosin β4 (Tβ4) stimulation, epicardium derived cells can upregulate embryonic epicardial genes, such as Wilm's tumor

A

Control

1 (Wt1) [33] and RALDH2 [34]. As explained in our previous study, CASCs are possibly related to the pro-epicardial originating CFU-Fs described by Chong et al. [22]. Experiments performed at that time, led us to conclude that ALDH+ cardiac cells were likely to be part of the

10% FCS

MSC-CM

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Fig. 6. MSC-CM stimulates CASC chemotaxis in transwell migration assays. A) Images representative for CASCs that migrated in either LG-DMEM with 10% FCS (positive control), serumfree LG-DMEM (control) or MSC-CM. Scale bar, 200 μm. B) Graphs depicting the percentage of migrating CASCs for each condition. Migration in the positive control condition was used for normalization. *p b 0.05 compared to control.

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Fig. 7. PDGFRα expression and phosphorylation on CASCs. A) Western blot depicting PDGFRα expression (190 kDa) on MSCs (positive control), HUVECs (negative control), CASCs in culture, CASCs in MSC-CM and CASCs directly isolated from human atrial appendages (resident). B) Representative immunofluorescence image illustrating PDGFRα expression (red, left) on CASCs directly isolated from human atrial appendages. Nuclei were stained with dapi (blue). For the negative control, only secondary antibody labeling was performed (right). Scale bar, 20 μm. C) Confirmation of RTK results by western blotting for phosphorylation of the PDGFRα (tyr849) (190 kDa) on expanded CASCs after MSC-CM treatment. α-Tubulin (α-Tub) served as loading control.

epicardial CFU-F stem cell hierarchy [4]. Currently, there's a lack of adult epicardium specific markers to identify these CFU-Fs in the adult heart. Known epicardial markers, such as Wt1 and T-Box 18 (Tbx18), are only expressed in the heart during embryonic stages and therefore cannot be used on adult tissue. In vivo examination of the reactivation of Wt1 in the adult injured heart after priming with thymosin β4 could give more insight on these unanswered questions [33]. Analysis of the migration response of different doses of MSC-CM illustrated a dose dependent effect of 10 × vs. 30 × MSC-CM based on area measurements after 6 days. It appears that beyond 10 × concentrated MSC-CM, a saturation phase is generally reached, explaining why we were unable to detect more statistical significant differences between the different doses of MSC-CM. Furthermore, our 3D collagen model in combination with transwell migration assays also suggested that the observed migratory response was due to chemotaxis rather than chemokinesis. Interestingly, recent studies already proposed chemotaxis of cardiac progenitors in response to MSC-CM. Hatzistergos et al. showed that MSC transplantation reduced infarct size in female Yorkshire pigs by directly interacting with host cardiac progenitors, promoting their recruitment, proliferation and differentiation. This was corroborated by a 20-fold increase in the number of endogenous CSCs after two weeks follow-up [35]. In contrast to our data, they were unable to reproduce similar effects with MSC-CM, but argumented that the injection of a single dose of MSC-CM was perhaps insufficient to produce sustained effects. Although MSC-CM was prepared in a similar way as in this study, Hatzistergos et al. assessed the effect of MSCCM in vivo after transendocardial injection into the infarct/border zone, followed by reperfusion. Possibly, MSC-CM was (partly) washed out during reperfusion, explaining its lower effect. More importantly, our data are consistent with findings described by Nakanishi et al., who illustrated that MSC-CM not only promoted proliferation, differentiation and cytoprotection of rat cardiosphere derived cells in vitro, but also enhanced their migration in a chemotaxis chamber assay [36].

Thus far, no specific mediator for stem cell migration in hearts has been identified. Although RTK screening revealed a mild increase in phosphorylation of the EGFR, the PDGFRβ, the InsR and the AxlR upon treatment with MSC-CM, we were unable to reproduce these findings. Conversely, the PDGFRα was consistently more activated as evidenced by a substantial increase in its phosphorylation status. The PDGFR family of proteins are reasonable candidates to stimulate CASC migration as they are known to be involved in coordinated movement of various progenitor cells [37,38]. In this study, we found that not only ex vivo expanded CASCs, but also CASCs directly obtained from human cardiac tissue express the PDGFRα, making the in situ induction of their movement towards the injured zone by PDGFRα ligands an attractive option. However, the signaling mechanisms that are involved in PDGFmediated cell motility are not fully understood, but are thought to be regulated by key signaling molecules such as phospholipase C-γ (PLCγ), phospho-inositide-3 kinase (PI3K), AKT, Ras and Focal Adhesion Kinase (FAK), as reviewed by Anand-Apte and Zetter [39]. Since several independent studies provided evidence for the secretion of significant amounts of PDGF-AA, but not -AB or -BB, by bone marrow MSCs, we focused our interest on the PDGF-AA/PDGFRα signalization pathway [26,27]. Here, we illustrated that MSC-CM contained PDGF-AA ligand, which is in line with the findings of Salazar et al. [26]. We found that, at the same time point, 100 ng/ml PDGF-AA was necessary to reproduce migration effects similar to that of MSC-CM, albeit with some temporal delay. However, lower and physiologically more relevant concentrates of 0.1 and 1 ng/ml PDGF-AA could effectively enhance migration as well, although after a longer period of time. Nevertheless, our findings illustrated a clear dose responsive effect of PDGF-AA on migration. Furthermore, the PDGFR tyrosine kinase inhibitor Imatinib Mesylate as well as a PDGFRα neutralizing antibody significantly attenuated PDGF-AA as well as MSC-CM induced migration, although the impact of Imatinib on MSC-CM induced migration was less pronounced. This strongly suggests that the PDGF-AA/PDGFRα is a promising, but probably not the sole pathway involved in CASC

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Fig. 8. PDGF-AA is able to reproduce the migrating effects of MSC-CM. A) Graphs depicting perimeter measurements illustrative of the cells' migration distances in either control, MSC-CM, 0.1 ng/ml PDGF-AA, 1 ng/ml PDGF-AA or 100 ng/ml PDGF-AA. Conditions as indicated. Number of cells in function of their migration into zone A, B or C at day 8 (B) and day 13 (C). *p b 0.05, **p b 0.01, ***p b 0.001 compared to control at the same time point. Connecting bars with asterisks illustrate statistical significances between indicated conditions. D) Photomicrographs representing PDGF-induced migrating cells (MCs, upper panel) in culture after incubation with activated Aldefluor® or DEAB (negative control, lower panel). Phase contrast (left) and ALDH (middle) images were taken and merged (right). Scale bar, 200 μm.

migration. Therefore, a synergistic relation between PDGF-AA and (an) other factor(s) secreted by MSCs has to be taken into account. For instance, a potential role could also be ascribed for vascular endothelial growth factor (VEGF), as we detected a 4,9-fold activation of the VEGFR by MSC-CM. Although we were unable to reproduce these results, it is already described that MSCs secrete VEGF [27]. Interestingly, VEGF is structurally related to PDGF and not only binds its own receptor, but is also able to interact with both PDGFRs [21]. Indeed, Zisa et al. identified VEGF as a key therapeutic trophic factor in MSC mediated cardiac repair through the migration of progenitor cells [40]. Moreover, Tang et al. found that VEGF promoted recruitment of CSCs in vitro via the PI3K/AKT pathway [41]. Another interesting candidate is the transforming growth factor-β (TGF-β), which becomes rapidly activated after MI and is known to play a catalytic role in the differentiation

of fibroblasts to activated myofibroblasts [42,43]. TGF-β has also been reported to stimulate migration of various cell populations, like (myo) fibroblasts [44,45], but also dental pulp stem cell [46] and MSCs [47]. Although the presence of TGF-β in MSC-CM was not investigated in this study because it was not included on the RTK array, migrating cells displayed strong expression of CD105 (endoglin), which is part of the TGF-β receptor complex. So, based on these results, it is interesting to further examine other chemotactic agents, like VEGF and TGF-β, in order to clarify their roles in stem cell recruitment/migration. In addition, PDGF is known to induce the expression and secretion of matrix metalloproteinases (MMPs) in several cell types, in order to perform their invasion/migration in collagen [48]. Although it is established that MSCs secrete MMPs that could degrade collagen and thereby enhance cell motility [27], the results from this study show that cell

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Fig. 9. Imatinib Mesylate attenuates PDGF- and MSC-CM induced migration. A) Graphs depicting perimeter measurements illustrative of the cells' migration distances in either control, 100 ng/ml PDGF-AA or MSC-CM, with or without addition of 1 μM Imatinib. Number of cells in function of their migration into zone A, B or C at day 8 (B) and day 13 (C). Conditions as indicated. *p b 0.05, **p b 0.01, ***p b 0.001 compared to its corresponding condition without Imatinib.

migration can be reproduced by recombinant PDGF-AA, in the absence of MSC-secreted MMPs.

Disclosure statement The authors declare no conflicts of interest.

5. Conclusions To our knowledge, this is the first study to demonstrate increased migration of progenitor cells from human cardiac tissue by MSC-CM or specific factors released by MSCs. The prospect of being able to recruit resident CASCs, that are inherently programmed to reconstitute the damaged myocardium, from their niches towards the injured infarct zone, might achieve better results than forcing non-cardiac originating stem cells to differentiate into contractile myocytes. Comprehensive understanding in the processes of cardiac progenitor migration/recruitment can lead to the identification of a specific factor or an optimal cocktail combination of paracrine acting factors. Indeed, according to our results, PDGF-AA is a promising but presumably not the only factor involved in the observed migrating response, making further migration studies recommended. Still, these results definitely designate the relevance for future in vivo models examining the migrating behavior of cardiac progenitors after administration of MSC-CM, and factors like PDGF-AA. Hereby, local injection or catheter based delivery of specific factors into the myocardium is aimed to first of all, influence guided trafficking of CASCs towards the site of injury and secondly, to mediate cardiac repair by coordinated stimulation of cellular processes involved in self-renewal, proliferation and cardiomyogenic/vasculogenic differentiation. This approach might provide a more attractive clinical option for patients with cardiomyopathy, avoiding the risks of isolation, ex vivo expansion and transplantation of stem cells into patients' hearts. Sources of funding This work was partially funded by a Ph.D. grant of the Agency for Innovation by Science and Technology in Flanders (IWT), and partially by the Limburg Clinical Research Program (LCRP) UHasselt-Jessa-ZOL, supported by the foundation Limburg Sterk Merk, Hasselt University, Jessa Hospital and Ziekenhuis Oost-Limburg.

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