ORIGINAL RESEARCH REPORT

STEM CELLS AND DEVELOPMENT Volume 00, Number 00, 2013  Mary Ann Liebert, Inc. DOI: 10.1089/scd.2013.0268

Hypoxia, Leptin, and Vascular Endothelial Growth Factor Stimulate Vascular Endothelial Cell Differentiation of Human Adipose Tissue-Derived Stem Cells Mohamed M. Bekhite,1,2,* Andreas Finkensieper,1,* Jennifer Rebhan,1 Stephanie Huse,1 Stefan Schultze-Mosgau,3 Hans-Reiner Figulla,1 Heinrich Sauer,4 and Maria Wartenberg1

The plasticity of human adipose tissue-derived stem cells (hASCs) is promising, but differentiation in vitro toward endothelial cells is poorly understood. Flow cytometry demonstrated that hASCs isolated from excised fat tissue were positive for CD29, CD44, CD70, CD90, CD105, and CD166 and negative for the endothelial marker CD31, and the hematopoietic cell markers CD34 and CD133. hASCs differentiated into adipocytes after cultivation in adipogenic medium. Exposure of hASCs for 10 days under hypoxia (3% oxygen) in combination with leptin increased the percentage of CD31 + endothelial cells as well as CD31, VE-Cadherin, Flk-1, Tie2, von Willebrand factor, and endothelial cell nitric oxide synthase mRNA expression. This was enhanced on coincubation of vascular endothelial growth factor (VEGF) and leptin, whereas VEGF alone was not sufficient. Moreover, hASCs cultured on a matrigel surface under hypoxia/VEGF/leptin, showed a stable branching network. Hypoxic conditions significantly decreased apoptosis as evaluated by cleaved caspase-3, and increased prolyl hydroxylase domain 3 mRNA expression. Hypoxia increased expression of VEGF as well as leptin transcripts, which were significantly inhibited on co-incubation with either VEGF or leptin or a combination of both. Furthermore, leptin treatment of hypoxic cells increased the expression of the long/signaling form of the leptin receptor (ObRL), which was augmented on co-incubation with VEGF. The observed endothelial differentiation was dependent on the Akt pathway, as co-administration with Akt inhibitor abolished the observed effects. In conclusion, our data demonstrate that hASCs can be efficiently differentiated to endothelial cells by mimicking the hypoxic and pro-angiogenic microenvironment of adipose tissue.

gous cell therapy for ischemia revascularization and for defined therapy of cardiovascular diseases [9–13]. For different reasons, human adipose tissue-derived stem cells (hASCs) are attractive stem cells for future clinical application. First, adipose tissue is abundant in most individuals and can be easily collected by local anaesthesia with less discomfort and donor site damage; second, adipose tissue has a significantly higher stem cell density [14]. Therefore, a small amount of adipose tissue can yield sufficient stem cells with proliferation and differentiation potential for autologous cell transplantation. Moreover, experimental studies showed that hASCs could differentiate into cell types of the three germ layers in vitro and in vivo [15,16]. These include bone, cartilage, cardiomyocytes, and vascular cells [17,18]. In vivo studies have boosted the interest in ASCs, as they may stimulate peripheral

Introduction

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evascularization of injured and ischemic organs is essential to restore organ function. The delivery of adult stem cells into ischemic tissue is emerging as a novel therapeutic option [1], as autologous adult stem cells obtained from the body of the patient appear to be immunologically compatible [2]. Investigators have identified adult stem cells that may participate in the process of new vessel formation in numerous organs and tissues [3–5]. However, isolation of human adult stem cells from the heart [6], skeletal muscle [7], or bone marrow [8] mainly yields a low number of stem cells and carries the risk of damage to donor organs. Recently, many publications described that adipose tissue contains multi-potential stem cells and holds great promise in autolo1

Clinic of Internal Medicine I, Department of Cardiology, University Heart Center, Jena University Hospital, Jena, Germany. Department of Zoology, Faculty of Science, Tanta University, Tanta, Egypt. 3 Department of Cranio-Maxillofacial Surgery and Plastic Surgery, Jena University Hospital, Jena, Germany. 4 Department of Physiology, Faculty of Medicine, Justus Liebig University, Giessen, Germany. *These two authors contributed equally to this work. 2

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2 nerve repair [19], recover the function of acute spinal cord damage [20], repair the liver injury [21], and reduce hyperglycemia in diabetic mice [22]. In addition, Lu et al. also reported that hASCs have the potential to enhance the blood supply in random-pattern skin flaps [23]. Hence, it is reasonable to expect that hASCs will become one of the preferred choices of human adult stem cells for future clinical applications. Previously, it was reported that hypoxia potentially enhances neo-angiogenesis of ASCs via a paracrine mechanism that involves an increased secretion of certain growth factors [24–26]. One growth factor that plays an essential role in the cellular response to hypoxia is vascular endothelial growth factor (VEGF) [27]. Notably, angiogenic and anti-apoptotic growth factors were secreted at a fivefold higher level by ASCs as compared with differentiated cells, and their secretion was significantly enhanced under hypoxic conditions [25,28]. Adipose tissue further actively participates in endocrine processes by secreting and releasing a vast array of protein signals, including hormones, growth factors, chemokines, and cytokines such as leptin [29]. Leptin is the product of the obese (Ob) gene and was considered one of the most important white adipose-derived adipokines that is not only important in the regulation of food intake and energy balance [30,31], but also acts as a potent angiogenic factor [32– 34]. In addition to adipose tissue, high levels of leptin were found in other angiogenic tissues such as the placenta, skeletal muscles, the mammary gland [35,36], and also cancer cells under hypoxic conditions [37]. Leptin exclusively binds to its receptor, while several isoforms of the leptin receptor (ObR) are found in diverse tissues and different cancer cells [38]. Interestingly, it was found that on leptin activation, the ObR including the long (ObRL) and the short (ObRS) isoform, can utilize a number of diverse signaling pathways relevant to angiogenesis [39]. Leptin receptors are expressed in endothelial cells and Flk-1 + cardiovascular progenitor cells derived from the bone marrow [32]. Consequently, in vitro studies demonstrated that leptin can stimulate growth and survival of endothelial cells, induce their migration, and organize them into capillary-like tubes [33,40,41]. Moreover, in vivo studies showed that leptin is able to stimulate the formation of new blood vessels in the chick chorioallantoic membrane [40]. In this study, we have examined the potential of hASCs to differentiate into endothelial cells in vitro by using 3% oxygen in combination with typical angiogenic factors, thus mimicking the in vivo microenvironment for tissue repair.

BEKHITE ET AL. two patients suffered from a mild primary hypertension (World Health Organisation grade I), and one of them was diabetic. Previously, Zhang et al. [43] showed no significant difference in the cell harvest depending on whether ASCs were obtained from different genders, elderly patients suffering from cardiovascular diseases, smokers, obese patients (BMI > 30), or healthy patients. Fat tissue was mainly taken from the abdomen or the area under the scapula. All patients provided written consent, and the isolation procedure was carried out following a routine operation according to the Declaration of Germany for the use of human tissue and was approved by the University of Jena Health Human Research Ethics Committee.

Isolation and cell culture of hASCs hASCs were isolated from *50 g surgical excised adipose tissue during a routine operation with small modifications following the protocol of Zuk et al. [44]. First, obtained adipose tissue was rinsed and washed with phosphate-buffered saline (PBS) to remove potential blood cells. Then, yellow fat tissue was minced into small pieces (*4 · 4 mm). Further, vascular and other fibroblastic structures were removed, and the obtained suspension was digested with 2 mg/mL collagenase B (PAA, Co¨lbe, Germany) at 37C on a conventional shaker for *60 min. After this, the suspension was diluted with Dulbecco’s-Modified Eagle Medium (DMEM) high glucose (Gibco, Life Technologies, Darmstadt, Germany) supplemented with 2 mM glutamine, 100 U/mL penicillin, and 100 mg/mL streptomycin (Biochrom, Berlin, Germany) and 15% heat-inactivated fetal bovine serum (FBS) (SigmaAldrich, Deisenhofen, Germany). In the next step, the cell suspension was centrifuged for 10 min at 2,000 rpm, and the floating adipocytes in the upper phase were separated from the obtained cell pellet. These cells were further resuspended in DMEM medium as described earlier, filtered through a 100 mm mesh, and, finally, treated with 160 mM NH4Cl for 10 min to lyse red blood cells. The remaining cells were then re-suspended in DMEM medium as described earlier, filtered through a 40-mm cell strainer, and seeded in conventional six-multiwell tissue culture plates. Cell culture was carried out at 37C in a humidified environment containing 5% CO2. Cells with 80% confluence were enzymatically dissociated using 0.25% trypsin dissolved in Hanks’ balanced salt solution (Gibco) supplemented with 1 mM ethylenediaminetetraacetic acid (EDTA). Cells were further passaged and cultured as previously described [45]. The passages of hASCs used in the presented work were 3, 6, 9, and 12.

Materials and Methods Nomenclature of ASCs (hASCs) Depending on the nomenclature of the International Fat Applied Technology Society [42], we decide to use the term hASCs, which identifies a plastic-adherent stroma fraction within the cell population obtained from adipose tissue.

Source of hASCs hASCs were isolated from nine different patients during routinely done plastic surgery. The average age of the donors was 58 – 8.5 years, and body mass index (BMI) was 26 – 4;

Growth factors and inhibitors used during differentiation experiments of hASCs For induction of differentiation, isolated hASCs at passage 6 were seeded at a cell density of *1 · 105 cells per well of a conventional six-well tissue culture plate and cultured for the respective time of the experiment. Cells were fed every day with DMEM medium supplemented as described earlier. Depending on the respective experiment, the medium was modified with human VEGF at a final concentration of 100 ng/mL [46,47] or with human leptin at a final concentration of 1 mg/mL (both from Sigma-Aldrich). In addition,

DIFFERENTIATION POTENTIAL OF HASCS TOWARD ENDOTHELIAL CELLS combinations of both factors with the same concentrations were used as described in the ‘‘Results’’ section. Further, depending on the experimental setting, cell culture was carried out under atmospheric oxygen (*21% oxygen that was defined as normoxia) [48] or under reduced oxygen (3% oxygen that was defined as hypoxia) [28] in a Cytoperm 2 incubator (Thermo Fisher Scientific, Bremen, Germany). For establishing hypoxic culture conditions (3% O2), the incubator was supplied with nitrogen gas in addition to CO2. The Akt inhibitor was applied at a final concentration of 50 mM [49–51] and the general VEGF receptor inhibitor, SU5614, was used at a final concentration of 1 mM [52] (both from Calbiochem, Bad Soden, Germany).

Flow cytometry analysis Cells were harvested from the culture dishes using trypsin EDTA as described earlier and fixed with 4% paraformaldehyde, pH 7.4, dissolved in PBS. Single cells for flow cytometry (FCM) were incubated for 30 min with the respective primary antibodies (Supplementary Table S1; Supplementary Data are available online at www.liebertpub.com/scd) and next washed with PBS supplemented with 1% FBS. After removing the primary antibody by washing twice, samples were re-incubated with either a Cy3- or Cy5-conjugated secondary antibody (Merck-Millipore, Schwalbach, Germany) at a dilution of 1:500 using an exposure time of 20 min. Before starting FCM, cells were again rinsed twice with wash buffer. FCM was carried out using an FACS Calibur (Becton Dickinson, Heidelberg, Germany).

Adipogenic differentiation of hASCs After a primary culture of hASCs in DMEM and expansion to three passages, cells were trypsinized and seeded onto gelatine-coated six-well tissue culture plates at a density of *1 · 105/well. Cells were then incubated in a small amount of DMEM for 1 day to adhere, and the medium was subsequently replaced with an adipogenic medium containing DMEM-F12 medium (Gibco) supplemented with human basic fibroblast growth factor (100 ng/mL) (Sigma-Aldrich), 20% knockout serum replacement (Gibco), 1% non-essential amino acids, 100 U/mL penicillin, and 100 mg/mL streptomycin (Biochrom). Cell culture was carried out in a humidified environment containing 5% CO2 at 37C. The medium was exchanged every day for approximately 20 days. Consecutive adipogenic differentiation was assessed by Oil Red O staining. Before staining, cells were rinsed with PBS and fixed in formaldehyde for 1 h. In the next step, samples were incubated in 2% (wt/vol) Oil Red O reagent (Sigma-Aldrich) for 5 min at room temperature. The remaining reagent was removed by washing with 70% ethanol, followed by several changes of distilled water. The ratio of positively stained cells in comparison to non-stained cells was analyzed using an Axiovert 25 microscope (Zeiss, Jena, Germany).

Immunocytochemistry Adherent or single cells were fixed for 30 min with 4% paraformaldehyde at 4C and permeabilized, if necessary, for 10 min with PBS containing 0.1% Triton X-100 (PBST; Sigma-Aldrich). Blocking against unspecific binding was performed using 0.01% PBST containing 10% milk for 1 h.

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Cells were incubated with the respective primary antibody dissolved in 0.01% PBST containing 10% milk for 120 min at room temperature. After washing and removing the primary antibody, samples were re-incubated with either a Cy3- or Cy5-conjugated secondary antibody (Millipore) at a dilution of 1:100 using an exposure time of 60 min. Immunocytochemistry (ICC) was recorded using a confocal laser scanning microscope (cLSM) LSM 510 (Zeiss).

Western blot analysis Samples were washed once in ice-cold PBS and lysed in ice-cold lysis buffer as previously described [50]. Protein aliquots of 40 mg were separated in 8% or 12% sodium dodecyl sulfate gels and transferred onto nitrocellulose membrane. The membranes were blocked with 5% dried fat-free milk in Tris-buffered saline containing 0.1% Tween 20. Incubation with primary antibodies was performed at 4C overnight. Immunoreactive bands were visualized by using peroxidaseconjugated secondary antibody, ECL detection kit (Amersham, Freiburg, Germany) and captured by a digital imaging system LAS 3000 (Fujifilm, Tokyo, Japan).

Matrigel angiogenesis assay Thawed matrigel (BD Bioscience, Heidelberg, Germany) was added in a 96-multiwell tissue culture plate without dilution and allowed to solidify at room temperature. Respective hASCs were seeded on these matrigel-coated wells and cultured at 37C. After completion of sprouting formation, five representative images for analysis were recorded from each well using the cLSM in combination with digital image capture software.

Reverse transcription-polymerase chain reaction and real-time polymerase chain reaction Total RNA was prepared using Quiashredder columns and an RNeasy mini kit (both Qiagen) according to the manufacturer’s recommendation, followed by a genomic DNA digestion using DNaseI (Invitrogen, Darmstadt, Germany). cDNA synthesis was performed using 0.5 mg RNA and SuperScript RTase II (Invitrogen). Further, a polymerase chain reaction (PCR) amplification was performed using the primer sets as shown in the Supplementary Data (Supplementary Table S2). Reverse transcription–polymerase chain reaction (RT-PCR) was performed under conditions as previously described by Zhang et al. [43], using 39 cycles at 59C instead of 40 cycles at 60C annealing temperature. Subsequently, gel images were captured after electrophoresis, preparing a 1% agarose gel (Fig. 5A, B, E, and F). Realtime RT-PCR (all other PCR data) was performed using SYBR-Green (Qiagen) Mastercycler ep Realplex (Eppendorf, Hamburg, Germany). Relative expression values were obtained by normalizing the CT values of the tested gene to CT values of the internal standard gene using the DDCTmethod. Human Galactosidase beta (GUSB) was used as the internal standard gene, which was also resistant to hypoxic side effects.

Statistical analysis Data were presented as mean values – standard deviation, with n denoting the number of experiments. All

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experiments were repeated at least thrice. GraphPad InStat-3 software (GraphPad Software, Inc., San Diego, CA) was applied for unpaired t-test and one-way analysis of variance data as appropriate. A value of P < 0.05 was considered significant (*).

Results Characterization of the isolated hASCs To guaranty identical characteristics of the isolated hASC population over prolonged cell culture times and passages, cells were characterized in addition to morphological criteria through the expression of nine different stem cellassociated surface antigens at passages 3, 6, 9, and 12 using FCM (Table 1). It was confirmed that isolated hASCs cultured in DMEM medium containing 15% FBS for 12 passages were positive for CD29, CD44, CD73, CD90, CD105, and CD166. In contrast, no significant expression of the haematopoietic lineage markers CD34 and CD133 and the endothelial cell marker CD31 was observed in any of the earlier and later passages. Moreover, these isolated hASCs were devoid of Flk-1 + cells (Supplementary Fig. S1). Therefore, our results underline that the characterized cells, isolated from human adipose tissue samples, represented specific mesenchymal cells rather than hematopoietic cells or other progenitors (Supplementary Fig. S2A–C).

Expression of stem cell markers in hASCs Further characterization studies were performed using real-time RT-PCR analyzing Oct3/4, Sox2 and Nanog transcripts. These markers are widely accepted for defining stem cells [53]. Interestingly, transcripts of Oct3/4, Sox2, and Nanog were detectable at passages 3, 6, 9, and 12 of isolated hASCs lines (Fig. 1A–C, n = 5). In parallel, Oct3/4, Sox2, Nanog, and Lin28 expression was localized in isolated hASCs by ICC. Lin28 localized to the cytoplasm of ASCs, whereas expression of Oct3/4, Sox2, and Nanog was exclusively found in the cell nucleus (Fig. 1D–F, n = 3). The overall mRNA expression of Oct3/4, Sox2, and Nanog remained stable over the passages, when cells were cultured in DMEM supplemented with 15% heat-inactivated FBS as described earlier.

Adipocyte differentiation The percentage of adipocytes was calculated by microscopic inspection after staining the cells with Oil red. The number of cells in a visual field with red lipid droplets was determined as a percentage relative to the total number of cells that were positively stained with hematoxylin. When isolated hASCs were cultured under conditions that induce adipogenesis for around 20 days, more than 78.7 – 5.3% of cells formed clusters of Oil red O-positive lipid-laden cells. This percentage was significantly higher in comparison to the standard cell culture protocol (Fig. 2A–C, n = 3).

Protection of hASCs from apoptosis by hypoxic culture conditions Caspase-3 is one of the critical effector caspases during apoptosis [54]. Moreover, there is evidence that hypoxia under certain conditions is associated with activation of caspase-3 [55]. Analyzing the protein level of cleaved caspase-3, we performed FCM of hASCs under hypoxic conditions in comparison to conventional cell culture conditions. Interestingly, significant inhibition of cleaved caspase-3 levels was observed during exposure of hASCs for 10 consecutive days to hypoxia (3% oxygen, P < 0.02) or co-incubated with leptin (1 mg/mL, P < 0.01), VEGF (100 ng/mL, P < 0.02), and both together (P < 0.03) in the presence of 3% oxygen in comparison to the respective controls (Fig. 3a, b, n = 5). To understand how hASCs sense oxygen and how this sensing is potentially regulated, the mRNA expression of prolyl hydroxylase isoforms [prolyl hydroxylase domain 1 (PHD1) - prolyl hydroxylase domain 3 (PHD3)] was analyzed. Interestingly, PHD3 mRNA transcripts were significantly upregulated when hASCs were cultured under hypoxic conditions as described earlier (Fig. 3c, n = 5). In contrast, expression of PHD1 and PHD2 mRNA transcripts was not altered if hASC were cultured under hypoxia (data not shown). This result suggests that during hypoxia, PHD3 potentially plays a role in regulating an anti-apoptotic pathway in hASCs. Furthermore, PHD3 signaling could be a key pathway that distinguishes stem cells from terminally differentiated cells, as one of the major advantages of stem cells compared with differentiated cells is the ability to become activated if surrounding differentiated cells suffer from oxygen depletion.

Table 1. Flow Cytometric Analysis of CD Marker Expression on Adipose Tissue-Derived Stem Cells Passage 3 CD antigen CD29 CD44 CD73 CD90 CD105 CD166 CD31 CD34 CD133

Passage 6

Passage 9

Passage 12

%

SD –

%

SD –

%

SD –

%

SD –

97.02 99.03 97.68 97.57 97.59 94.83 0.02 0.25 0.01

1.34 0.59 1.82 1.25 2.11 2.87 0.05 0.15 0.01

94.5 98.96 99.08 93.80 97.16 98.64 0.02 0.18 0.06

2.85 0.52 0.55 3.83 1.58 1.28 0.02 0.09 0.07

94.61 98.85 99.48 93.28 96.47 98.87 0.01 0.25 0.10

3.10 1.22 0.14 4.31 2.99 1.43 0.02 0.23 0.07

95.23 99.20 98.79 92.27 99.10 98.53 0.01 0.32 0.01

2.72 0.53 1.49 3.56 1.06 1.72 0.01 0.28 0.02

Data are presented as a percentage (%) – SD obtained from five donors. SD, standard deviation.

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FIG. 1. Expression of stem cell markers in human adipose tissue-derived stem cells (hASCs). The stem cell-associated genes Oct3/4 (A), Sox2 (B), and Nanog (C) were detectable in the isolated hASCs. These genes were used as a further factor to characterize the obtained hASCs from different donors. Interestingly, real-time polymerase chain reaction (PCR) showed that expression of the mentioned genes was present in isolated hASCs and was further stable over several passages during cell culture (A–C). Immunocytochemistry (ICC) clearly localized the respective proteins of Lin28 (D) in combination with Oct4 (D) as well as Sox2 (E) and Nanog (F), respectively. Data are considered non-significant (n.s.) at P > 0.05. Color images available online at www.liebertpub.com/scd

FIG. 2. Adipogenic differentiation of hASCs. To demonstrate that hASCs can undergo adipogenic differentiation, hASCs were cultured for a defined time period in the presence of adipogenic medium as described in the ‘‘Materials and Methods’’ section. A successful adipogenic differentiation was assessed by clusters of lipid-containing adipocytes that were detected by Oil red-O staining (B) and compared with untreated cells (A) for the same time period. Adipogenesis was quantified using light microscopy by counting the percentage of lipid-loaded cells in comparison to native hASCs (C). Data are considered significant at P < 0.05 (*) and n.s. at P > 0.05

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FIG. 3. To exclude a potential apoptotic side effect mediated through activation of cleaved caspase 3 during exposure of hASCs to 3% oxygen in comparison to 21% oxygen, flow cytometry (FCM) analysis was performed (A, B). hASCs showed a significant inhibition in the expression level of cleaved caspase 3 if they were exposed to 3% oxygen (A, B). Furthermore, down-regulation of cleaved caspase-3 was observed if hASCs were additionally treated with vascular endothelial growth factor (VEGF) and leptin in different combinations (A, B). To elucidate how hASCs sense a reduced oxygen tension of 3% mRNA, the expression of prolyl hydroxylase 3 (PHD3) was analyzed. It was found that mRNA expression of PHD3 was significantly increased in all samples that were exposed to 3% oxygen (C). Data are considered significant at P < 0.05 (*) and n.s. at P > 0.05.

hASCs exposed to 3% oxygen and leptin show a dose- and time-dependent increase of CD31 + cells To determine the optimal stimulus for the differentiation of CD31 + cells from isolated Flk-1 - hASCs, the effect of different concentrations of leptin and VEGF, applied for a defined time period, was analyzed. hASCs were grown in the presence of the respective stimulus and were harvested at day 0, 2, 4, 6, 8, 10, 12, and 14 of differentiation. Leptin was applied in a concentration range of 0–2 mg/mL (Fig. 4A–D), whereas VEGF was used at 0–300 ng/mL (Fig. 4E). The maximum culture period was 14 days, and conditions of atmospheric oxygen concentration were compared with cell cultures under 3% oxygen. To quantify the respective effects, RT-PCR (Fig. 4A, B, n = 3) and FCM (Fig. 4C, D, n = 3) were performed. First of all, it was obvious that neither leptin (Fig. 4A, B, n = 3) nor VEGF (Fig. 4E, n = 3) under normoxia, even if applied with the maximum dose, increased the CD31 mRNA expression if incubated for 10 days. Likewise, no significant increase in the number of CD31 + cells was found under normoxic conditions using FCM (Fig. 4C, D). In contrast, exposure of hASCs to 3% oxygen in combination with leptin dose-dependent significantly increased mRNA expression of CD31 after 10 days of incubation (Fig. 4A, B). CD31 mRNA

expression was only marginally detectable on day 2 and 4 of differentiation (Fig. 4F, G, n = 3). Starting from day 6 of differentiation, hASCs showed—if exposed to 1 mg/mL leptin— a significant increase of CD31 mRNA expression from day 6 to 10. A longer exposure time under the described conditions did not lead to a further increase of mRNA transcripts in comparison to the respective controls. In conclusion, the highest increase of CD31 mRNA expression was observed if isolated hASCs were exposed to 1 mg/mL leptin in combination with 3% oxygen for 10 consecutive days (Fig. 4D).

Stimulation of endothelial cell differentiation from hASCs by hypoxia, leptin, and VEGF FCM analysis for CD31 + cells after treatment with different concentrations of VEGF revealed that 10 ng/mL [56] or 50 ng/mL [43] VEGF increased CD31 + cell numbers less efficiently as compared with 100 ng/mL (Supplementary Fig. S3A, B). Concentrations of 100 ng/mL VEGF have been previously shown to increase vascular permeability, and to stimulate cell migration, proliferation, and differentiation of endothelial cells [46,47,57–59]. To further increase endothelial cell differentiation of hASCs under controlled conditions, hASCs were cultured with either 100 ng/mL VEGF or 1 mg/

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FIG. 4. Endothelial cell differentiation of hASCs. To identify the optimal dose and exposure time of leptin in combination with 3% oxygen to induce endothelial differentiation of isolated hASCs, CD31 mRNA expression was monitored by reverse transcription–polymerase chain reaction (RT-PCR) (A, B) and FCM (C, D). First, mRNA expression of CD31 clearly showed a significant increase if hASCs were exposed to at least 1 mg/mL leptin in combination with 3% oxygen for 10 days (A, B). (A) Shows a representative RT-PCR blot of CD31 mRNA expression under conditions of normoxia (21% oxygen) and hypoxia (3% oxygen). (B) Bar chart of the means – standard deviation of three experiments showing CD31 mRNA expression after incubation with different concentrations of leptin for 10 days. Human umbilical vein endothelial cells (HUVECs) were used as positive control. (C, D) FCM analysis of CD31 + cells on treatment of hASCs with different doses of leptin. Note that doses of leptin exceeding 1 mg/mL did not further increase the percentage of CD31 + cells. (E) Cell culture conditions based on 21% oxygen or 3% oxygen and only using VEGF at 0–300 ng/mL did not increase the mRNA expression of CD31. (F, G) CD31 mRNA expression showed a maximum at day 10 if hASCs were treated with leptin (1 mg/mL) and 3% oxygen in comparison to the respective control. Data are considered significant at P < 0.05 (*) and n.s. at P > 0.05. mL leptin, or a combination of 1 mg/mL leptin, 100 ng/mL VEGF, and 3% oxygen for 10 consecutive days. To evaluate the percentage of CD31 + cells after this experimental procedure, FCM was performed (Fig. 5B, C, n = 5). It was found that leptin alone increased the number of CD31 + cells to

5.5%. However, the combination of leptin, VEGF, and hypoxia was most effective and approached *30% CD31 + cells, indicating pronounced endothelial cell differentiation. These results were confirmed using ICC (Fig. 5A, n = 5). To further characterize the differentiated endothelial cells, ICC against

8 VEGFR2 (Flk-1) (Fig. 5D, E, n = 3) and von Willebrand factor (Fig. 5F, G, n = 3) was performed and compared with undifferentiated hASCs. Interestingly, we also found that these endothelial markers were significantly up-regulated if hASCs were exposed to leptin, VEGF, and hypoxia for 10 consecutive days. To further confirm these findings, realtime RT-PCR for CD31 (Fig. 5H, n = 5), Flk-1 (Fig. 5I, n = 5),

FIG. 5. Expression of endothelial cell markers in hASCs under conditions of hypoxia, leptin, and VEGF treatment. hASCs exposed to 3% oxygen in combination with 1 mg/mL leptin with or without 100 ng/ mL VEGF for 10 consecutive days differentiated to a significant percentage of CD31 + cells that was analyzed by ICC (A) and FCM (B, C). Cell culture conditions based on 21% oxygen or treatment with VEGF alone did not increase the percentage of CD31 + cells (A–C). To verify an endothelial cell phenotype, a further ICC was performed (D–G). Interestingly, Flk-1 + (D, E) and vWf + (F, G) could be identified if hASCs were exposed to to 3% oxygen in combination with 1 mg/mL leptin with or without 100 ng/mL VEGF for 10 consecutive days. Furthermore, real-time RT-PCR clearly showed an increase of typical endothelial cell transcriptions factors, including CD31 (H), Flk-1 (I), Tie-2 ( J), endothelial cell nitric oxide synthase (eNOS) (K), and VE-Cadherin (L). In addition, protein expressions of the endothelial markers were evaluated using western blot analysis in hASCs. HUVECs under normoxia were used as an endothelial control. Significant increases in CD31, VE-Cadherin, and Flk-1 (M, N) were detected under hypoxia. In contrast, eNOS protein could not be detected in endothelial cells under normoxia or hypoxia (M, N). Data are considered significant at P < 0.05 (*) and n.s. at P > 0.05. Color images available online at www.liebertpub.com/scd

BEKHITE ET AL. Tie2 (Fig. 5J, n = 5), endothelial cell nitric oxide synthase (eNOS) (Fig. 5K, n = 5), and VE-Cadherin (Fig. 5L, n = 5) was performed. Transcripts of these endothelial markers were significantly up-regulated after exposure of hASCs for 10 days to leptin alone and even more pronounced after exposure toward the combination of leptin, VEGF, and hypoxia. In contrast, treatment with VEGF alone did not show a

FIG. 5.

(Continued). 9

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FIG. 5.

(Continued).

significant increase of endothelial cell differentiation (Fig. 5H–L, n = 5). In addition, protein expression of endothelial markers on incubation of hASCs under hypoxia, leptin, and VEGF was evaluated using western blot analysis. Human umbilical vein endothelial cells (HUVECs) were used as an endothelial control. We found a significant increase in CD31, VE-Cadherin, and Flk-1 that was comparable to the mRNA expression level (Fig. 5M, N, n = 3). In contrast, the mRNA expression level of eNOS could not be detected as protein level under the same condition of treatment (Fig. 5M, N, n = 3).

hASCs exposed to hypoxia, VEGF, and leptin show a precisely regulated mRNA expression pattern of the respective receptors and agonists To further understand the signaling mediated by leptin in isolated hASCs, mRNA expression of known leptin receptor isoforms that are distinguished in an ObRL and ObRS isoform were analyzed. Both isoforms were identifiable in all isolated hASCs. Interestingly, the relative mRNA expression of the ObRL isoform was significantly up-regulated after an exposure of hASCs to leptin (1 mg/mL) alone or the combi-

nation of leptin, VEGF (100 ng/mL), and hypoxia for 10 consecutive days (Fig. 6B, n = 3). Furthermore, it was found that the relative leptin mRNA expression was significantly up-regulated if hASCs were cultivated only under hypoxic conditions for 10 consecutive days (Fig. 6A, n = 3). In contrast, the addition of leptin or VEGF alone reduced the relative leptin mRNA expression if exposed to hypoxia for the same time period (Fig. 6A, n = 3). Since VEGF is the most relevant growth factor that stimulates vascular differentiation and growth, the relative mRNA expression of VEGF was analyzed under conditions of VEGF and leptin treatment of hASCs. As expected, hypoxia treatment significantly increased VEGF mRNA expression. Interestingly, upregulation of VEGF transcripts under hypoxia were abolished under conditions in which either leptin or leptin + VEGF were present in the cell culture medium (Fig. 6C, n = 3).

The effect of endothelial cell differentiation from hASCs on hypoxia and leptin is mediated via an Akt-dependent pathway To evaluate a potential signaling pathway mediating the effect of hypoxia, VEGF, and leptin during the differentiation

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FIG. 6. Expression of leptin and long/signaling form of the leptin receptor (ObRL) in hASCs. The expression of leptin (A) as well as ObRL (B) was analyzed by RT-PCR. It was found that exposure of hASCs to 3% oxygen for 10 consecutive days increased leptin mRNA expression, which was again reduced if leptin or VEGF were applied (A). The obtained data clearly showed that the ObRL isoform transcripts were generally expressed in hASCs under control conditions (B). Higher transcript expression was noticed if hASCs were exposed to 3% oxygen in the absence of leptin for 10 consecutive days (B). (C) VEGF mRNA expression was up-regulated if hASCs were cultured under 3% oxygen in the absence of leptin for 10 consecutive days. This was abolished when either leptin or leptin + VEGF was present in the incubation medium. Data are considered significant at P < 0.05 (*) and n.s. at P > 0.05.

of hASCs toward endothelial cells, we co-incubated hASCs with an Akt pathway (50 mM) inhibitor and the VEGF receptor inhibitor SU5614 (1 mM). The incubation with the respective inhibitor was performed singular or in combination with hypoxia, leptin (1 mg/mL), and VEGF (100 ng/mL) for 10 consecutive days, and the relative CD31 + positive immunofluorecence area in cell monolayers was analyzed. Interestingly, the results showed that co-incubation with Akt inhibitor completely abrogated hASC differentiation toward endothelial cells, which was observed under the stimulus of hypoxia and leptin and the combination of leptin, VEGF, and hypoxia (Fig. 7A, B, n = 3). It was obvious that inhibition of the VEGF receptor did not significantly affect the differentiation of CD31 + cells from hASCs on treatment with leptin in combination with hypoxia, but significantly abolished the increase of endothelial cell differentiation when cells were treated with the combination of leptin, VEGF, and hypoxia (Fig. 7A, B, n = 3).

Endothelial cells differentiated from hASCs on hypoxia, leptin, and VEGF form typical endothelial sprouts in vitro using matrigel-dependent angiogenesis assays To further characterize the angiogenic potential of hASCs, endothelial cell sprouts were assessed by light microscopy and quantified on a respective matrigel surface (Fig. 8A, B, n = 3). This in vitro angiogenesis assay showed significant stimulatory effects on sprout- and branch formation of hASCs if co-incubated with leptin (1 mg/mL) and VEGF (100 ng/mL) in the presence of 3% oxygen in comparison to the respective controls or on only leptin treatment (Fig. 8A, B, n = 3). Furthermore, it was obvious that inhibiting the Aktpathway totally abolished the effect of leptin and VEGF on hASC cell sprouting. In consistence with the results presented earlier, inhibition of the VEGF receptor by SU5614 alone resulted in a significant reduction in sprouting and

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FIG. 7. Involvement of the Akt pathway in endothelial differentiation of hASCs. To verify that the ObRL-Akt pathway was involved in endothelial differentiation if hASCs were exposed to 3% oxygen and leptin, CD31 expression was analyzed in the presence or absence of an Akt inhibitor (50 mM) by ICC. The obtained data illustrate that co-incubation of hASCs with 3% oxygen and leptin with or without VEGF in the presence of an Akt inhibitor abolished the previously observed endothelial cell differentiation (A, B). On coincubation of leptin with the VEGFR-2 inhibitor SU5614, endothelial cell differentiation was reduced to the level achieved with leptin alone (A, B). Data are considered significant at P < 0.05 (*) and n.s. at P > 0.05. Color images available online at www.lie bertpub.com/scd

branching if hASCs were co-incubated with leptin and VEGF. In conclusion, the presented data suggest that the Akt pathway is essential for endothelial cell differentiation and cell sprouting of hASCs if exposed to hypoxia and leptin for 10 consecutive days.

Discussion Therapeutic enhancement of neovascularization is one of the important strategies needed to limit the complications of post-ischemic injury [60]. Recently, several in vitro and in vivo studies focused on the development of new therapeutic approaches to increase neo-vessel growth in an ischemic microenvironment by using adult stem cells [61,62]. The purpose of the present work was to study whether isolated hASCs that were initially negative for VEGFR2 (Flk-1) were able to differentiate into endothelial cells in vitro by simulating the microenvironment of adipose tissue in vivo. For this reason, hASCs were exposed to 3% oxygen in comparison to 21% oxygen (control) in the presence of the known angiogenic factors leptin and VEGF. It is well established that cells sense oxygen and that on ischemia and hypoxia, multiple organs can undergo neo-angiogenesis by increasing the capillary density or even by attracting new collateral vessels [63–66]. Experimental evidence from both in vitro and in vivo studies demonstrate the multipotency of ASCs that are isolated from humans and other species. Moreover, it has been

shown that the number of stem cells in adipose tissue is 100– 500 times larger than the number of mesenchymal stem cells (MSCs) derived from bone marrow [14,67]. In general, adipose tissue derives from the mesodermal layer of the embryo and continues to develop after birth [68,69]. Interestingly, these stem cells have the ability to differentiate into several mesodermal [70,71] as well as non-mesodermal lineages [29,72–76]. hASCs can be easily obtained either by liposuction aspirates or by surgically excised fat from the visceral and subcutaneous adipose tissue of patients using local anesthesia [77,78]. Moreover, Harris et al. [79] showed that hASCs obtained from patients with cardiovascular disease can be differentiated toward the endothelial cell lineage. Thus, adipose tissue is considered a promising source of stem cells that could be potentially used for therapeutic strategies of several vascular-related diseases. In the present study, the cell surface marker characteristics of isolated hASCs were analyzed and assessed over several passages to guarantee similar and standardized conditions, consistent with previously described data [80,81]. Isolated hASCs from excised fat displayed a typical surface marker profile for several passages. In detail, they were positive for CD90, CD105, and CD166, which are commonly used markers to identify cells with multi-lineage differentiation potential [70,82]. We also found that the isolated hASCs were positive for CD29 (b1 integrin), which plays a critical role in therapeutic angiogenesis by regulating cell survival, differentiation and incorporation through growth factor receptor

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FIG. 8. Sprout formation of hASCs in the matrigel angiogenesis assay. To further underline the ability of hASC-derived endothelial cells to form primitive vascular sprouts, an in vitro angiogenesis assay on a matrigel-coated cell culture surface was performed (A, B). It was obvious that hASCs if exposed to 3% oxygen with or without VEGF formed vascular sprouts if grown on a matrigelcoated cell surface (A, B). This sprout formation was significantly reduced in the presence of Akt inhibitor, but only partially reduced in the presence of the Flk-1 inhibitor SU5614 (A, B). Data are considered significant at P < 0.05 (*) and n.s. at P > 0.05.

cross-talk after implantation into ischemic tissue [83,84]. Furthermore, ASCs were positive for CD44, the primary receptor responsible for hyaluronan binding [85], which is crucial for the development of neo-extracellular matrix [85]. In contrast, isolated hASCs lacked the expression of known haematopoietic and endothelial surface antigens such as CD34, CD133, and CD31. Interestingly, hASCs isolated from liposuction aspirates but not from excised fat showed around 4% CD34 + and around 1% Flk-1 + cells (data not shown). The difference in the cell surface marker expression could potentially result from the different isolation technique. According to our experiences, surgically excised fat tissue provides a high number of vital cells with a minimal amount of tissue needed for sufficient isolation of hASCs. Furthermore, by using surgically excised fat tissue, it was easier to reduce the possibility of contamination with CD31+ and CD34 + cells by removing vascular structures before enzymatic dissociation. Therefore, hASCs used in this study were only obtained from surgically excised fat tissue.

In the present study, we demonstrated that isolated hASCs can be cultured in DMEM medium for multiple passages without significantly losing their stemness gene mRNA expression. To integrate a further quality parameter for the homogeneity of the isolated hASCs, the general adipogenic potential was investigated using special adipogenic cell culture conditions. One major criterium of adipose cell differentiation are lipid-filled intracellular vacuoles, which are known as well-defined markers of adipogenesis [44,71]. Having characterized the stem cell characteristics of isolated hASCs and analyzed their adipogenic differentiation potential, we focused on the influence of leptin and VEGF in the presence or absence of hypoxic conditions on the endothelial differentiation potential. It is well known that leptin is secreted by white adipocytes into the bloodstream [86] and is associated with the regulation of food intake and energy balance [30]. In addition, leptin has been shown to stimulate proliferation and angiogenesis in cells that express leptin receptors by activating a tyrosine kinase-dependent

14 intracellular pathway [32,39,87]. Leptin was further found to be associated with the pathogenesis of cardiovascular diseases; for example, by stimulating vascular inflammation and oxidative stress [88,89]. It was found that overweight (obese) individuals were associated with an increase in Creactive protein, which was associated with a greater risk of cardiovascular disease [90,91]. In addition, Singhal et al. [92] found that leptin is another factor that may underlie the association of obesity with cardiovascular disease. Accordingly, it is possible that chronically high levels of leptin in obesity could contribute to its adverse effects on cardiovascular health. Weighting these findings in the context of our data an in vitro pre-differentiation of isolated hASC to endothelial cells using hypoxia and leptin should be the favored method in the future. In the present study, we showed that adding VEGF to the cell culture medium of Flk-1 - hASCs did not per se induce endothelial differentiation. In contrast, addition of VEGF (100 ng/mL) to hASCs cells expressing Flk-1 after treatment with leptin under hypoxia displayed significantly increased numbers of endothelial cells, which was abolished in the presence of Flk-1 inhibitor. These results clearly confirm that Flk-1 expression is necessary for endothelial cell differentiation of hASCs cells on VEGF treatment. Moreover, experiments from others previously showed that endothelial growth medium-2 (EGM-2) supplemented with VEGF (10 ng/mL) did not significantly increase endothelial cell marker expression in ASCs that were Flk-1 + [56]. Interstingly, Zhang et al. [43] showed that adding additional VEGF (50 ng/mL) to EGM-2 originally containing VEGF significantly increased endothelial cell markers in Flk-1 + ASCs. Therefore, these results confirm our observation that Flk-1 expression in hASCs and high (100 ng/mL) concentrations of VEGF are crucial for endothelial differentiation. Moreover, our results showed that Flk-1 - hASCs only differentiated to endothelial cells under conditions of reduced oxygen if hypoxia was combined with leptin and VEGF. In this context, previous studies showed that hASCs differentiated to endothelial cells when they were transplanted into the ischemic hind limb or were introduced into other ischemic settings in vivo [74,93]. Based on the present and previous data, we suggest that defined hypoxic conditions, in general, play a role for endothelial differentiation in vivo and in vitro [94,95]. This is presumably due to the up-regulation of pro-angiogenic factors, for example, VEGF, and downregulation of several endogenous angiogenic inhibitors [96,97]. Thus, it is reasonable to speculate that growth and differentiation of hASCs toward endothelial cells is definitely associated with local hypoxia. One of the major known oxygen sensor proteins is HIF-1a, which is significantly expressed in our isolated hASCs and is well known to regulate VEGF expression under hypoxic conditions [98]. Our results emphasize that hASCs sense the surrounding oxygen level that was additionally documented by the significant up-regulation of PHD3 mRNA expression. PHD3 was previously described to be an important modulator of the HIF-1a response toward low oxygen levels, and it has been previously shown to be up-regulated on the mRNA and protein level by experimental hypoxia [99]. Presumably, the increased PHD3 expression during long-term exposure to hypoxia functions as an adaptive mechanism for protecting differentiated hASCs from apoptosis and cell death [100]. To

BEKHITE ET AL. validate this hypothesis, cleaved caspase 3 activation of hASCs, exposed to 3% oxygen for 10 consecutive days, was assessed. As expected, our results demonstrated that hypoxia significantly decreased cleaved caspase 3 levels in comparison to cells cultured with 21% oxygen, suggesting a protective effect of low oxygen concentrations. Oxygen concentrations in different human organs, for example, lung, liver, kidney, and heart, range from 4% to 14% [101–104]; whereas within bone marrow, very low oxygen tensions from 0% to 4% prevail [105]. The general oxygen concentration in adipose tissue was previously determined to be 3% [63–66], suggesting that the experimental conditions of the present study represented physiological hypoxia for hASCs. It was previously described that a defined pre-conditioning of hASCs and other MSCs in a hypoxic environment leads to an up-regulation of HIF dependent genes [106,107]. In addition, our findings demonstrated that exposure of hASCs to leptin in combination with 3% oxygen for 10 days significantly increased the expression of Flk-1 and CD31. This effect was enhanced in the presence of VEGF. In contrast, the combination of VEGF and hypoxia alone was without effects. The additional effect of VEGF may be explained by the up-regulation of Flk-1 on treatment with leptin under hypoxic conditions, which facilitates the cellular VEGF response toward endothelial cell differentiation [108]. To refine endothelial cell differentiation characteristics after exposure of hASCs to 3% oxygen, leptin, and VEGF, the expression of further endothelial marker genes was analyzed. Our data suggested that the differentiated cells carried an endothelial mRNA expression pattern for CD31, Flk-1, Tie2, eNOS, and VE-Cadherin. To further confirm these findings, defined endothelial markers that were known from the profile of HUVECs were analyzed using western blot. We found that CD31, VE-Cadherin, and Flk-1 protein levels increased comparably to the mRNA expression level. However, we could not detect eNOS protein, while its mRNA expression level increased significantly after inducing differentiation toward the endothelial lineage from hASCs. Several experimental studies indicated that nitric oxide (NO) produced in endothelial cells is synthesized by eNOS, and this fact plays a crucial role in cardiovascular function [109– 112]. In accordance to our findings, Cao et al. [74] demonstrated that endothelial cells differentiated from hASCs express eNOS at the gene but not the protein level and did not display significant NO production. Moreover, Fischer et al. [113] showed that endothelial cells differentiated from hASCs did not express significant amounts of eNOS in response to endothelial cell growth supplement and/or exposed to physiological shear force. In consequence to our findings, we suggest that alternative sources of NO production may prevail in hASCs. In this regard, it has been recently shown that hASCs express NOS 1, which is involved in the regulation of intercellular coupling [45]. Further studies have demonstrated that by injecting S-nitroso-Nacetyl-d,l-penicillamine-treated or eNOS-overexpressing ASCs into the heart after infarction, local NO availability could be restored in the injured area, thus leading to induced angiogenesis and a reduced infarction size [114,115]. Moreover, McIlhenny et al. [116] showed that hASC transfected with eNOS gene produced NO at levels similar to endothelial cell controls in vitro and were capable of causing vasorelaxation of aortic specimens ex vivo. This combination of

DIFFERENTIATION POTENTIAL OF HASCS TOWARD ENDOTHELIAL CELLS genetically embedded eNOS in hASCs resulted in efficient NO production and increased the efficiency of endothelial cell differentiation. Our data demonstrated that mRNA expression of the ObRL isoform of the leptin receptor was up-regulated under hypoxic conditions in the presence of either leptin alone or the combination of leptin and VEGF. Interestingly, leptin expression was up-regulated under hypoxia, but was reduced under conditions of either leptin or VEGF alone and completely abolished on co-administration of VEGF and leptin. These findings indicate that leptin and VEGF are involved in negative feedback mechanisms that control leptin expression. To further understand the underlying mechanisms, we analyzed the relevance of the Akt pathway that is known to be activated downstream of the ObRL isoform of the leptin receptor [117,118]. This was done using the sprouting and a network structure formation assay on matrigel in the presence or absence of the respective inhibitor. The effects of leptin or leptin co-incubated with VEGF in combination with 3% oxygen could clearly be abolished in the presence of Akt inhibitor. This was monitored by analyzing the percentage of CD31 + cells in monolayer cell culture as well as the branching network formation on matrigel. In contrast, co-incubation with the VEFGR2 inhibitor SU5614 did not completely blunt the endothelial differentiation and branching network, but significantly abolished the increase of endothelial differentiation observed in the combination of hypoxia, VEGF, and leptin. These data suggest that an ObRL-Akt pathway was involved in endothelial differentiation and migration if hASCs were treated with leptin in combination with 3% oxygen. Thus, our experiments support the recent data of others, suggesting that a functional endothelial p38/Akt/COX-2 signaling pathway is obligatory for the pro-angiogenic response of leptin in human endothelial cells [119]. Taken together, our data demonstrate that the in vivo microenvironment of low hypoxia in combination with leptin and VEGF regulates the differentiation potential of hASCs toward endothelial cells in vitro by utilizing an ObRL-Akt signaling pathway. In future, these findings could especially lighten and accelerate the healing process during critical limb ischemia using a defined in vitro hypoxic preconditioning protocol in combination with an additional leptin treatment of hASCs.

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Sources of Funding

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This study was supported by the Interdisciplinary Center for Clinical Research (IZKF) of the Medical Faculty University Jena, the German Foundation for Heart Research, and the Excellence Cluster Cardiopulmonary System (ECCPS) of the German Research Foundation (DFG).

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Author Disclosure Statement

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The authors declare that they have no competing financial interests.

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19 Address correspondence to: Dr. Mohamed M. Bekhite Clinic of Internal Medicine I Department of Cardiology University Heart Center Jena University Hospital Erlanger Allee 101 Jena 07743 Germany

E-mail: [email protected] Received for publication June 13, 2013 Accepted after revision October 16, 2013 Prepublished on Liebert Instant Online XXXX XX, XXXX