Isolation and Characterization of CD276+/HLA-E+ Human Subendocardial Mesenchymal Stem Cells from Chronic Heart Failure Patients: Analysis of Differentiative Potential and Immunomodulatory Markers Expression

Abstract

Mesenchymal stem cells (MSCs) are virtually present in all postnatal organs as well as in perinatal tissues. MSCs can be differentiated toward several mature cytotypes and interestingly hold potentially relevant immunomodulatory features. Myocardial infarction results in severe tissue damage, cardiomyocyte loss, and eventually heart failure. Cellular cardiomyoplasty represents a promising approach for myocardial repair. Clinical trials using MSCs are underway for a number of heart diseases, even if their outcomes are hampered by low long-term improvements and the possible presence of complications related to cellular therapy administration. Therefore, elucidating the presence and role of MSCs that reside in the post-infarct human heart should provide essential alternatives for therapy. In the current article we show a novel method to reproducibly isolate and culture MSCs from the subendocardial zone of human left ventricle from patients undergoing heart transplant for post-infarct chronic heart failure (HSE-MSCs, human subendocardial mesenchymal stem cells). By using both immunocytochemistry and reverse transcriptase-polymerase chain reaction (RT-PCR), we demonstrated that these cells do express key MSCs markers and do express heart-specific transcription factors in their undifferentiated state, while lacking strictly cardiomyocyte-specific proteins. Moreover, these cells do express immunomodulatory molecules that should disclose their further potential in immune modulation processes in the post-infarct microenvironment. Another novel datum of potentially relevant interest is the expression of cardiac myosin heavy chain at nucclear level in HSE-MSCs. Standard MSCs trilineage differentiation experiments were also performed. The present paper adds new data on the basic biological features of heart-resident MSCs that populate the organ following myocardial infarction. The use of heart-derived MSCs to promote in-organ repair or as a cellular source for cardiomyoplasty is a fascinating and challenging task, which deserves further research efforts.

Introduction

M esenchymal stem cells (MSCs) have been isolated from different sources, such as bone marrow (BM) [1,2] and adipose tissue [3,4], and are virtually present in all postnatal organs [5], as well as in embryo-associated (perinatal) tissues [6,7]. In recent years, multiple evidences highlighted their differentiative ability toward a number of mature cytotypes, therefore making them reliable candidates for cellular therapy. In addition, MSCs have also promising immunomodulatory features in vitro and in vivo, another favorable factor for their administration in a number of diseases [8 –10].

MSCs feature the expression of a reproducible set of markers that are consistently found in populations isolated from different organ sources. Indeed, these cells must express molecules, such as CD29, CD44, CD73, CD90, and type I major histocompatibility complex (type I MHC), while remaining negative for endothelial (CD31) and hematopoietic (CD34, CD45, and type II MHC) markers [11,12]. In addition, it has been shown that apart classical markers, MSCs can express diverse molecules depending on the tissue of provenance, and this should disclose further potential applications of these same cells [13,14]. For example, we [15] and others [16] demonstrated that some MSC populations can also express the CD117 molecule, receptor for the stem cell factor encoded by c-kit gene, at both protein and mRNA levels.

Myocardial infarction results in severe tissue damage, comprising a massive loss of cardiomyocytes, formation of scar tissue, and consequent remodeling of the ventricular wall, and eventually heart failure [17 –21].

Cellular therapy using different populations of MSCs or progenitor cells, mainly BM-derived ones, is currently underway for heart diseases, and also clinical trials are being performed to assess the feasibility of this therapeutic option with respect to whole-organ transplantation [22 –24]. Different cell types were used to obtain differentiation toward cardiomyocytes, such as skeletal muscle stem cells [25], embryonic stem cells [26], circulating mesenchymal progenitors [27], adult MSCs [28], perinatal stem cells [29 –31], and cardiac stem cells [19].

Indeed, several literature data are present regarding the time-limited results obtained with these approaches, mainly due to paracrine actions of infused cells rather proper differentiation to functional myocytes [32,33]. Moreover, safety issues on the potential tumorigenesis of these cells were recently reported, therefore, suggesting caution in the interpretation of the results and in the evaluation of the risk/benefit balance [34].

Moreover, moving on from data concerning animal models [35], there are several reports of the presence of resident progenitor populations in the human heart, which can have or not the features of MSCs [36,37]. This is an important aspect since these cells are present at the infarcted area, and can have a role in tissue remodeling and perhaps tissue regeneration processes. Therefore, a new way to consider cellular therapy for heart diseases should be the definition of the properties of these resident cells and the development of strategies that can help directing their behavior to reduce the outcomes of heart infarct.

In the current article, we report a novel isolation method that allowed to reproducibly obtain MSCs from the subendocardial zone of human left ventricle from patients undergoing heart transplant for post-infarct chronic heart failure. We demonstrated that these cells feature marker expression and differentiation potential similar to other MSC populations. In addition, these cells do express immunomodulatory molecules and cardiac transcription factors that should disclose their further potential in both immune modulation processes in the post-infarct microenvironment and potential to differentiate toward cardiomyocytes or other heart-resident cells.

Materials and Methods

Tissue samples and cellular isolation protocol

All samples were obtained after patients' informed consent and treated in accordance with the tenets of the Declaration of Helsinki and local ethical regulations.

Transmural left ventricle samples (n=6) were obtained immediately after heart transplant surgery for the treatment of post-infarct chronic heart failure. Tissue samples were taken from those used for our preceding work [38]. Samples were stored aseptically in transport buffer [38] and then cellular isolation started within 6 h. To perform cellular isolation, the samples were rinsed in warm Hank's balanced salt solution (HBSS) (Gibco) and then processed to extract endocardial endothelial cells with a standard collagenase incubation. After protease withdrawal and HBSS rinse, transmural sections were cut into pieces of appropriate dimensions and posed, with the subendocardial layer facing down, in standard culture medium, in six-well plates (Corning) precoated with gelatin, one cord piece for each well; and covered completely with culture medium. The isolation and subculture of cells were made using Dulbecco's modified Eagle's medium (DMEM) low-glucose (Sigma), supplemented with 10% fetal bovine serum (FBS) Gold (PAA), 1× nonessential aminoacids (NEAAs; Sigma), 1× antibiotic–antimycotic (Sigma), and 2 mM L-glutamine (Sigma). The isolation method made no use of proteases to detach cells from the embedding matrix. Therefore, based on the “mesenchymal” migratory capability of cells, heart fragments were left in the culture medium for 15 days, with medium change every second day. Cellular exit from the tissue and attachment to the surface of the tissue culture slide was monitored by phase-contrast microscopy. Finally, after 15 days of culture, the remnants of the tissue fragments were removed from the wells, and cells were cultured until reaching the confluence.

Cell culturing and passaging

After reaching confluence, primary cells were subcultured routinely in culture medium. Cellular detachment from tissue culture flasks has been performed using TrypLe Select (Invitrogen) instead of standard trypsin solution. Primary populations of human subendocardial MSCs (HSE-MSCs) were cultured for up to 15 passages. To establish freezing conditions for long-term storage of cells, complete medium with 10% dimethylsulfoxide (DMSO) showed the best results as deep-freezing medium in terms of replating efficiency and cellular survival. For characterization experiments described below, cells at different passages were used in order to ensure the maintenance of marker expression along the population doublings of cells. For immunocytochemical analyses, cells were plated in gelatin-coated eight-well chamber slides (BD Biosciences) and were subjected to immunochemistry when they reached 90% confluence. For RNA extraction, cells were cultured either in six-well tissue culture plates or in 25-cm² tissue culture flasks (Corning).

Immunocytochemical analysis

After culturing, cells grown in chamber slides were washed with phosphate buffer saline (PBS) and fixed in methanol for 20 min at −20°C. Air-dried slides were then stored at −20°C until use. For the immunocytochemical procedure, cells were permeabilized with 0.1% TritonX-100 in PBS (Sigma). After a subsequent rinse with PBS, slides were exposed to 0.3% hydrogen peroxide in PBS, blocked with 1% FBS in PBS, and incubated for 2 h with the primary antibody. The detection was performed using an avidin–biotin complex kit (LSAB2; Dako); 3,3-diaminobenzidine (DAB chromogenic substrate solution; Dako) was used as developer. Nuclear counterstaining was obtained using hematoxylin (Dako). Immunopositivity was scored using a semiquantitative approach. Three independent observers (R.A., G.L.R., and F.C.) evaluated the immunocytochemical results and semiquantified the percentage of positive cells for each specimen. Ten high-power fields were examined in each culture slide and counting of the cells was performed at 40× magnification. Negative controls were run simultaneously by omission of the primary antibody.

The antibodies used in the present study, with indications of the working conditions used, are listed in Table 1.

Table 1.

List of Antibodies Used in the Present Study

Antigen	Host	Manufacturer	Dilution	Clone/cat No.
Cardiac myosin heavy chain	Mouse	Upstate	1:100	2F4
CD13	Mouse	Chemicon	1:50	WM 15
CD31	Mouse	Dako	Prediluted	JC/70A
CD31	Mouse	Santa Cruz	1:100	JC70
CD34	Mouse	Dako	Prediluted	QBEnd-10
CD45	Mouse	Dako	Prediluted	2B11+PD7/26
CD68	Mouse	Dako	1:100	KP1
CD79	Mouse	Dako	1:100	JCB117
CD117	Rabbit	Stressgen	1:200	N/A (KAP-TK005)
CD117	Rabbit monoclonal	Epitomics	1:50	YR145
CD276 (B7-H3)	Rabbit	Santa Cruz	1:100	N/A (H-300)
CK-7	Mouse	Dako	1:100	OV-TL 12/30
CK-8	Mouse	Sigma	1:200	M20
CK-18	Mouse	Sigma	1:800	CY-90
CK-19	Mouse	Chemicon	1:100	RCK 108
Collagen II	Mouse	Merck Millipore	1:100	6B3
Connexin-43	Mouse	BD Laboratories	1:50	2
Connexin-43	Rabbit	Santa Cruz	1:100	N/A (H-150)
GATA-4	Mouse	Santa Cruz	1:200	N/A (H-112)
GFAP	Mouse	Dako	1:500	6F2
HLA-A/B/C	Mouse	Santa Cruz	1:50	D-2
HLA-DR	Mouse	Santa Cruz	1:50	TAL 1B5
HLA-E	Mouse	Santa Cruz	1:50	MEM-E/02
MyoD	Mouse	NeoMarkers	1:100	5.2F
α-SMA	Mouse	Dako	Prediluted	1A4
α-SMA	Mouse	Santa Cruz	1:100	4i221
Nestin	Mouse	BD Laboratories	1:50	25
Nestin	Mouse	Santa Cruz	1:100	10c2
NSE	Mouse	Dako	Prediluted	BBS/NC/VI-H14
Prol-4-hydroxylase	Mouse	Dako	1:50	c5b5
Vimentin	Mouse	Santa Cruz	1:100	V9
Von Willebrand factor	Mouse	Dako	1:50	F8/86
Von Willebrand factor	Mouse	Santa Cruz	1:50	F8/86

CK, cytokeratin; GFAP, glial fibrillar acidic protein; HLA, human leukocyte antigen; α-SMA, alpha-smooth muscle actin; NSE, neuron-specific enolase.

Immunohistochemistry

Paraffin-embedded sections were stained with immunohistochemical methods. Four-micrometer-thick sections were stained with a streptavidin-peroxidase kit (LSAB2 system peroxidase; Dako), using primary antibody versus Collagen II (Merck Millipore). Negative controls and human vimentin as positive controls were run simultaneously.

Total RNA extraction

Total cellular RNA was isolated using the Quick Prep Total RNA Extraction Kit (GE Healthcare) as described previously [39]. RNA yield was evaluated spectrophotometrically (A260/280) and RNA aliquots were stored at −80°C until use. Total RNA fractions were used for subsequent experiments only if the A260/A280 ratio exceeded 1.7.

Reverse transcriptase-polymerase chain reaction analysis

Qualitative reverse transcriptase-polymerase chain reaction (RT-PCR) was performed using Phusion High-Fidelity RT-PCR kit (Finnzymes). RT-PCR consists of two phases: retro-transcription where RNA is converted into complementary DNA (cDNA) and cDNA was amplified. After treatment with DNAse, 2 μg of RNA was added, oligo dT and oligo N, to select only mRNA from total RNA. Subsequently, 5 μL of 10× Phusion Buffer, 1 μL of dNTP mix, 1 μL of RNAse inhibitor, and 1 μL of AMV reverse transcriptase and RNAse/DNAse free water were added until to reach a final volume of 50 μL. The reaction comprised a reverse transcription step of 50 min at 42°C and an inactivation phase of 5 min at 92°C.

Subsequently, at 2 μL cDNA, 10 pM of specific primers, 4 μL of 5× Phusion Buffer, 0.4 μL of 10 mM dNTP, 0.6 μL of DMSO, 0.2 μL of Phusion DNA Polymerase, and water were added until to reach a final volume of 20 μL. The amplification reaction was performed according to five steps. The initial denaturation of 30 s at 98°C, followed by another denaturation step of 10 s at 98°C, the annealing phases of 30 s at specific-primer temperatures, the extension step of 30 s at 72°C, and final extension for 10 min at 72°C.

The primer pairs used in this study are listed in Table 2.

Table 2.

List of PCR Primers Used for the Present Study

Name	Accession number	Product Size	Forward primer	Reverse primer
ABCG2	NM_004827	255	5′-ATGGTGTATAGACGCCCTGA-3′	5′-GGGACAGGTATGTGAAAAGC-3′
Actin, beta	NM_001101	350	5′-AAACTGGAACGGTGAAGGTG-3′	5′-TCAAGTTGGGGGACAAAAAG-3′
CD29	NM_033666	186	5′-CTGATTGGCTGGAGGAATGT-3′	5′- TTTCTGGACAAGGTGAGCAA-3′
CD44	NM_00610	282	5′-TCTCAAGGGCGTAACTCTGG-3′	5′-GCCAATTCTACCAGGCTTGA-3′
CD73	NM_002526	308	5′-CCTGCTCAGCTCTGCATAAGTA-3′	5′-CCCTATTTTACTGGCCAAGTGT-3′
CD80	NM_005191	259	5′-AGGGCCTCCTTAGATCCCTA-3′	5′-TTAGCTGCCATGAGATGTGC-3′
CD86	NM_175862	250	5′-TCCTGGCTGAGAGAGGAAGA-3′	5′-AGACTGCCCCATCCCTTAGT-3′
CD90	NM_006288	265	5′-TTTGGCCCAAGTTTCTAAGG-3′	5′-AGATGCCATAAGCTGTGGTG-3′
CD105	NM_000118	179	5′-TCCAGCACTGGTGAACTGAG-3′	5′-TGTCTCCCCTGCCAGTTAGT-3′
CD117	NM_000222	268	5′-ACTTCAGGGGCACTTCATTG-3′	5′-ACGTGGAACACCAACATCCT-3′
CD133	NM_006017	255	5′-GCATGCAAAAGCCATCATAG-3′	5′-ATCCATGCTGGACACCAGA-3′
CD166	NM_001627	283	5′-TGGTGTGGGAGATCAAAGGT-3′	5′-TGTGGCTGCCATTAAACAAG-3′
Connexin-26	NM_004004	215	5′-ACTGTGGTAGCCAGCATCG-3′	5′-AGGCTGAAGGGGTAAGCAA-3′
Connexin-32	NM_000166	218	5′-TCAGTGAGGAGGGATGTGG-3′	5′-TGGGGACTAGAGGCAGAGG-3′
Connexin-40	NM_005266	203	5′-GTGTGTGTGTGGGTGCTGA-3′	5′-GATGGGCAGGTGAGTCAGA-3′
Connexin-43	NM_000165	240	5′-CTTCAAGCAGAGCCAGCAG-3′	5′-TACCCCATACACCCCCAGT-3′
Connexin-45	NM_005497	234	5′-GCCAACATGGCAAAACTGT-3′	5′-CCTGGTTCAACAAGCCAAC-3′
Desmin	NM_001927	191	5′-AGCAGGGTGTTGGGATACTG-3′	5′-AGCCCCTGCTTTCTAAGTCC-3′
FABP4	NM_001442	252	5′-CATCAGTGTGAATGGGGATG-3′	5′-GTGGAAGTGACGCCTTTCAT-3′
GATA-4	NM_002052	270	5′-CCAGAGATTCTGCAACACGA-3′	5′-ATTTTGGAGTGAGGGGTCTG-3′
GATA-5	NM_080473	259	5′-GAATGGCCGGTGATGTATGT-3′	5′-TGAAGCTGATGCCAGACAAC-3′
GATA-6	NM_005257	259	5′-ACTAACCCACAGGCAGGTTG-3′	5′-GGTACAAAACGGCTCCAAAA-3′
HLA-A	NM_002116	262	5′-TGGGACTGAGAGGCAAGAGT-3′	5′-ACAGCTCAGTGCACCATGAA-3′
HLA-DR-B1	NM_002124	349	5′-GCACAGAGCAAGATGCTGAG-3′	5′-AGTTGAAGATGAGGCGCTGT-3′
HLA-E	NM_005516	245	5′-CAAGGGCCTCTGAATCTGTC-3′	5′-CGTGTTAGCCAGGATGGTTT-3′
HLA-F	NM_001098479	202	5′-TGGAGTTGCTCCGCAGATA-3′	5′-TCCACAAGCTCTGTGTCCTG-3′
ISL-1	NM_002202	360	5′-TCAAGAAGTCTGAAGCGACT-3′	5′-AAGACCACCGTACAACCTTT-3′
Keratin 8	NM_002273	216	5′-TCTGGGATGCAGAACATGAG-3′	5′-AGACACCAGCTTCCCATCAC-3′
Keratin 18	NM_000224	263	5′-CTGCTGCACCTTGAGTCAGA-3′	5′-GTCCAAGGCATCACCAAGAT-3′
Keratin 19	NM_002276	295	5′-ATGAAAGCTGCCTTGGAAGA-3′	5′-CCTCCAAAGGACAGCAGAAG-3′
Leptin	NM_000230	193	5′-CCAGATCCTCACAACCACCT-3′	5′-CTCCCAAAGTGCTGGGATTA-3′
MEF2C	NM_002397	430	5′-AGGACCCCCAAATGTCACT-3′	5′-AGCGGCAGCCTTTTACAAT-3′
Myocardin	NM_153604	193	5′-CTCGGCTTCCTTTGAACAAG-3′	5′-CTTCCCAGAGAATCCATCCA-3′
MDR-1	NM_000927	425	5′-ACAAAGCGCCAGTGAACTCT-3′	5′-TCACAGGCAGTTTGGACAAG-3′
MYBPC3	NM_000256	300	5′-CAGCAAGCAGGGAGTGTTG-3′	5′-GAGGGGTTTCCCCAACTTC-3′
MYL-2	NM_000432	250	5′-CAAGGAGGAGGTTGACCAGA-3′	5′-CAAAGAAGATGGAGGTGGA-3′
Nkx 2.5	NM_004387	450	5′-CATGGTATCCGAGCCTGGTA-3′	5′-GAGCTCAGTCCCAGTTCCAA-3′
NANOG	NM_024865	209	5′-CTCCATGAACATGCAACCTG-3′	5′-CTCGCTGATTAGGCTCCAAC-3′
Nestin	NM_006617	275	5′-TATAACCTCCCACCCTGCAA-3′	5′-AGTGCCGTCACCTCCATTAG-3′
Osteocalcin	NM_199173	296	5′-AGAGTCCAGCAAAGGTGCAG-3′	5′-GCAAGGGGAAGAGGAAAGAA-3′
Periostin	NM_006475	185	5′-TGGAGTTAGCCTCCTGTGGT-3′	5′-ACAAGGCTCGGTCTTTTCAA-3′
RYR-2	NM_001035	332	5′-CCCCATATGCTCCTGCTATT-3′	5′-CTGATCACAGGTGGCTGAAA-3′
TNNI3	NM_000363	244	5′-TGACCTTCGAGGCAAGTTT-3′	5′-CAGGAAGGCTCAGCTCTCA-3′
TNN3K	NM_015978	221	5′-CATTTTCATTCTTGCCGAAA-3′	5′-CACAAATCCAAAGCCTGCTA-3′

FABP4, fatty acid binding protein 4; ISL-1, Islet-1; MYBPC3, myosin binding protein C; MYL-2, myosin regulatory light chain 2; Nkx 2.5, NK2 transcription factor related 5; RYR-2, ryanodine receptor 2; TNNI3, troponin I type 3; TNN3K, TNNI3 interacting kinase.

Semiquantitative evaluation of mRNA expression

To determine quantitative differences in the expression of mRNAs of osteocalcin as well as periostin in osteogenic-differentiated versus undifferentiated HSE-MSCs, we performed densitometry analysis on multiple RT-PCR experiments, using cells at different culture passages, with subsequent normalization of the band intensities for a housekeeping gene (β-actin). Briefly, after total RNA extraction and DNAse treatment, reverse transcription was performed as stated earlier. The amplification reaction was performed according to five steps. The initial denaturation of 30 s at 98°C, followed by another denaturation step of 10 s at 98°C, the annealing phases of 30 s at specific-primer temperatures, the extension step of 30 s at 72°C, and final extension for 10 min at 72°C. The products of interest were visualized on 2% agarose gels and stained with SYBR safe (Invitrogen), and densitometry was performed in order to calculate the normalized band intensity for each PCR product at the different experimental conditions. Kruskal–Wallis test was used to assess the statistical significance of differences with respect to cellular differentiation.

Semiquantitative multiplex PCR analysis

To confirm the quantitative differences in the expression of fatty acid binding protein 4 (FABP4) mRNA following adipogenic differentiation, we used the protocol described by Spencer and Christensen [40] with minor modifications [39]. Briefly, after reverse transcription, we amplified the genes for β-actin and FABP4 for a range of cycles starting from the minimum number necessary to visualize the product on 2% agarose gel. The cycles were as follows: the initial denaturation of 30 s at 98°C, followed by another denaturation step of 10 s at 98°C, the annealing phases of 30 s at specific-primer temperatures, the extension step of 30 s at 72°C, and final extension for 10 min at 72°C. The cDNA of FABP4 was coamplified with β-actin cDNA over a range of cycles in the exponential phase of the PCR (maintaining the intensity of the band of β-actin constant by adding the related primers at the appropriate number of cycles before the end of the reaction). After 2% agarose electrophoresis, and SYBR safe staining, the bands corresponding to the FABP4 amplification product were quantified using the 1D Scan EX software and, after normalization for those of β-actin, the intensities were plotted as function of cycle number. GraphPad Prism software package was used to calculate the exponential regression equations fitted to the curves. Calculation of the R ² values was performed to ensure that the range of cycles used was in the exponential phase of the amplification reaction.

Clonogenicity assays

To give a formal demonstration of the self-renewal capability of HSE-MSCs, a limiting dilution method was applied [41]. Briefly, cells at different culture passages were plated as single cells into each well of a 96-well plate. The addition of a single cell per well was confirmed by phase-contrast microscopy. After 2 weeks in culture, with medium change each second day, the presence of clones was assessed by phase-contrast microscopy.

Osteogenic differentiation

Differentiation of cells was performed by culturing HSE-MSCs at different passages in osteogenic differentiation medium, mainly based on previous reports [15]. Briefly, culture medium was supplemented with 50 μM ascorbate-2-phosphate (Sigma), 10 mM β-glycerophosphate (Sigma), 0.1 μM dexamethasone (Sigma), 10% fetal bovine serum (FBS) (PAA), 1× NEAAs (Sigma), and 1% antibiotic–antimycotic (Sigma). Cells were cultured in six-well tissue culture plates for 3 weeks and medium was replaced every second day. The formation of cell clusters resembling direct ossification was monitored by phase-contrast microscopy along culturing. Controls included HSE-MSCs cultured in normal growth medium for 3 weeks to monitor the eventual spontaneous formation of bone-like nodules.

Adipogenic differentiation

Differentiation of cells was performed by culturing HSE-MSCs at different passages in adipogenic differentiation medium, mainly based on previous reports [15]. Culture medium was supplemented with 0.5 mM isobutylmethylxanthine (Sigma), 1 μM dexamethasone (Sigma), 10 μM insulin (Sigma), 200 μM indomethacin (Sigma), 10% FCS (PAA), 1× NEAAs (Sigma), and 1% antibiotic–antimycotic (Sigma).

Cells were cultured in six-well tissue culture plates for 3 weeks, and medium was replaced every second day. The formation of cytoplasmic lipid vacuoles was monitored by phase-contrast microscopy along culturing. Controls included HSE-MSCs cultured in standard growth medium for 3 weeks to monitor the spontaneous formation of lipid vacuoles.

Chondrogenic differentiation

Differentiation of cells was performed by seeding HSE-MSCs into alginate beads, using slight modifications of previously published protocols [42].

Briefly, HSE-MSCs were resuspended in sodium alginate (Sigma-Aldrich) (4×10⁶ cells/mL at a final concentration of 1.2% sodium alginate in sterile physiologic solution). Beads were formed by slowly dispensing droplets of the alginate cell suspension from a 22-gauge needle syringe into a 100 mM calcium chloride (CaCl₂) solution. After washes with 0.15 M sodium chloride, the beads were rinsed with DMEM. Then, beads were cultured either in standard growth medium (controls) or chondrogenic medium, prepared using published protocols with slight modifications (DMEM supplemented with 1% FBS, 6.25 μg/mL insulin, 10 ng/mL transforming growth factor beta 1, 50 nM ascorbate-2-phosphate, 1% antibiotic–antimycotic, and 1× NEAAs) [43]. Beads were maintained in culture for 3 weeks, with medium changes every second day. For fixation and paraffin embedding, beads were processed as previously described [42]. The beads were fixed in 4% paraformaldehyde and 0.1 M cacodylate buffer (pH 7.4) with 10 mM CaCl₂ for 4 h at 20°C and then washed overnight at 4°C in 0.1 M cacodylate buffer (pH 7.4) containing 50 mM barium chloride. The beads were standard dehydrated through alcohols and xylene and embedded in paraffin. Sections (6 μm) were processed for histology (Alcian blue and nuclear fast red staining) and immunohistochemistry (IHC).

Histochemical staining

To demonstrate the acquisition of the osteogenic phenotype, the Alizarin Red S staining was performed. Briefly, cells were fixed in 4% paraformaldehyde and stained with 1% solution of Alizarin Red S (Sigma). Stained cells were rinsed with water for three times to remove excess stain, and then photographed at the photomicroscope.

To demonstrate the adipogenic differentiation, cells were stained with Oil Red O (Sigma), and photographed at the photomicroscope. After medium aspiration, a brief wash was performed with PBS. Cells were fixed with 10% formalin (Sigma) for 30 min at room temperature, followed by subsequent washes with distilled water and 60% isopropanol. Oil Red O working solution was added to the cells for 5 min, followed by four washes (5 min each) with distilled water. The wells were viewed and photographed using an inverted phase-contrast microscope. Following a further step of counterstaining (Meyer's hematoxylin, 1 min), lipid vacuoles appeared red and nuclei appeared blue.

To demonstrate the acquisition of the chondrogenic phenotype, the Alcian blue–nuclear fast red staining was performed. Cells cultured in alginate beads were formalin fixed and paraffin embedded using standard protocols. Sections were de-paraffinized and hydrated to distilled water; then, Alcian blue (Sigma-Aldrich) solution [1% Alcian blue 8GX in 3% acetic acid (pH 2.5)] was added for 30 min. After washing, nuclei were counterstained in nuclear fast red (Sigma-Aldrich) solution (0.1%). Sections were then dehydrated and mounted for observation.

Results

Cellular isolation and culturing

The novel cellular isolation protocol described in the “Materials and Methods” section allowed to reproducibly isolate, from each heart specimen, a population of fibroblast-like cells, which were subcultured giving cells at various tissue culture passages. Figure 1 shows a panel of representative phase-contrast micrographs of HSE-MSCs at different culture passages. Panels are representative of what obtained with different cell lines isolated from different patients' samples. Cells at first passage (Fig. 1A, C, and D) showed a typical mesenchymal morphology, with cell–cell contacts and processes extending between adjacent cells. Confluent cells showed a typical arrangement in parallel rows. Cells at later passages (Fig. 1B) maintained their morphological features and steadily grew in the standard culture medium. Cells were routinely passaged and maintained in culture up to 15th passage, with a mean population doubling time of 6.2 days. Cultured cells were routinely freezed, stored in liquid nitrogen, and defrosted at different passages, thus demonstrating their ability to survive deep freezing and therefore their long-term storability (not shown).

FIG. 1.

Light microscopic micrographs of HSE-MSCs in monolayer culture at different culture passages. Cultured cells assumed a polymorphic, fibroblast-like morphology, which was maintained throughout the passaging process. The panels show cells at passage 1, prior to reach confluence (A); confluent cells at passage 4 (B); and confluent monolayer at passage 1 (C, D). Magnification (A, B)×10; (C, D)×20. Scale bar=100 μm. HSE-MSCs, human subendocardial mesenchymal stem cells.

Immunocytochemistry analysis of MSC markers

The initial characterization of HSE-MSCs was performed by immunocytochemistry (ICC) analysis, on triplicate samples of cells derived from different heart specimens, in order to limit the extent of subjective differences. Figure 2 shows a representative result obtained in the analysis of cells for a panel of markers previously used to study mesenchymal populations. Present data show that these cells feature the expression of structural molecules as alpha-smooth muscle actin (α-SMA) and nestin (Fig. 2D, E). Nestin is an intermediate filament, characterized in the neuroectodermic lineage as precursor of neurofilament proteins. Its expression in MSCs and also pancreas-specific progenitors has been recognized by different authors [15,44]. In addition, CD117 (also known as c-kit, receptor of the stem cell factor) is highly expressed by these cells (Fig. 2A and Supplementary Fig. S1A, B; Supplementary Data are available online at www.liebertpub.com/scd). CD117 has been characterized as a key marker in hematopoietic stem cells (HSCs), but its expression has been reported in various populations of adult- and fetal-derived MSCs [16,45]. As reported for other mesenchymal populations, HSE-MSCs are negative for CD34 (Fig. 2B), while show an intense expression of connexin-43, a key molecule in the formation of gap junctions in both the embryo [46] and the human heart [47] (Fig. 2C). We also determined the expression of a key molecule of the differentiated cardiomyocyte phenotype as cardiac myosin heavy chain. As visible (Fig. 2F and Supplementary Fig. S1C, D) this molecule is expressed by HSE-MSCs, showing a diffuse weak cytoplasmic staining and a nuclear envelope localization, which has been already demonstrated for several members of both actin and myosin families [48]. Our data show for the first time that also cardiac-specific myosin heavy chain (encoded by MYH1 gene) can be expressed at the nuclear level, and also suggest that in undifferentiated cells, this molecule appears not engaged in the formation of sarcomeric structures, which are typical of the differentiated working myocardium.

FIG. 2.

Representative panels of immunocytochemical detection of mesenchymal markers on HSE-MSCs. The isolated cells showed a strong positive signal for CD117 (c-Kit, A), Connexin-43 (C), α-SMA (D), and Nestin (E). Cardiac myosin was amply expressed by cultured cells, with clear nuclear envelope positivity (F). CD34 was not detected (B). Magnification ×20. Scale bar=100 μm. α-SMA, alpha-smooth muscle actin. Color images available online at www.liebertpub.com/scd

Another molecule that is involved in the developmental pathways of the heart is GATA-4, which we demonstrated to be expressed in other mesenchymal populations. As shown in Fig. 3A and in Supplementary Fig. S2, HSE-MSCs are also positive for the expression of this molecule, which showed a cytoplasmic/perinuclear localization. The distribution of this molecule outside of the nucleus is consistent with the undifferentiated phenotype of HSE-MSCs, where in the absence of a specific differentiation stimulus, the transcription factor is expressed but is maintained in inactive form outside of the nucleus, therefore being unable to transactivate cardiomyocyte-specific promoters [49].

FIG. 3.

Representative panels of immunocytochemical detection on HSE-MSCs. The isolated cells showed a strong positive signal for GATA-4 perinuclear (A), CD68 (B), and CD276 (B7-H3, F). Cells showed a light and diffuse staining for HLA-A/B/C (C) and HLA-E (E). HLA-DR (D) was not detected. Magnification ×20. Scale bar=100 μm. HLA, human leukocyte antigen. Color images available online at www.liebertpub.com/scd

We further wanted to analyze the expression of molecules that give MSCs hypoimmunogenicity and/or tolerogenicity, a feature that should be useful for the application of heart-derived cells in cellular therapy applications. We demonstrated that HSE-MSCs are strongly positive to CD68, which is classically known as a macrophage-specific antigen [50], but for which extra-myeloid expression has been demonstrated by us and others [51,52]. It is widely acknowledged that MSCs feature reproducibly a pattern of expression of MHC molecules in which type I MHC [class Ia human leukocyte antigen (HLA), namely, HLA-A, -B, and -C] are expressed, while type II MHC (class II HLA-D molecules) are not [9,53]. We demonstrated that HSE-MSCs do express class Ia type I MHC, while are negative for HLA-DR (type II MHC). In addition, we demonstrated for the first time in heart-derived MSCs the expression of two members of the class Ib type I MHC molecules. Figure 3E shows the diffuse staining of HLA-E in HSE-MSCs. HLA-E is a nonclassical type I MHC molecule that has been implicated in the selective binding to NK cells exerting immunomodulatory functions in diverse cell types [54,55]. In addition, we demonstrated for the first time that HSE-MSCs do express B7-H3 (CD276) (Fig. 3F). This molecule is a member of the B7 family of costimulatory molecules for which coinhibitory roles have been demonstrated [56 –58]. Other members of the family with similar immunomodulatory potential have been characterized in BM-MSCs (as B7-H1 and -H4) [9], but this is the first demonstration of the expression of B7-H3 in heart-derived MSCs.

Table 3 shows the global results of the ICC analysis of HSE-MSCs. The cells did not express endothelial-specific markers (CD31 and vWF) or hematopoietic ones (CD34, CD45, and CD79) [38]. These data are in good agreement with those reported in literature regarding the expression pattern of MSCs. HSE-MSCs did express CD13 (aminopeptidase N), a surface antigen typically expressed in endothelial cells and monocytes, whose expression has been characterized in MSCs populations by us and others [15,59]. Also, vimentin was expressed in HSE-MSCs. This is an intermediate filament found in a broad number of cells of mesenchymal origin, as well as historically in marrow stromal cells [60], and in MSCs from perinatal tissues [15]. Vimentin is not the only type of intermediate filament to be expressed by HSE-MSCs: a new and potentially interesting observation is the positivity of cells for the expression of cytokeratins (CKs) (namely, CK8 and CK18). These epithelial markers have not been characterized previously in heart-derived MSCs, while we and others demonstrated their expression in Wharton's jelly–derived MSCs (WJ-MSCs) and in situ in the umbilical cord [15,61].

Table 3.

Immunocytochemistry Results of Marker Expression by Human Subendocardial Mesenchymal Stem Cells

Cardiac myosin	+++
CD13	+
CD31	−
CD34	−
CD45	−
CD68	+++
CD79	−
CD117	+++
CD276 (B7-H3)	+++
CK-7	−
CK-8	++
CK-18	+
CK-19	−
Connexin-43	+++
GATA-4	+++
GFAP	+++
HLA-A/B/C	++
HLA-DR	−
HLA-E	+++
MyoD	−
α-SMA	+++
Nestin	+++
NSE	+
Prol-4-Hydroxylase	+++
Vimentin	+++
Von Willebrand factor	−

Results of the immunocytochemical analysis are represented semiquantitatively.

Putative neural markers have been discovered to be expressed also in MSCs of different origin. Therefore, we aimed to determine their expression also in HSE-MSCs. Nestin is a neural filament protein in neural precursor cells, which is replaced by neurofilaments upon neuron differentiation. We already demonstrated its expression in perinatal MSCs [15], and nestin+MSCs constitute a particular perivascular population in the BM niche, where they are strictly associated with HSCs [62]. Table 3 shows that, besides nestin, undifferentiated HSE-MSCs did express neuron-specific enolase (NSE) and glial fibrillar acidic protein (GFAP). NSE is a glycolytic enzyme present in neurons and neuroendocrine cells, whose levels may increase in case of neural damage [63]. Previous studies demonstrated that MSCs from umbilical cord do express NSE in the undifferentiated state and are able to upregulate its protein levels following neural differentiation protocols [15,64]. GFAP is an intermediate filament protein expressed in glial cells, whose expression has been demonstrated also in undifferentiated BM-MSCs [65] and WJ-MSCs [15,64]. In addition, cells are negative for the mature muscle transcription factor MyoD, while being extensively positive for cardiac myosin heavy chain, as outlined previously. The expression of another mesenchymal marker, namely, prolyl-4-hydroxylase, known as a fibroblast marker, strongly suggests the presence of an active collagen production by these cells. Further experiments have shown that alongside with passages, cultured cells were morphologically and phenotypically similar to the parental cells (not shown).

Qualitative RT-PCR analysis of gene expression

To confirm and extend the data obtained by ICC analysis, we performed total RNA extraction, followed by retrotranscription and specific amplification of gene fragments by qualitative RT-PCR. Figure 4 shows two representative electrophoretic gels of HSE-MSC gene expression pattern. As visible, the GATA-4 and GATA-6 factors are expressed by the cord cells. The expression of GATA-4 at the RNA level confirms the datum obtained by immunocytochemistry. GATA-6 is also involved in the developmental processes of human heart [49]. In addition, HSE-MSCs do express Mef2C transcription factor. This molecule is involved in the cardiogenic differentiation process, and therefore its expression in undifferentiated mesenchymal cells may indicate the possibility for these cells to undergo myocyte differentiation. RT-PCR experiments allowed also assessing the expression of other “core markers” of human MSCs: CD29, CD44, CD73, CD90, and CD105. Table 4 summarizes the global results of qualitative RT-PCR analyses. As shown, we first extended the analysis for MSC markers, assessing the expression of CD106, CD117 (confirmed by ICC), CD133, and CD166. Moreover, we further investigated the expression of molecules involved in the immune recognition of MSCs by the immune system of the host. It has been reported that in “in vivo” settings the host response toward differentiated stem cells was related to the expression of B7 costimulatory molecules, such as CD80 (B7-1) and CD86 (B7-2). We determined by RT-PCR (Table 4) that HSE-MSCs did lack CD86, while expressing CD80. Previous reports strongly suggest that this feature should be of key importance in the instauration of a tolerogenic response of the host to avoid transplant rejection [66,67]. This, together with the data presented earlier on the expression of HLA-E and CD276/B7-H3, clearly points to the potential immunomodulatory effects that can be provided by these cells upon transplantation. In addition, we determined by RT-PCR (see Table 4) that another class Ib type I MHC molecule, namely, HLA-F, is expressed by HSE-MSCs. For this molecule, immunomodulatory functions at the fetomaternal interface have been proposed [68]. In addition, recent studies demonstrated that HLA-F may have additional functions independent from antigen peptide loading [69,70]. Connexin molecules are also of key importance in the functional phenotype of cardiac cells. Apart connexin-43, which is typically expressed by cardiomyocytes [71], RT-PCR allowed demonstrating also the expression of connexin-26 and connexin-45. In particular, the latter is typically expressed by specific myocardium cells [72], and its expression has not been demonstrated in mesenchymal populations so far. Another interesting datum obtained with RT-PCR analysis is the characterization of expression of other myocardial-specific transcription factors. As depicted in Table 4, these cells do express cardiomyocyte-specific factors, such as Islet-1 (ISL-1), Myocardin, and NK2 transcription factor related 5 (Nkx 2.5) [73]. The presence of all of these myocyte-specific factors does not directly imply that undifferentiated cells have the traits of a functional contractile cell. We already demonstrated for GATA-4 that the factor is kept in a cytoplasmic location, thus being inactive in HSE-MSCs. Further data corroborate this hypothesis. In fact, HSE-MSCs resulted negative for strictly myocardial-specific markers, such as myosin regulatory light chain 2 (MYL-2, ventricular/cardiac muscle isoform), myosin binding protein C (MYBPC3, cardiac), ryanodine receptor 2 (RYR-2), troponin I type 3 (TNNI3, cardiac), and TNNI3 interacting kinase (TNNI3K), thus suggesting the lack of a proper terminal differentiation without the presence of specific inducers in the culture medium.

FIG. 4.

Representative panel of RT-PCR analysis of HSE-MSCs. Cultured cells showed the expression of markers of MSCs (CD29, CD73, CD90, CD105, CD133, and CD166, panels A,B) and transcription factors involved in the cardiomyocyte differentiation (GATA-4, -6, and MEF2C, panels A,B). In addition, cells were positive for intermediate filaments, such as desmin and CK-18. B7 costimulators showed alternative expression since CD80 was expressed while CD86 was not detected. M: 50 bp ladder. RT-PCR, reverse transcriptase-polymerase chain reaction; HSE-MSCs, human subendocardial mesenchymal stem cells.

Table 4.

Expression of Markers of Different Lineages by Human Subendocardial Mesenchymal Stem Cells Assessed by RT-PCR

Name	Accession number	Expression in HSE-MSCs
ABCG2	NM_004827	+
Actin, beta	NM_001101	+
CD29	NM_033666	+
CD40	NM_001250	−
CD44	NM_00610	+
CD73	NM_002526	+
CD80	NM_005191	+
CD86	NM_175862	−
CD90	NM_006288	+
CD105	NM_000118	+
CD117	NM_000222	+
CD133	NM_006017	+
CD166	NM_001627	+
Connexin-26	NM_004004	+
Connexin-32	NM_000166	−
Connexin-40	NM_005266	−
Connexin-43	NM_000165	+
Connexin-45	NM_005497	+
Desmin	NM_001927	+
GATA-4	NM_002052	+
GATA-5	NM_080473	−
GATA-6	NM_005257	+
HLA-A	NM_002116	+
HLA-DR-B1	NM_002124	−
HLA-E	NM_005516	+
HLA-F	NM_001098479	+
ISL-1	NM_002202	+
Keratin 8	NM_002273	−
Keratin 18	NM_000224	+
Keratin 19	NM_002276	+
MEF2C	NM_002397	+
Myocardin	NM_153604	+
MDR-1	NM_000927	+
MYBPC3	NM_000256	−
MYL-2	NM_000432	−
Nkx 2.5	NM_004387	+
NANOG	NM_024865	+
Nestin	NM_006617	+
RYR-2	NM_001035	−
TNNI3	NM_000363	−
TNN3K	NM_015978	−

Qualitative RT-PCR analyses determined the expression of key markers of MSCs together with molecules responsible of the immunologic features of these cells.

RT-PCR, reverse transcriptase-polymerase chain reaction; HSE-MSCs, human subendocardial mesenchymal stem cells.

Clonogenicity assays

One of the key properties of stem cells is self-renewal, which allows to maintain a population of cells with stemness features alongside cell divisions. The clonogenicity ability of cells, that is, the ability to derive a cellular colony (clone) from a single cell, is a formal demonstration of self-renewal ability. To perform clonogenicity assay, cells at different culture passages were seeded in 96-well plates using the limiting dilution method, in order to deliver a single cell per well. After 15 days of culture, colonies were counted, and we obtained a cloning efficiency of 10% (not shown), in line with the previous results of us and others with MSCs [15,41]. Clonal cell lines were no further characterized.

Osteogenic differentiation of HSE-MSCs

Multipotent stem cells are able to differentiate toward multiple mature cell types. In particular, MSCs are defined as being able to undergo trilineage differentiation, toward osteoblasts, chondrocytes, and adipocytes.

To demonstrate HSE-MSC capacity do undergo osteoblast differentiation, cells were cultured for 3 weeks in osteogenic medium. Control cells were cultured for the same time period in standard growth medium. After 3 weeks, the formation of extracellular calcium deposits has been assessed by Alizarin Red S staining. As visible in Fig. 5B, D, and F, osteogenic-differentiated HSE-MSCs showed extensive areas of red staining, indicating the presence of calcium deposits in correspondence to de-novo-formed cellular aggregates (similar to bone nodules). On the contrary, undifferentiated control cells, cultured for the same time in standard growth medium (Fig. 5A, C, and E), did not show any positive staining, and maintained their fibroblastoid morphology.

FIG. 5.

Light microscopic demonstration of bone tissue formation by HSE-MSCs following osteogenic differentiation. When cultured in osteogenic medium, cellular morphology changed from a fibroblastic appearance (A, C, E) to a more cuboidal shape (B, D, F); in addition, cells were surrounded with an abundant matrix and formed mineralized nodules. The formation and deposit of a mineralized matrix in differentiated osteoblast-like cells (B, D, F) has been assessed by Alizarin Red S staining. Control cells cultured in standard medium did not show any staining (A, C, E). Magnification (A–D)×10; (E, F)×20. Scale bar=100 μm. Color images available online at www.liebertpub.com/scd

Moreover, the acquisition of an osteoblast-like phenotype was assessed by semiquantitative RT-PCR. As shown in Fig. 6, osteoblast-differentiated HSE-MSCs significantly upregulated the expression of osteoblast-specific genes (osteocalcin and periostin) with respect to undifferentiated cells (p=0.0159 for osteocalcin and p=0.0286 for periostin), thus confirming data from histochemical staining also at the molecular level.

FIG. 6.

Semiquantitative RT-PCR analysis of the expression of specific markers following osteogenic differentiation of HSE-MSCs. Following the osteogenic differentiation protocol, cells significantly increased the expression of both osteocalcin (A) and periostin (B) with respect to control cells cultured for the same time in standard growth medium.

Adipogenic differentiation of HSE-MSCs

To demonstrate their ability to differentiate toward multivacuolar adipocytes, HSE-MSCs were cultured for 3 weeks either in adipogenic medium or in standard culture medium (control cells). Figure 7 shows the results of the adipocyte differentiation of HSE-MSCs. Differentiated cells after 3 weeks of culture in adipogenic medium showed multiple intracellular vacuoles at the microscopic examination by phase-contrast microscopy (Fig. 7B), which were not present in undifferentiated control cells (Fig. 7A). Oil Red O staining, which specifically evidences the presence of neutral lipid vacuoles in cells, further confirmed morphological data. As visible in Fig. 7D, F, and H, differentiated cells showed a high number of intracellular lipid vacuoles of diverse sizes, resembling multivacuolar adipocytes. On the contrary, no positive staining was detected in control cells (Fig. 7C, E, and G).

FIG. 7.

Light microscopic demonstration of adipocyte differentiation with Oil Red O staining. HSE-MSCs cultured for 3 weeks in adipogenic medium showed variations in cellular morphology (B, D, F, H) and accumulation of neutral lipid vacuoles (demonstrated by Oil Red O staining in D, F, H) with respect to control cells. The latter (A, C, E, G) were cultured for the same time in standard culture medium, and retained the normal fibroblast-like morphology, without any positivity for the lipid-specific staining procedure (C, E, G). Magnification (A, B, E, F)×20; (C, D)×10; (G, H)×100. Scale bar=100 μm. Color images available online at www.liebertpub.com/scd

In addition, we performed the technique of semiquantitative multiplex PCR to assess the effects of differentiation protocol on the expression of the FABP4 (adipocyte) gene. As visible in Fig. 8A, we coamplified the PCR products corresponding to FABP4 and β-actin, over a range of cycles starting from 31 (the minimum required for the detection of FABP4, as determined in control experiments). Primers for β-actin were added after two cycles, in order to keep the reference PCR product in the exponential phase of PCR. As shown in Fig. 8B, the densitometry data obtained after normalization of band intensity for the reference gene were plotted to extrapolate the fitted exponential curve. R ² coefficients were calculated for both curves and confirmed the fitting of the curves with the exponential one. As visible, the expression of FABP4 was higher in differentiated versus undifferentiated cells, further confirming the acquisition of the adipocyte phenotype by HSE-MSCs.

FIG. 8.

Semiquantitative multiplex PCR analysis of the expression of FABP4 in adipogenic-differentiated cells with respect to undifferentiated controls. (A) Representative electrophoretic migration of the co-amplified products; (B) Representative plots of normalized data versus cycle numbers fit with an exponential curve for untreated and adipocyte-differentiated cells. M: 50-bp ladder. FABP4, fatty acid binding protein 4.

Chondrogenic differentiation of HSE-MSCs

To demonstrate their ability to differentiate toward chondroblasts, HSE-MSCs were seeded into alginate beads and cultured for 3 weeks either in chondrogenic medium or in standard culture medium (control cells). As visible in Fig. 9B, differentiated cells showed the presence of lacunae outside each cell, resembling mature chondrocytes. This feature was absent in alginate-seeded cells cultured in standard medium (controls; Fig. 9A). Moreover, we performed Alcian blue staining, which specifically evidences the deposits of glycosaminoglycans in the cartilage matrix. No positive staining was detected in control specimens (Fig. 9C, E) where a pale blue stain was homogeneously visible in alginate matrix. On the contrary, the matrix seeded with HSE-MSCs cultured in chondrogenic medium did show an intense blue staining over a large bead area (Fig. 9D, F).

FIG. 9.

Light microscopic demonstration of chondrogenic differentiation. HSE-MSCs cultured for 3 weeks in chondrogenic medium in alginate matrix showed variations in cellular morphology (appearance of rounded lacunae: B, arrows ) and accumulation of glycosaminoglycans (demonstrated by Alcian blue staining in D, F) with respect to control cells. The latter (A, C, E) were cultured for the same time as alginate-embedded cells in standard culture medium, without showing any morphological variation nor positivity for the specific staining procedure. Magnification (A, B) ×20; (C, D) ×10; (E, F) ×40. Scale bar=100 μm. Color images available online at www.liebertpub.com/scd

To further confirm the acquisition of the differentiated phenotype, we performed IHC for type II collagen, a specific marker of cartilage tissues [74,75]. As visible in Fig. 10, type II collagen was expressed in undifferentiated HSE-MSCs exclusively at the intracellular level. On the contrary, differentiated cells did show a stronger intracellular staining, and further positive staining on the cellular margin of the lacunae. This particular pattern strictly resembles the organization of collagens in vivo and in chondrocytes cultured in alginate matrix [42,75].

FIG. 10.

Immunohistochemical demonstration of collagen type II expression by chondrogenic-differentiated HSE-MSCs. HSE-MSCs cultured in alginate matrix with standard growth medium did not form lacunae-like structures around cells, and did express both vimentin (A) and, at lower levels, type II collagen (C). Alginate-seeded cells grown in differentiation medium showed the appearance of pericellular lacunae, and maintained the expression of both vimentin (B) and type II collagen (D). The latter appeared at higher levels than control cells, and interestingly it was also deposited at the edge of the lacunae structures (arrows) resembling its typical pattern in vivo in cartilage. Magnification: ×40. Scale bar=100 μm. Color images available online at www.liebertpub.com/scd

Discussion

Cellular therapy using MSCs has been proposed as a valid alternative to whole-organ transplantation for the management of diverse heart diseases. Despite the initial boost in the number of clinical trials using such approach, more data are needed to ensure the expected outcomes by cellular cardiomyoplasty. In addition, several literature data report the existence of heart-resident stem (progenitor) cells, which can be isolated from post-infarct hearts and may also be stimulated to modulate the processes downstream the initial tissue damage [76]. These cellular populations may be differentiated towards heart-resident cell types both in vitro and in vivo [77].

Also, cells with MSCs features can be isolated from human heart, and some recent data suggested that these populations may be very closely related to cardiac progenitor cells, perhaps residing in the same cellular lineage [78]. Since one of the outcomes of cellular activation following infarct is the recruitment of local or blood-borne progenitors to the lesion site, it should be worth to investigate the basic biological features of these cells to understand how to stimulate them to favor heart tissue regeneration, in absence of exogenous cell administration. This method should overcome the poor results that seem to hamper cellular therapy for heart diseases.

In this article, we present the results obtained with a novel isolation protocol of HSE-MSCs, from the infarct and peri-infarct zone of left ventricles. We used a nonenzymatic protocol that was applied to left ventricle transmural sections after the removal of the endocardial endothelial cells. This procedure exposes the subendothelial region, that is, the basement membrane underlying the endothelium and the well-vascularized loose connective tissue that bridges endothelium to myocardium [79]. Therefore, cells are left free to exit from the heart tissue (subendocardium and myocardium) based only on their migratory capability (which is a key property of all cells named as “mesenchymal”). Lack of lytic enzymes during cellular isolation avoids the undesired effects that have been reported previously; in fact, overdigestion may alter cellular function and viability [80,81]. As showed, the isolation protocol allowed to reproducibly isolate from all the specimens a population of fibroblastoid cells, which showed adherence to gelatine-coated tissue culture flasks and were maintained in culture using standard media we previously tested with other MSCs [15].

Cellular characterization was carried out by means of both ICC and RT-PCR. Apart assessing the expression of classical MSCs markers, we aimed to extend the characterization of these cells well over the classical markers that are currently used to define the mesenchymal phenotype. HSE-MSCs do express markers that belong to mature cell types that are of mesenchymal origin (vimentin and α-SMA), neuroectodermal origin (Nestin, NSE, and GFAP), and endodermal origin (GATA-4, GATA-6, CK-8, and CK-18). Such molecules are quite uncommon in heart-derived cells. This is particularly true for CKs; this is in our opinion a first evidence of the hidden properties of HSE-MSCs, which feature an undifferentiated phenotype resulting in the expression of markers of different lineages. Whether the expression of these markers might imply a differentiative ability toward cell types other than heart-resident ones is a hypothesis that goes over the scope of the present work. Interestingly, we demonstrated that HSE-MSCs do express at the protein level the myosin heavy chain, heart-specific isoform. We showed that subcellular localization of this protein is both in the cytoplasmatic and in the nuclear envelope region. It is well known that contractile filaments may have alternative subcellular localizations, in particular, in cell nuclei, where they exert different functions [48]. Nuclear actin is mostly found as a monomer, even if fibrillar structures have been also reported [48]. Nuclear myosins (six different types) have been located in the nuclear compartment where they mediate transcription by polymerases I and II, compartmentalization of chromatin, and many other functions [48]. To the best of our knowledge, this is the first report showing nuclear localization of the isoform coded by MYH1 gene in any cell type, and, in particular, in MSCs. Further studies are required to better define the implications of this nuclear expression for heart-derived MSC biology. In a very recent report, Asumda and Chase described the presence of nuclear actin, tropomyosin, cardiac troponin C, and cardiac troponin I in the nuclei of rat cardiomyocytes and the same proteins were detected in the nuclei of rat BM-MSCs upon coculture with cardiomyocytes [82]. The authors hypothesized a novel function of this calcium-sensing complex in nuclear signaling.

A key aspect of MSCs biology is their capacity to modulate host immune responses upon transplantation. This process may favor the engraftment of infused cells and promote their resistance to immunological scavenging by the host immune system. The regulation of immune response may involve different mechanisms, such as dendritic cell maturation, T-cell recognition, and promotion of lymphocyte tolerance or anergy [10]. Therefore, we assessed the expression of immunomodulatory molecules at both protein and RNA levels. We showed that HSE-MSCs do express MHC type I molecules (HLA ABC), while being negative for type II molecules (such as HLA-DR). This pattern was reproducibly characterized in MSCs populations of different origin [reviewed in Ref. 9], and can explain a number of favorable features with respect to immune recognition of MSCs. In fact, a low expression of class I MHC protects cells from NK-mediated lysis, such as that occurring in tumor cells where class I MHC are downregulated. Moreover, class II MHC are powerful alloantigens, which mediate recognition by alloreactive T-cells. To further detail the features of HSE-MSCs, we demonstrated for the first time that they do express also other class I MHC molecules, namely, HLA-E and HLA-F. Both are nonclassical type I MHC molecules (class Ib HLA). HLA-E and -F have been also implicated in tolerogenic processes occurring at the fetomaternal interface, together with other factors such as HLA-G [83 –88]. Recent studies demonstrated that HLA-F may have additional functions independent from antigen peptide loading [69,70]. Both HLA-E and HLA-F share a low genetic polymorphism and their primary structures are highly conserved in different species, for example, between primates [89].

In addition, we analyzed the expression of three molecules belonging to the B7 family of costimulators. HSE-MSCs did express CD80 (B7-1) while lacking CD86 (B7-2). It has been suggested that this pattern of expression of the two main constituents of the B7 family may result in favorable outcome from an immune modulation point of view [66]. In fact, CD80 may exert an inhibitory role on lymphocytes via CD152 (CTLA-4) binding, thus attenuating lymphocyte response [90,91]. This process may be enhanced in the absence of CD86, which binds CD28; further reports [67] suggested that CD86 blockade, in presence of intact CD80, results in alloantigen-specific tolerance. Another molecule that has been implicated in coinhibition mechanisms is CD276 (B7-H3). This molecule has been implicated in tumor evasion from immune response, but above all showed inhibitory effects on lymphocyte activation [56]. Its expression on HSE-MSCs should act in parallel to that of HLA-E in inducing lymphocyte downregulation. Further studies are needed to elucidate the role of these molecules in the post-infarction events taking place in human heart (e.g., for the evolution of inflammatory and remodeling processes) as well as for the possible therapeutic applications of heart-derived MSCs.

Finally, we aimed to characterize the expression of a number of heart-specific (and, in particular, cardiomyocyte-specific) markers in undifferentiated HSE-MSCs. This should be of particular importance if the use of these cells upon differentiation in contractile myocardial cells is prospected as a future application.

We demonstrated that HSE-MSCs do express Cx-43, which is known as the main connexin of the working myocardium, and Cx-45, which is characteristic of conduction cells of the specific myocardium, above all in the ventricles [72]. Interestingly, Zhang and coworkers reported a marked increase of these proteins in cardiac fibroblasts post-infarct, as a mechanism to maintain electrical coupling following myocardial damage [92]. This particular pattern of expression may suggest that HSE-MSCs may exert a similar function in maintaining the physiological properties in damaged myocardium. In addition, expression of Cx-26, together with CX-43 and -45, has been demonstrated in human heart valve interstitial cells [93].

Transcription factors intervene at different developmental points in the specification of cardiomyocyte phenotype. Their expression and nuclear translocation are key events in the establishment of a differentiated phenotype. In this work, we determined the expression of different transcription factors involved in muscle/heart development. ICC demonstrated that HSE-MSCs do express GATA-4, which is kept outside the nucleus, thus not performing cardiomyocyte-specific transactivation. By the same means, we demonstrated that the muscle-specific factor MyoD is not expressed in these cells. RT-PCR demonstrated that other cardiac-specific transcription factors are expressed. In fact, HSE-MSCs expressed Isl-1 mRNA. ISL-1 is a molecule belonging to the LIM homeodomain subfamily, which is essential in embryonic heart formation [73]. Another factor that was expressed in HSE-MSCs is Nkx 2.5. This homeoprotein is expressed in the precardiac mesoderm together with GATA-4 and GATA-6. These two factors lie upstream to Nkx 2.5 in the transcriptional regulatory cascade that affects cardiogenesis [94]. Another transcription factor that is expressed in HSE-MSCs is MEF2C. This is a key factor in cardiomyocyte differentiation and recent data demonstrated that transfection of mouse fibroblasts with four factors (comprising GATA-4 and MEF2C) results in a beating myocyte phenotype [95]. In addition, we demonstrated that another potent myogenic factor, myocardin, is expressed in HSE-MSCs. This molecule is involved, together with serum response factor, in the promyogenic differentiation of precursor cells [96]. Recent data from other groups support our hypothesis that the sole expression of these factors is not sufficient to elicit a proper cardiomyocyte differentiation program in absence of specific inducers (i.e., in undifferentiated cells). In fact, Armiñán and coworkers [28] demonstrated that MSCs can start a cardiac differentiation program upon nuclear translocation of both Nkx 2.5 and GATA-4.

Since HSE-MSCs are mainly positive for the expression of many cardogenic factors, with absence of phenotypical features of mature cardiomyocytes, we aimed to determine whether specific molecules of the cardiomyocyte phenotype were expressed in undifferentiated cells. RYR-2 is a key molecule involved in the outflow of calcium from the sarcoplasmic reticulum (SR). Spontaneous Ca(2+) release through RYR-2, a huge tetrameric protein, during diastole leads to a decrease in the SR Ca(2+) content [97]. This molecule is of key importance also for the large class of diseases known as channellopaties [98,99]. Other strictly myocardial markers, such as MYL-2 (slow cardiac myosin regulatory light chain 2, ventricular/cardiac muscle isoform), MYBPC3, TNNI3 (cardiac), and TNNI3K, were similarly not expressed by undifferentiated HSE-MSCs. In particular, MYL-2 is normally expressed in ventricular cardiomyocytes, and its downregulation has been linked to the development of heart failure [100]. MYBPC3 binds titin and myosin heavy chains in the C region of the A band in heart muscle sarcomere [101,102]. TNNI3 is the cardiac isoform of troponin I, which takes part in the formation of the calcium-sensitive regulatory complexes in the sarcomere, together with Troponins C and T and tropomyosins [103]. TNNI3K is a recently discovered kinase gene encoding a protein specifically interacting with TNNI3 [104].

Present data therefore suggest that HSE-MSCs do express a number of inactive transcription factors when kept in an undifferentiated state. This is also strongly suggested by the lack of highly specific myocardial proteins involved in the contractile phenotype of cardiomyocytes. The investigation of possible myocardial differentiation of HSE-MSCs is an intriguing possibility, which goes out of the scopes of the present work.

The demonstration of the adherence of HSE-MSCs to the general MSC phenotype was also given by their differentiability toward main mesenchymal tissues, namely, bone, cartilage, and adipose.

In conclusion, we demonstrated that the subendothelial endocardium, in patients affected by post-infarct chronic heart failure, contains a cellular population of MSCs that can be isolated by a novel nonenzymatic protocol, after removal of the endothelial cells. We demonstrated that these cells, apart classical MSC markers, do express new molecules belonging to a number of differentiated cytotypes, together with promising molecules whose immunomodulatory activity is recognized in vitro and in vivo. In particular, this is the first report demonstrating the parallel expression of HLA-E, HLA-F, and CD276 in MSCs derived from human heart. These cells feature several cardiac-specific transcription factors and lack most of the markers that are expected in mature cardiomyocytes. In addition, we demonstrated that these cells feature the expression of cardiac myosin heavy chain in an unusual nuclear location, thus adding a new potential member to the nuclear myosin group of proteins. Nuclear MYH1 functions in basic stem cell biology, and cardiomyocyte physiology or differentiation, may be hypothesized also on the basis of recent literature reports and deserve in our opinion more research efforts.

Defining the in vitro and in vivo biology of these cellular populations will be in our opinion a challenging task, for the potential role that these cells may have in the processes following myocardial infarction. Recent data shed new light on the roles of MSCs in tissue remodeling, infarct scar formation, maintenance of electrical coupling, and modulation of immune and inflammatory reactions occurring after myocardial damage. Once we have gained a sufficient knowledge on these processes and the contribution of heart-derived MSCs in vivo, perhaps new avenues will open on their use in therapy of chronic heart failure.

Footnotes

Acknowledgments

This work in part was supported by Fondazione S. Maugeri, Ricerca Corrente (A.D.S. and P.G.), and Istituto Euro-Mediterraneo di Scienza e Tecnologia (G.L.R.). The funders had no role in study design, data collections and analysis, decision to publish, or preparation of the article.

Author Disclosure Statement

Dr. La Rocca is member of the scientific board of Auxocell Laboratories, Inc. No competing financial interests exist for other authors.

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