H9C2 cell line originates from the European Collection of Authenticated Cell Cultures and it was purchased from Sigma-Aldrich (St. Louis, MO, United States). H9C2 cells were plated on 96-well plates (2*10⁴ cells/well) and incubated for 24 h in Dulbecco’s Modified Eagle’s Medium (Corning, NY, United States) supplemented with 10% fetal bovine serum (FBS) (Euroclone SpA. Milan/Stockholm).

The primary culture of adult rat cardiomyocytes was prepared as described previously (Makkos et al., 2019). Briefly, male Wistar rats (150–200 g) were anesthetized with pentobarbital (60 mg/kg) and heparinized (500 IU/kg). Hearts were excised and perfused with Krebs-Henseleit solution, followed by digestion with collagenase II (8000 U/mL) for 30–45 min. Ventricles were cut into small pieces and digested for 10 min. Cell suspension was filtered and pelleted under gravity. Ca²⁺ concentration was increased gradually up to a final of 1 mM. Cells (7,500 cells/well) were plated on laminin-coated glass coverslips in a 24-well plate (Thermo Fisher Scientific, Waltham, MA, United States) and maintained in proliferation media for 3 h (5% FBS containing M199) and in growth media (serum-free M199) for 24 h.

Study protocols are demonstrated in Figures 1A, 2A. The concentrations of SST were selected for the in vitro experiments on the basis of competition binding and G-protein activation results on CHO cells (Markovics et al., 2012). Cells were treated with either SST (1, 10, 100, and 300 nM and 1 µM) or its vehicle in growth media in a CO₂ incubator for 1 h. Then H9C2 cells and adult rat cardiomyocytes were covered either with normoxic solution (in mM: NaCl 125, KCl 5.4, NaH₂PO₄ 1.2, MgCl₂ 0.5, HEPES 20, CaCl₂ 1, glucose 15, taurine 5, creatine-monohydrate 2.5, bovine serum albumin (BSA) 0.1%, and pH 7.4) in CO₂ incubator (normoxia groups) or with hypoxic solution (in mM NaCl 119, KCl 5.4, NaH₂PO₄ × 1H₂O 1.2, MgCl₂ × 6H₂O 0.5, HEPES 5, MgSO₄ × 7H₂O 1.3, CaCl₂ × 2H₂O 0.9, Na-lactate 20, BSA 0.1%, and pH 6.4) in a three-gas (mixture of 95% N₂ and 5% CO₂) incubator (simulated ischemia/reperfusion groups), both solutions containing SST or its vehicle for 16 and 3 h, respectively. Then, during the simulated reperfusion phase, cells were kept in growth media containing vehicle or SST in a CO₂ incubator for 2 h. Cell viability was determined using calcein staining (Makkos et al., 2019). After rinsing with Dulbecco’s phosphate-buffered saline (D-PBS), cells were incubated with calcein solution (1 µM) for 30 min at room temperature. Fluorescence intensity of each well was detected by Varioskan Lux multimode microplate reader (Thermo Fisher Scientific, Waltham, MA, United States) at room temperature; excitation wavelength: 490 nm; emission wavelength: 520 nm. Four independent experiments were performed (n = 4) averaging at 70% confluence of cell cultures. Regarding H9C2 cells, a 96-well format was applied and an average of six-well relative fluorescence units (RFU) was used in one treatment group. Regarding adult rat cardiomyocytes, a 24-well format was applied and an average of four wells RFU was used in one treatment group. Results are shown as RFU (Brenner et al., 2020).

Porcine left ventricle and plasma samples of the present study have been obtained from one of our previous studies in a closed-chest pig model of acute myocardial infarction and cardioprotection by ischemic pre-, post-, and remote conditioning (Baranyai et al., 2017). The study was conducted according to the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH publication No. 85–23, revised 1996) and to the EU Directive (2010/63/EU) and was reported according to the ARRIVE guidelines (Kilkenny et al., 2010). The study was approved by the Animal Ethics Committee of the Hungarian National Food Chain Safety Office (SOI/31/26–11/2014). Briefly, pigs were randomly divided into five groups: ischemia/reperfusion, ischemic preconditioning, ischemic postconditioning, remote ischemic conditioning, and sham-operated groups. In the ischemia/reperfusion group, 90 min of myocardial ischemia was applied, which was induced by left anterior descending coronary artery (LAD) occlusion. The ischemic preconditioning group was subjected to 3 × 5 min myocardial ischemia prior to 90 min LAD occlusion. The ischemic postconditioning group was subjected to 6 × 30 s of myocardial ischemia after 90 min LAD occlusion at the start of the reperfusion. The remote ischemic conditioning group was subjected to 4 × 5 min of hind limb ischemia during the 90 min LAD occlusion. In the sham group, the balloon catheter was inserted in the LAD coronary artery, but it was not inflated (Figure 3A). After 3 h reperfusion, plasma samples and myocardial tissue samples were collected from the ischemic region of the left ventricle myocardium. Group sizes are as follows: sham (n = 6–7), ischemia/reperfusion (n = 5–7), ischemic preconditioning (n = 4–5), ischemic postconditioning (n = 4–5), and remote ischemic conditioning (n = 4–5). Samples were frozen immediately and stored at −80°C. Infarct size measurements confirmed that ischemic preconditioning decreases myocardial necrosis significantly compared to ischemia/reperfusion after 3 h of reperfusion. There was no difference in the area at risk between groups. Ischemic postconditioning and remote ischemic conditioning decrease edema significantly compared to ischemia/reperfusion in the myocardium after 3 days of reperfusion, and ischemic preconditioning shows only a tendency of decrease. For details of the phenotype of the pig model, see (Baranyai et al., 2017).

All experiments were designed and implemented according to the ethical standards of the Declaration of Helsinki (1975). Patients gave their written informed consent to be involved in the study. The protocol was approved in 2018 by the Polish Local Ethics Committee of the National Institute of Cardiology in Warsaw with the identification code IK-NPIA-0021-14/1426/18 project 4.

Human left ventricle and interventricular septum samples were collected in the Department of Heart Failure and Transplantology, Cardinal Stefan Wyszyński National Institute of Cardiology, Warszawa, Poland, as previously described (Varga et al., 2017). Human hearts not used for transplantation for various reasons were obtained from organ donors (control group, CON, n _female = 6 and n_male = 4). The donors with any relevant previous cardiovascular history or any abnormalities in ECG and echocardiography were not included in the present study. Failing hearts were obtained from patients suffering from advanced heart failure of ischemic cardiomyopathy (ICM, n _female = 1 and n _male = 9). Detailed clinical parameters of the patients are summarized in Figure 5.

Left ventricle and interventricular septum samples were collected during heart explantation, avoiding the inclusion of scarred/fibrotic or adipose tissues, endocardium, epicardium, or coronary vessels. The samples were rinsed immediately in saline, blotted dry, frozen in liquid nitrogen, and kept at −80°C until processing for further molecular assays. Another series of left ventricle samples were fixed in neutral buffered formalin and embedded in paraffin for histological assays.

Left ventricle tissue samples were homogenized in 8x volumes (in µL) of distilled water containing 10 µL protease inhibitor (Gordox, 10,000 KIE/mL, Gedeon Richter Plc) using a tissue homogenizer (IKA T25 Digital ULTRA TURRAX). Distilled water shows a similar protein extraction efficiency with pig heart tissue as described in the literature with a different buffer composition (Parés et al., 2020). Afterward, samples were centrifuged at 10,000 rpm at 4°C for 15 min. Protein was extracted from each 1 ml of the plasma with the mixture of 3 ml absolute ethanol and 10 µL 96% acetic acid. Tubes were incubated at room temperature for 30 min, followed by centrifugation at 3,000 rpm for 20 min at 4°C. The supernatant was collected into a reaction tube and dried under a nitrogen flow for 6 h at room temperature. Then, it was resuspended in 300 μl assay buffer and centrifuged at 10,000 rpm at 4°C for 15 min before radioimmunoassay (RIA) determination.

SST-like immunoreactivity was determined by a specific and sensitive RIA method developed in our laboratory as described earlier in detail (Nemeth et al., 1996). Briefly, each reaction tube contained 25 µL tissue homogenates or 750 µl suspended dry plasma sample, 100 µl ¹²⁵I-labeled somatostatin-14 as tracer (3,000 cpm/tube), and 100 µl antiserum (1:445,000). Tubes were filled to 1,000 µl with RIA assay buffer. Antiserum has been raised in sheep against somatostatin-14-bovine thyroglobulin. Following 72 h incubation at 4°C, antigen-bound and free peptides were separated. Tubes were centrifuged at 4°C at 4,000 rpm for 20 min. Radioactivity was measured by a gamma counter (Gamma NZ-310, Hungary). The results were expressed as fmol SST-like immunoreactivity per mg total protein weight in the tissue and per mL in the plasma samples.

Total RNA from pig left ventricle samples (ischemic zone) was extracted using Direct-Zol RNA Mini Prep (Zymo Research, Irvine, CA, United States) according to the manufacturer’s instructions. RNA was then treated with DNase I (Zymo Research, Irvine, CA, United States), and RNA concentration was determined by spectrophotometer (NanoDrop ND-1000, NanoDrop Technologies Inc., Wilmington, DE, United States). 1 µg of total RNA/sample was reverse transcribed into complementary DNA (cDNA) using Maxima™ First Strand cDNA Synthesis Kit for real-time quantitative PCR (RT-qPCR) (Thermo Scientific, Waltham, MA, United States).

The expression of somatostatin and its receptors was assessed using Biometria TProfessional Basic Gradient PCR equipment (Biometra GmbH, Göttingen, Germany). PCR products were identified by their size using 2% agarose gel electrophoresis.

Relative gene expression ratios were determined with Stratagene Mx3000P QPCR System (Agilent Technologies, Santa Clara, CA, United States) using β-ACTIN as a reference gene based on our pilot experiments and literature data (Nygard et al., 2007). Target genes were amplified using 1x Luminaris HiGreen Low ROX qPCR Master Mix (Thermo Scientific, Waltham, MA, United States). Amplifications were carried out under the following conditions: 95°C 10 min, followed by 40 cycles of 95°C 30 s, 60°C 30 s, and 72°C 45 s. Measurements included dissociation curve analysis to ensure amplification specificity. The corresponding group sizes are ischemia/reperfusion group (SSTR1: n = 5, SSTR2: n = 6), ischemic preconditioning group (n = 4), ischemic postconditioning group (n = 5), remote ischemic conditioning group (SSTR1: n = 5, SSTR2: n = 4), and sham group (n = 7). Primers and product lengths for each gene are listed in Supplementary Table S1.

Samples were collected and lysed in 1 ml of QIAzol Lysis Reagent (QIAGEN, Hombrechtikon, Switzerland). Total RNA was extracted from 350 µl of the lysates using Direct-Zol RNA Mini Prep System with on-column DNase I treatment according to the manufacturer’s (Zymo Research, Irvine, CA, United States) protocol. The RNA integrity numbers and RNA concentration were determined by RNA ScreenTape system with 2,200 Tapestation (Agilent Technologies, Santa Clara, CA, United States) and RNA HS Assay Kit with Qubit 3.0 Fluorometer (Thermo Fisher Scientific, Waltham, MA, United States), respectively.

For Gene Expression Profiling (GEx) library construction, QuantSeq 3’ mRNA-Seq Library Prep Kit FWD for Illumina (Lexogen GmbH, Wien, Austria) was applied according to the manufacturer’s protocol. The quality and quantity of the library were determined by using High Sensitivity DNA1000 ScreenTape system with 2,200 Tapestation (Agilent Technologies, Santa Clara, CA, United States) and dsDNA HS Assay Kit with Qubit 3.0 Fluorometer (Thermo Fisher Scientific, Waltham, MA, United States), respectively. Pooled libraries were diluted to 1.8 pM for 1 × 86 bp single-end sequencing with 75-cycle High Output v2.5 Kit on the NextSeq 500 Sequencing System (Illumina, San Diego, CA, United States) according to the manufacturer’s protocol. RNA-sequencing datasets of the human samples are stored in the ArrayExpress database with the following accession number: E-MTAB-10720. Out of this data set, here, we present the SST and SSTR1-5 genes-related sequencing data obtained from control human cardiac samples.

RNA Scope^® in situ hybridization assay was performed on left ventricle tissue slides harvested from human control hearts (the same study as described previously: IK-NPIA-0021–14/1426/18 project 4) using RNA Scope^® Multiplex Fluorescent Kit v2 according to the manufacturer’s instructions (Advanced Cell Diagnostics Pharma Assay Services, Newark, CA, United States). Briefly, 4 µm formalin-fixed paraffin-embedded tissue sections were pretreated with heat and protease prior to hybridization with the following target oligo probes: 3plex-Hs-Positive Control Probe (catalog number: 320861), 3plex-Hs-Negative Control Probe (catalog number: 320871), Hs-SSTR1-C1 (catalog number: 310581, accession no.: NM_001049.2), Hs-SSTR2-C1 (catalog number: 310571, accession no.: NM_001050.2), Hs-TAGNL-C3 (catalog number: 498961-C3, accession no.: NM_003186.3), Hs-PECAM1-O1-C3 (catalog number: 487381-C3, accession no.:. NM_000442.4), and Hs-RYR2-C2 (catalog number: 415831-C2, accession no.: NM_001035.2; see also Supplementary Table S2 for the list of probes). Cell type-specific markers were used to identify cardiomyocytes with a probe recognizing the mRNA of Ryanodine receptor 2 (RYR2) (Lanner et al., 2010), endothelial cells with a probe recognizing the mRNA of platelet endothelial cell adhesion molecule 1 (PECAM-1) (Feng et al., 2004), and vascular smooth muscle cells with a probe recognizing the mRNA of Transgelin (TAGLN) (Camoretti-Mercado et al., 1998), respectively. Preamplifier, amplifier, and AMP-labeled oligo probes were then hybridized sequentially, followed by chromogenic precipitate development. Each sample was quality controlled for RNA integrity with a positive control probe specific to the housekeeping genes with a negative control probe. The pretreatment conditions were optimized to establish the maximum signal-to-noise ratio. Specific RNA staining signal was identified as red/green punctate dots. Nuclei were stained with 4’,6-diamidino-2-phenylindole (DAPI) appearing light purple. Imaging was performed with Leica DMI8 Confocal microscope.

GO enrichment analysis (database version released on August 10, 2020) was performed for each possible comparison of the experimental groups. To obtain GO biological process terms enriched among differentially expressed genes compared to the Sus scrofa reference gene list, online PANTHER Overrepresentation Test [geneontology.org, version released on July 28, 2020 (Mi et al., 2019)] was utilized. Enrichment analysis was conducted by applying Fisher’s exact test with false discovery rate adjustment for multiple comparisons (Benjamini and Hochberg, 1995).

Statistical analyses and Rout outlier analyses were performed and graphs were created using GraphPad Prism version 8 (GraphPad Software, San Diego, CA, United States). The Kruskal-Wallis test with Dunn’s post hoc test, one-way analysis of variance (ANOVA), and unpaired t-test were used to find statistically significant differences. Differences were considered significant at values of p < 0.05 (p values are indicated as *p < 0.05; **p < 0.01; ***p < 0.001). Unless noted otherwise, all data represent the mean ± SEM. To avoid the possibility of overlooking significant differences due to small group sizes, ANOVA-like nonparametric bootstrap-based comparison of means with 1,000 times resampling (Davison and Hinkley, 1997) was also performed on relative expression ratios assessed by RT-qPCR.

In order to investigate the cardiocytoprotective effect of SST, in vitro simulated ischemia/reperfusion experiments were performed in H9C2 cells and adult rat cardiomyocytes, respectively. Simulated ischemia/reperfusion significantly decreased the H9C2 and adult rat cardiomyocyte cell viability (Supplementary Figure S1) compared to normoxic controls. The simulated ischemia/reperfusion-induced cell death was reversed by SST treatment at 100 and 300 nM (Figure 1C) showing a concentration-dependent, bell-shaped cardiocytoprotective effect in H9C2 cells. SST did not influence the viability of normoxic cells (Figure 1B), except for the highest concentration of SST (1 µM) that caused a significant increase of cell death in H9C2 cells. SST also shows a concentration-dependent, bell-shaped cardiocytoprotective effect against simulated ischemia/reperfusion-induced injury in adult rat cardiomyocytes (Figure 2C) where SST treatment at 1, 10, 100, and 300 nM concentrations increased the viability significantly. Although the cell viability shows a similar dose-response pattern in the SST-treated groups in normoxic conditions (Figure 2B), these changes were not statistically significant.

To determine SST-like immunoreactivity in pig plasma and left ventricle samples, RIA was performed. SST-like immunoreactivity was detected in pig left ventricle samples, but no differences were determined between groups (Figure 3C). Significantly increased plasma SST-like immunoreactivity was measured in ischemic preconditioning samples compared to the sham group (Figure 3B).

In order to detect SST and its receptor mRNA expression in pig left ventricle, RT-qPCR and bioinformatics analysis of mRNA-sequencing data were performed. We also performed a bootstrap-based comparison of means to confirm the RT-qPCR results. Expression of SSTR1 and SSTR2 mRNA was detected by PCR, but not SST and its other receptors (SSTR3, SSTR4, and SSTR5) (Figure 4A). The results of the bioinformatics analysis of mRNA-sequencing data confirmed all these results (Table 1). The relative expression of SSTR1 was significantly upregulated in the remote ischemic conditioning group compared to the ischemia/reperfusion group (Figure 4B). There were no significant differences in SSTR2 mRNA expression between any groups (Figure 4C). These results were also confirmed by bootstrap analysis.

To determine SST-like immunoreactivity in human interventricular septum samples, RIA was performed. The patient’s clinical characteristics are described in Figure 5A. A significant decrease of tissue SST-like immunoreactivity was measured in ICM samples compared to the control group (Figure 5B).

To detect SST and its receptor mRNA expression in human left ventricles bioinformatics analysis of the mRNA-sequencing data was performed. The expression of the mRNA of SSTR1, SSTR2, and SSTR5 receptors was detected, but not SST and its other receptors (SSTR3 and SSTR4) (Table 2).

To determine the cell type-specific localization of mRNA of SSTR1 and SSTR2 in human healthy left ventricular samples, RNA Scope^® In Situ Hybridization assay was performed (Figures 6–8). Expression of mRNA of both SSTR1 and SSTR2 was shown primarily in PECAM-1 mRNA-positive endothelial cells (Figures 6A,B, respectively); however, both SSTR1 and SSTR2 mRNA were detected in other cell types, including TAGLN mRNA-positive vascular smooth muscle cells (Figure 7) and RYR2 mRNA-positive cardiomyocytes (Figure 8). There was no detectable signal on the negative control slides (Supplementary Figure S2).

To identify the biological processes that could play a role in the cytoprotective effect of SST, GO analysis of all differentially expressed mRNAs was performed for each possible comparison of the experimental groups. The results of the comparison between remote ischemic conditioning and ischemia/reperfusion groups—where SSTR1 expression significantly increased—are presented (Supplementary Table S3). Differentially expressed mRNAs were significantly associated with, e.g., cardiac muscle differentiation, skeletal muscle development/regeneration, and response to oxidative stress (Figure 9).

SSTR4-specific primer pairs were designed using a partial coding sequence. The available mRNA sequence of pig SSTR4 is not covering the complete region encoding the protein sequence from start to stop codon. Furthermore, the corresponding SSTR4 gene is missing from the used Sus scrofa reference annotation; therefore, it could not be measured by RNA-sequencing. Furthermore, we could not provide functional evidence for the SSTR receptor-mediated mechanisms due to the lack of selective receptor antagonists.