Engineered extracellular vesicle-mediated delivery of miR-199a-3p increases the viability of 3D-printed cardiac patches

Abstract

316In recent years, extrusion-based three-dimensional (3D) bioprinting is employed for engineering cardiac patches (CP) due to its ability to assemble complex structures from hydrogel-based bioinks. However, the cell viability in such CPs is low due to shear forces applied on the cells in the bioink, inducing cellular apoptosis. Herein, we investigated whether the incorporation of extracellular vesicles (EVs) in the bioink, engineered to continually deliver the cell survival factor miR-199a-3p would increase the viability within the CP. EVs from THP-1-derived activated macrophages (MΦ) were isolated and characterized by nanoparticle tracking analysis (NTA), cryogenic electron microscopy (cryo-TEM), and Western blot analysis. MiR-199a-3p mimic was loaded into EVs by electroporation after optimization of applied voltage and pulses. Functionality of the engineered EVs was assessed in neonatal rat cardiomyocyte (NRCM) monolayers using immunostaining for the proliferation markers ki67 and Aurora B kinase. To examine the effect of engineered EVs on 3D-bioprinted CP viability, the EVs were added to the bioink, consisting of alginate-RGD, gelatin, and NRCM. Metabolic activity and expression levels of activated-caspase 3 for apoptosis of the 3D-bioprinted CP were evaluated after 5 days. Electroporation (850 V with 5 pulses) was found to be optimal for miR loading; miR-199a-3p levels in EVs increased fivefold compared to simple incubation, with a loading efficiency of 21.0%. EV size and integrity were maintained under these conditions. Cellular uptake of engineered EVs by NRCM was validated, as 58% of cTnT⁺ cells internalized EVs after 24 h. The engineered EVs induced CM proliferation, increasing the ratio of cell-cycle re-entry of cTnT⁺ cells by 30% (Ki67) and midbodies+ cell ratio by twofold (Aurora B) compared with the controls. The inclusion of engineered EVs in bioink yielded CP with threefold greater cell viability compared to bioink with no EVs. The prolonged effect of EVs was evident as the CP exhibited elevated metabolic activities after 5 days, with less apoptotic cells compared to CP with no EVs. The addition of miR-199a-3p–loaded EVs to the bioink improved the viability of 3D-printed CP and is expected to contribute to their integration in vivo.

1. Introduction

317Heart failure (HF) is considered a growing public health concern, affecting millions of people worldwide^[1]. HF is the result of a myocardial infarction (MI), defined by extensive cardiac muscle death due to ischemia^[2]. The adult human heart lacks regenerative capacity, since adult cardiomyocytes (CM) do not proliferate after damage. In the last three decades, research in the field of tissue engineering (TE) has focused on the design of engineered cardiac patches (CP)^[3,4]. These myocardium substitutes, fabricated and cultured ex vivo, should be functional, contractile heart tissue, and transplantable to replace the injured heart tissue^[5,6]. CP designs were shown to improve cellular delivery rates, while also exhibiting clinically relevant cardiac graft size and appropriate functionality^[6-9]. Within TE approaches, 3D bioprinting has emerged as a fabrication method which successfully captures the intricacy of the native cellular composition and matrix structure of mimetic heart tissue^[10-13]. In particular, extrusion-based 3D printing is a relatively simple and affordable fabrication approach; it allows the use of multiple biomaterials including hydrogels, while cell deposition density can reach 10⁸ cells/mL, crucial for cardiac TE^[14-16]. Nevertheless, extrusion-based 3D-bioprinted CPs exhibit relatively low cell viability. The cell-laden bioink is extruded through a needle by force, producing high shear forces applied onto the cells, damaging their membrane and reducing survival rates post-printing by ~50%^[17].

Stimulation of heart regeneration-related processes was explored as a therapeutic approach and has been incorporated within investigated TE treatments. Inclusion of growth factors and small non-coding RNA species within hydrogel-based delivery systems was shown to promote cardiac recovery through induction of anti-apoptotic signals, CM proliferation, modulation of inflammatory response, and attenuation of fibrotic scar formation^[18-23]. There are advantages of using miRNA-based therapy, such as their small size, the ability to target several genes in a given pathway, and the ability to rapidly develop new therapies for many genes that are not “drugable”^[24,25]. Thus, the ability to therapeutically manipulate miRNA expression and function through systemic or local delivery of miRNA inhibitors, referred to as anti-miRs, has triggered enthusiasm for miRNAs as novel therapeutic targets in the treatment of cardiovascular diseases^[26]. Eulalio et al. performed a functional screening study to identify which human miRNAs can promote CM proliferation in neonatal stages. Two of these miRNAs, miR-199a-3p and miR-590-3p, were further investigated, which were shown to activate the transcriptional cofactor YAP, a known signaling pathway regulating CM proliferation, and were demonstrated to induce cardiac regeneration in an adult mouse MI model^[27,28]. MicroRNA miR-199a-3p was also connected to cell survival through inhibition of caleolin-2 activity^[29].

An emerging, natural carrier of such signaling biomolecules are extracellular vesicles (EVs). EVs are nanoscale membrane-enclosed vesicles that are originated and secreted by living cells. The cargo of EVs frequently includes proteins, mRNA, and miRNA. The binding of EVs to the surface of recipient cells is mediated by the classical adhesion molecules involved in cell–cell interactions, including integrins, intercellular adhesion molecules (ICAMs), and heparan sulfate proteoglycans (HSPGs), increasing target efficiency^[30].

EVs function in cellular communication has been extensively investigated^[31]; prominent examples were illustrated by cancer cells^[32,33] and in the cardiovascular system^[34]. For instance, monocyte-derived EVs were shown to influence migration of endothelial cells (ECs) via EV- mediated delivery of miR-150, promoting angiogenesis^[35], while administration of mesenchymal stem cell-derived EVs was demonstrated to contribute to CM survival, reduced inflammation, and diminished oxidative stress in vitro and in vivo^[36,37]. Monocyte-derived macrophages (MΦ) participate in cardiac repair after MI, as part of an acute inflammatory response. Activated M1-MΦs remove dead cells and their surrounding extracellular matrix by phagocytosis, while also secreting proteases and reactive oxygen species (ROS). Another signaling pathway MΦ used to communicate with surrounding cells is by EV secretion and uptake. Alongside their role in the inflammatory response, M1-MΦs were also shown to secrete an abundant amount of EVs, promoting fibroblast inflammation and suppressing cardiac fibroblast proliferation following MI, the latter was also affected by EVs secreted by other cell types^[38].

Here, an engineered, M1-like MΦ derived-EV system for sustained delivery of miR-199a-3p, known for inducing proliferation and survival of CM, is proposed. It is suggested that incorporation of the engineered EVs within the bioink would increase cell viability and improve survival rate within 3D-bioprinted CPs.

2. Materials and methods

2.1. Materials

Roswell Park Memorial Institute (RPMI) 1640 medium was obtained from Gibco (Gaithersburg, MD). Sodium Pyruvate, L-glutamine, penicillin/streptomycin, and heat-inactivated fetal bovine serum (FBS) were obtained from Biological Industries (Beit-Haemek, Israel). Other reagents, unless specified otherwise, were purchased from Sigma-Merck (Rechovot, Israel). Sodium alginates (VLVG, 318LVG; >65% guluronic acid monomer content) were from FMC Biopolymers (Drammen, Norway). All reagents were of analytical grade.

2.2. Cell culture

Human monocytic THP-1 cells (ATCC^® TIB-202) were cultured in RPMI 1640 supplemented with 2 mM L-glutamine, 1 mM sodium pyruvate, penicillin (100 U/mL), and streptomycin (100 µg/mL). THP-1 monocytes were seeded at a density of 0.3 × 10⁶ cells/mL and differentiated into MΦs by incubation with 100 ng/mL phorbol 12-myristate 13-acetate in serum-free RPMI medium for 2 days followed by a 24-h incubation in RPMI culture medium. MΦ activation was performed by 24-h incubation with serum-free RPMI medium, supplemented with 20 ng/mL human recombinant interferon gamma (IFN-γ; Peprotech^®, #300-02) and 10 pg/mL of lipopolysaccharide.

Primary human umbilical vein endothelial cells (HUVEC) from ATCC^® (PCS-100-013) were cultured with Vascular Cell Basal Medium (ATCC^® PCS-100-030), supplemented with the Endothelial Cell Growth Kit-VEGF (ATCC^® PCS-100-041) following the kit’s instruction.

2.3. Neonatal rat cardiomyocytes isolation

The study was performed with the approval and according to the guidelines of the Institutional Animal Care and Use Committee of Ben-Gurion University of the Negev, Beer Sheva Israel (IL-72-09-2020A). Neonatal cardiac cells were isolated from the ventricles of 1–2-day-old neonatal Sprague-Dawley rats using 6–7 cycles (20 min each) of enzymatic digestion with collagenase type II (95 U/mL; Worthington, Lakewood, NJ) and pancreatin (0.6 mg/mL; Sigma). The cell pellets were resuspended in M-199 culture medium, supplemented with 100 U/mL penicillin and 100 mg/mL streptomycin and 5% (v/v) FBS. To enrich the CM population, the cell suspension was pre-plated twice for 1 h each on an uncoated plate to separate the non-CM. After pre-plating, the non-attached cells (mostly CM) were collected. The isolated cells consisted of over 60% CMs (Figure S1 ^{(899.4KB, pdf)} in Supplementary File).

2.4. EV isolation and characterization

2.4.1. EV isolation

EVs were isolated from culture media using differential centrifugation and ultrafiltration methodologies. THP-1 derived MΦs were washed with phosphate-buffered saline (PBS), and the medium was replaced with fresh, serum-free RPMI 1640 medium. To increase EV yield, the medium was supplemented with 100 mM ethanol. After incubation for 48 h, culture media was collected on ice, centrifuged at 3,500 ×g for 10 min to remove cell debris. Supernatant was collected and centrifuged at 10,000 ×g for 30 min for apoptotic body elimination. Supernatant was filtered through a 0.22-µm filter, followed by two-step ultrafiltration, using 100-kDa MWCO Amicon™ centrifugal filters (Sigma- Merck), according to manufacturer’s protocol.

2.4.2. Nanoparticle tracking analysis (NTA) of size and concentration

Size distribution and concentration of isolated EVs were measured using NanoSight NS300 system (Malvern, UK). EV samples were diluted in PBS until individual nanoparticles could be tracked. The samples were captured for 60 s at room temperature. NTA software was used to measure particle concentration (particles/mL) and size distribution (in nanometers). For each sample, five measurements were taken, and the mean value was determined.

2.4.3. Cryo-transmitting electron microscopy (Cryo-TEM)

For cryo-TEM, 5 μL of the EV sample was applied to a copper grid coated with perforated lacy carbon 300 meshes (Ted Pella) and blotted with filter paper to obtain a thin liquid film of solution. The blotted sample was immediately plunged into liquid ethane at its freezing point (-183°C) with an automatic plunger (Lieca EM GP). The vitrified samples were stored in liquid nitrogen until analysis. Sample analysis was carried out with a FEI Tecnai 12G2 TEM at 120kV with a Gatan cryo-holder maintained at -180°C. Images were recorded digitally using the Digital Micrograph 3.6 software (Gatan, Munich, Germany). Crystal size of purified EVs was measured using ImageJ 1.53q software (U.S. National Institutes of Health, Bethesda, Maryland, http://imagej.nih.gov/ij/).

2.4.4. Western blotting for exosome markers

THP-1-derived MΦs and isolated EVs were lysed in RIPA Buffer (Thermo Fisher Scientific) supplemented with protease and phosphatase inhibitor cocktails (Roche), followed by centrifugation at 14,000 ×g for 15 min at 4°C. Protein quantities of the lysates were quantified using the Pierce BCA Protein Assay Kit (Thermo Fisher Scientific). For Western blotting, fractionation of lysates was carried in sodium dodecyl sulfate polyacrylamide gel electrophoresis on NuPAGE 4%–12% Bis-Tris Protein Gels (Thermo Fisher Scientific). Then, fractionated samples were transferred onto nitrocellulose membranes for immunoblotting. The membranes were blocked in Tris-buffered saline containing 5% powdered bovine serum albumin (BSA) for 1 h at room temperature, and then incubated overnight at 4°C with anti-TSG101 rabbit polyclonal antibody (Abcam) and anti- Calnexin rabbit polyclonal antibody (Abcam), which were diluted 1:1000. After incubation with an appropriate anti- HRP conjugate (Thermo Fisher Scientific), bands were visualized with a chemiluminescence reagent (Biological Industries) and detected using the ChemiDoc system (Bio Rad, CA).

3192.5. Electroporation of EVs

EVs were directly enriched using electroporation performed on a Neon Transfection System (Thermo Fisher Scientific) following the manufacturer’s protocol. Briefly, 1 × 10¹⁰ EVs and 240 pmol miRNA were mixed, and the final volume was adjusted to 100 µL using the electroporation buffer. EVs were loaded with two miRNAs in selected experiments: cel-miR-39 spike-in (ArrayControl™ RNA Spikes, Invitrogen) and hsa-miR-199a-3p (Syntezza Bioscience, Jerusalem, Israel). Electroporation of the EV- miRNA mixture was carried using a pulse width of 20 ms and different voltages (500, 750, 850 V) and numbers of pulses (5 and 10), according to the manufacturer’s protocol. The mixture was then incubated for 30 min at 37°C and overnight at 4°C. Naïve EVs (nEVs) and simple incubation of EVs with miRNAs (iEVs) were used as controls in selected experiments. To remove unbound miRNA residues, samples were filtered using 100 kDa MWCO Amicon™ centrifugal filters, according to manufacturer’s protocol. The concentration and size distribution of engineered EVs was determined by NTA as described above. The yield of engineered EVs following electroporation was calculated as follows: (amount of EVs in electroporated sample) / (amount of EVs in simple incubation sample), based on the area under the curves (AUC) of the size distribution plots.

2.6. RNA isolation and RT-PCR

Total RNA was extracted and isolated from EVs samples using the miRNeasy Mini Kit (QIAGEN) according to the manufacturer’s protocol. RNA concentration of samples was quantified using a spectrophotometer (Nanodrop).

For miRNA analysis, Taqman PCR Kit (Invitrogen) was used. Briefly, RNA samples were reverse-transcribed using the high-capacity cDNA Reverse Transcription Kit (Applied Biosystems, Foster City, California, USA) and obtained cDNA samples were used to quantify miRNAs levels. Experiments were performed using 10 ng of cDNA for each reaction according to the manufacturer’s protocol (Applied Biosystems). Reactions were run on a StepOnePlus‐applied detection system (Applied Biosystems). Relative miRNA expression levels were calculated by the ΔΔCt method using U6 small nucleolar RNA as control.

To generate of a calibration curve for absolute quantification of miRNA, synthetic hsa-miR-199a- 3p (Syntezza) was quantified spectrophotometrically (NanoDrop One, Thermo Fisher Scientific), and 240 pmol were reverse-transcribed. The obtained cDNA was serially diluted 1:5 to have 10 dilutions. Serial dilutions were run in three replicates. The calibration curve was used to convert the Ct values of each sample into the corresponding amount of miRNA molecules or mole. The Pearson’s correlation coefficient calculated for the calibration curve was above 0.99. The loading efficiency into EVs was calculated as follows: (miR-199a-3p mol/EV in sample) / (miR-199a-3p mol/EV used to engineer EVs).

2.7. EV labeling and cellular uptake

Isolated EVs were fluorescently stained using CFSE cell division Tracker kit (BioLegend). Equal volumes of EVs resuspended in PBS were mixed with 40 µM of CFSE solution in PBS, then incubated for 1 h at 37°C. Excess dye was removed using 100-kDa MWCO Amicon™ centrifugal filters, according to manufacturer’s protocol. The same protocol was applied on PBS samples without EVs as a control.

EV internalization by neonatal rat cardiomyocytes (NRCM) and HUVECs was visualized by laser scanning confocal microscopy (Nikon C1si). Cells were seeded onto eight-well µ-slides (ibidi^®, 80826) coated with 0.1% Gelatin (w/v). After 24 h post-seeding, cells were washed once with PBS, then incubated with stained EVs (6 × 10⁹ EVs/mL, in culture medium without serum addition) for 24 h. Cells were then washed twice with PBS, fixed, and stained as outlined below. For EC culture, the percentage of cells containing EVs was quantified by flow cytometry, compared with controls of non-treated cells and cells treated with filtered CFSE dye. A population of single cells was gated, and a minimum threshold of fluorescence was set according to negative control samples in each experiment. All results above the minimum threshold were considered positive. For NRCM culture, EV internalization rate was determined by the number CM (cTNT⁺ cells) containing EVs normalized to the total number of cTNT⁺ cells per field. Multiple fields were analyzed in each sample.

2.8. Proliferation and cytokinesis assays

For the proliferation assay, miR-199a-3p-engineered EVs or controls (cel-miR-39-engineered EVs or PBS) were added to NRCM monolayers and incubated for 24 h. After an additional 24 or 48 h, cells were fixed, permeabilized, and blocked as detailed below. Cells were then stained overnight at 4°C with cardiac troponin T (ab8295, Abcam) and Ki67 (ab16667, Abcam) or Aurora B (ab2254, Abcam) primary antibodies and then secondary antibodies conjugated to Alexa Fluor488 and 647. Images were acquired with Nikon C1si laser-scanning confocal microscope. To calculate the ratio of proliferating CMs, the amount of Ki67⁺ nuclei in the CMs (cTNT⁺ cells) was quantified using ImageJ thresholding and normalized to the total number of cTNT⁺ cells per field. To quantify the ratio of mitotic CMs, midbodies⁺ CMs were counted and normalized to the total number of cTNT⁺ cells per field. Multiple fields were analyzed in each sample.

2.9. Tube formation assays

320HUVECs of passages 5 were resuspended in growth medium without vascular endothelial growth factor (VEGF), supplemented with either miR-199a-3p-engineered EVs (6 × 10⁹ EVs/mL) or controls (naïve MΦ-EVs or PBS), and then seeded onto Matrigel (Corning)-coated, 48-well plates at a concentration of 4 × 10⁴ cells/ cm² for 18 h. For positive control, HUVECs resuspended in growth medium supplemented with 20 ng/mL VEGF were used. After 18 h, the cells were washed in PBS and labeled with 5 µM CFSE (in PBS) for 20 min, and then washed in growth medium. Image acquisition was carried using a fluorescent microscope. The total number of junctions and cumulative vessel length in each image were quantified using Angiotool software (version 0.6a, NIH/ NCI, USA)^[39] and normalized to the values of the positive control.

2.10. Cardiac patches 3D bioprinting

2.10.1. Bioink preparation

Alginate (LVG) was covalently modified with RGD peptide as previously described^[40]. RGD-modified alginate solution (LVG-RGD) was prepared by dissolving lyophilized alginate-RGD in double-distilled water under stirring for 2 h. Fifteen percent gelatin stock solution was prepared by dissolving bovine gelatin (Type B) in Dulbecco’s Modified Eagle Medium (DMEM) under stirring for 1 h at 37°C. Crosslinking of alginate was achieved by mixing 1.8 mL of 2.5% w/v LVG-RGD with 0.3 mL of 3% w/v D-gluconic acid salt solution, using a homogenizer to equally distribute the calcium ions throughout the solution. The mixture was further stirred at 37°C until used. Next, 0.6 mL of gelatin stock solution was added to the cross-linked alginate, allowed to mix for 10 min before being transferred to a sterile syringe. Following gelatin addition, 0.3 mL of isolated NRCM, either mixed with 1.2 × 10¹¹ engineered EVs/mL solution or PBS, were loaded into an additional sterile syringe, which was then immediately connected to the polymer-loaded syringe through a Luer-to-Luer connector. The solutions were mixed by gently pushing the pistons back and forth for 1 min, until the solutions were homogenously mixed. The final bioink solution consisted of 1.5% LVG-RGD (w/v), 0.3% (w/v) D-gluconic acid salt, 3% Gelatin, 1 × 10⁷ NRCM/mL, and 6 × 10¹⁰ EVs/mL.

2.10.2. Bioink rheological characterization

The viscoelastic properties of bioink were analyzed using a stress control rheometer (TA Instruments, model AR 2000), operated in the cone-plate mode with a cone angle of 1° and a 60-mm diameter. Storage (G′) and loss (G″) moduli were measured in a frequency range of 0.1–10 Hz, while the apparent viscosities (Pa*s) of the bioink solutions were assessed at different shear rates between 0.1 and 10 s⁻¹. The measuring device was equipped with a temperature control unit (Peltier plate, ±0.05°C) operated at 28°C (Figure S2 ^{(899.4KB, pdf)} in Supplementary File)

2.10.3. 3D bioprinting and cardiac patches maintenance

Prior to bioprinting, the bioink was deposited into a sterilized 30 mL printer cylindrical cartilage sealed with a fit plunger, using a Luer-to-Luer connector. The printer cartilage was then sealed and centrifuged for 1 min at 150 ×g to remove remaining air bubbles. A sterile 25-gauge needle tip was connected to the cartilage, and a cap connecting the print head to the barrel was added. The barrel was put in the low-temperature head of the bioprinter (EnvisionTEC 3D-bioplotter Developer Series, Germany), set to 28°C. The bioink solution was allowed to reach the printing head temperature for 30 min before bioprinting began.

Three-dimensional bioprinting was performed using the freeform reversible embedding of suspended hydrogel (FRESH) approach, as previously described^[41]. The cardiac patches were printed onto a 12-well plate filled with FRESH gelatin supporting bath at room temperature, using applied pressure of 0.5 bar and a printing velocity of 12 mm/s. Patches were 1 cm in diameter and 2 mm thick, printed in 14 layers, and fabricated using an infill pattern of 90° grids with 0.6 mm spacing (center-to-center) and a 30% overlap between printed layers. Patches were printed in triplicates. Computer-aided design models of the patches and grids printed were created using SOLIDWORKS software and imported as STL files to the printing control system through the Bioplotter RP software. Following bioprinting, patches were incubated for 1 h at 37°C with 5% CO₂ to free the 3D-printed constructs from the gelatin support bath. The patches were then washed and incubated for 10 min at 37°C in DMEM supplemented with 0.1 M CaCl₂ to further crosslink the 3D-bioprinted constructs. Following incubation, the cardiac patches were incubated in DMEM growth medium supplemented with 20% FBS in new 12-well plate. Medium was changed daily for each cardiac patch until analysis.

2.11. Mechanical stiffness of 3D-bioprinted CPs

The elastic modulus (Young’s modulus) of the 3D-bioprinted CPs was analyzed by an Instron 4505 mechanical tester (courtesy of Prof. Ronit Bitton, Ben-Gurion University of The Negev, Israel) equipped with a 100 N load cell. The crosshead speed was set to 5 mm/min, and load was applied until the specimens were compressed to approximately 100% of the original thickness. The elastic modulus was calculated as the slope of the initial linear portion of the stress–strain curve (n = 4).

2.12. Printability analysis

321The printability analysis examined the effectiveness of the extruded strands in the test grids to form square holes between strands, as previously described^[42]. Circularity (C) of an enclosed area is based on the shape perimeter and area, where a perfect circle has a circularity of 1. For a square shape, circularity is equal to π/4, and the printability is given by Equation I:

$$Pr=\frac{L^2}{16A}$$ (I)

where L is the perimeter, and A is the area of a shape. A printability of 1 is equal to a perfect square and indicates optimal printing conditions of a bioink. Confocal images of three-layer high test grids, printed as described above, were evaluated by measuring the perimeter and area of several holes in each sample, and printability was calculated, with 4 constructs and at least 25 holes measured.

2.13. Cryogenic scanning electron microscopy (Cryo-SEM)

Samples of a cellular, 3D-bioprinted CPs, were fabricated as described above. Dissected samples were placed and sandwiched between two aluminum disks (3 mm in diameter, each 25-μm thick) and cryo-immobilized in a high-pressure freezing device (EM ICE; Leica).

The frozen samples were then mounted on a holder under liquid nitrogen in a specialized loading station (EM VCM; Leica) and transferred under cryogenic conditions (EM VCT500; Leica) to a sample preparation freeze fracture device (EM ACE900; Leica). In that device, the samples were fractured by the rapid stroke of a cryogenically cooled knife, exposing the inner part of the sandwiched disks. After fracturing, the samples were etched at -100°C for 10 min to sublime ice from the sample surface, and were coated with 3 nm carbon.

Samples were imaged in a Gemini SEM (Zeiss) by a secondary electron in-lens detector, while maintaining an operating temperature of −120°C (Figure S3 ^{(899.4KB, pdf)} in Supplementary File). The measurements were done at the Ilse Katz Institute for Nanoscale Science and Technology at Ben-Gurion University of the Negev, Beer Sheva, Israel.

2.14. Flow cytometry

On days 1 and 5 post-printing, the cardiac patches were dissociated with citrate buffer (4% w/v in PBS, pH 7.4) to release encapsulated cells, and were washed twice in FACS buffer (2% FBS in PBS). For cellular viability analysis, cells were stained using the Live/Dead viability/cytotoxicity kit (Invitrogen) according to the manufacturer’s protocol.

For protein analysis, suspended cells were fixed and permeabilized with the Cytofix/Cytoperm kit (BD Pharmingen) according to the manufacturer’s protocol. The cells were immunostained with relevant antibodies in FACS buffer (2% FBS in PBS); 1 h with primary antibodies followed by 30 min secondary antibody. Data acquisition and analysis were performed using a FACS-Canto machine (BD Biosciences) utilizing Cellquest Pro software (BD Biosciences) and FlowJo X 10.0.7. For quantification of cardiac protein expression levels, mean fluorescent intensity was calculated based on the intensity distribution plot of each protein marker. Gating of CM population was determined according to the intensity of sarcomeric α-actinin (Clone EA-53, 1:300, Sigma), setting the threshold based on secondary antibody control and pre-plated cardiac fibroblasts culture population expression levels.

2.15. Cell metabolic activity and DNA content

Cell metabolic activity was measured by PrestoBlue reagent (Invitrogen). The 3D-bioprinted constructs were transferred into new wells and incubated with 600 µL of PrestoBlue reagent mixture (1:9 in DMEM culture medium supplemented with 20% v/v FBS) for 3 h at 37°C. Aliquots (300 µL) were placed in a 96-well plate and the fluorescence was measured at excitation wavelength of 560 nm and emission wavelength of 590 nm using a plate reader (model ELX 808, BIO TEK Instruments, Winooski, VT). Samples of the reagent mixture incubated under the same conditions without cells were used as a blank. Cell DNA content was quantified using bisbenzimidazole Hoechst 33258 fluorescent dye (Sigma). The constructs were dissolved in 500 µL sodium citrate (4% w/v in PBS) to release encapsulated cells, and then centrifuged for 10 min at 6000 rpm and 4°C. The cell pellet was suspended in 100 µL lysis buffer (0.02% v/v SDS) in sodium citrate (0.015 M-saline, pH 7) and incubated for 1 h at 37°C. Then, 100 µL of Hoechst 33258 assay solution (2 mg/mL) was added, followed by a 10-min incubation. Aliquots (180 µL) were placed in a 96-well plate and the fluorescence was measured at excitation wavelength of 485 nm and emission wavelength of 530 nm using a plate reader. Samples of the reagent mixture incubated under the same conditions without cells were used as a blank. Specific metabolic activity was calculated as cell metabolic activity intensity normalized to DNA content quantified by Hoechst.

2.16. Immunocytochemistry and confocal imaging

For immunofluorescence of cell monolayers, cells were fixed in 4% v/v warm methanol-free formaldehyde in PBS for 20 min, washed twice in PBS, permeabilized with 0.1% Triton-X, and blocked with 1% BSA (Millipore, Bedford, MA) for 1 h at room temperature. The samples were incubated overnight with the primary antibodies, 322followed by a 2-h incubation with secondary antibodies, with 10-min PBS washing for three times in between the incubation sessions.

The antibodies details are as follows: mouse anti-sarcomeric α-actinin (Clone EA-53, 1:300, Sigma), mouse anti-cardiac troponin T (cTnT, ab8295, 1:200; Abcam), rabbit anti-ki67 (ab16667, 1:250, Abcam), rabbit anti-active caspase-3 (#9661, Life signaling), Alexa 488-conjugated donkey anti-mouse (715-545-150, Jackson Immuno-Research Labs Inc.), Alexa 488-conjugated chicken anti-rabbit (A21441, Invitrogen), Alexa 647-conjugated donkey anti-rabbit (ab150115, Abcam), and Alexa 647-conjugated goat anti-mouse (ab150075, Abcam).

Alexa-Fluor 546-conjugated phalloidin (A22283, 1:1000, Life Technologies) was used for staining F-actin and NucBlue (Invitrogen) for nuclei detection. Image acquisition was performed with a Nikon C1si laser-scanning confocal microscope.

2.17. Statistical analysis

Statistical analysis was performed with GraphPad Prism version 8.43 for Windows (GraphPad Software, San Diego, CA). All variables are expressed as mean ± SEM from at least two independent experiments. Fold change in miRNA levels, midbodies ratios and total vessel length were compared by one-way analysis of variance (ANOVA) with Tukey’s post-hoc test. Ki67 ratios, specific metabolic activities and apoptotic CM ratios were compared using two-way ANOVA with Tukey’s post-hoc test. Protein expression analysis, DNA content and cellular viability were compared using two-tailed student t-test. P < 0.05 was considered statistically significant.

3. Results and discussion

3.1. Isolation and characterization of macrophage-derived EVs

Considering their role in heart tissue after MI, it was speculated that MΦ-secreted EVs may be taken up by other cells in the cardiovascular system, including CMs. First, isolated EVs were characterized and verified to include only EVs, without cellular contaminations (e.g., cell debris and apoptotic bodies). Nanoparticle tracking analysis (NTA) was used to evaluate EV hydrodynamic diameter and yield (Figure 1A). Isolated nanoparticles exhibited varied size distribution, ranging between 50 and 200 nm in diameter, which are common for purified EVs. Mode hydrodynamic diameter was 84 ± 4.3 nm (mean ± 95% CI, n = 25), corresponding with the typical size range of EVs (30–200 nm).

Figure 1.

Characterization of macrophage-derived EVs. (A) Representative size distribution plot of EVs isolated from activated macrophages. (B) Cryo-transmission electron micrographs of EVs (black arrows; scale bar: 200 nm). (C) Immunoblots of EVs and cell lysate fractions for the EVs marker TSG101 and ER protein calnexin (n = 4).

Cryo-TEM analysis showed typical exosome morphology, exhibiting round shape and a visible double membrane (Figure 1B). Crystal size of isolated EVs (mode size of 106 ± 6 nm, mean ± SEM) was comparable to the hydrodynamic diameter measurements.

Western blot analysis confirmed the exosomal marker tumor susceptibility gene 101 (TSG-101, ESCRT-0 protein) is present on purified EVs and secreting cells, while endoplasmic reticulum (ER) protein calnexin was present in secreting cells but absent in purified EVs (Figure 1C), verifying that isolated nanoparticles are indeed EVs^[43].

3.2. Optimization of electroporation settings for loading miRNA into EVs

In order to engineer the EVs to promote desired cellular processes (i.e., CM proliferation), direct electroporation of 323EVs following isolation was used. Even though electroporation is a widely used methodology for cargo loading into EVs, it also presents major disadvantages, including reduction in EV concentration and damaged membrane integrity^[44,45]. Hence, electroporation settings should be evaluated not only in means of loading efficiency, but also in terms of yield and quality of obtained engineered EVs.

To identify which electroporation setting presents the best efficacy in means of miRNA loading, different voltages and pulses were examined using cel-miR-39 model^[46]. miR-39 expression level was quantified using RT-PCR (Figure 2A). Naïve EVs (nEVs) and EVs simply incubated with miRNA (iEVs) were examined to determine basal levels of inserted miRNA without electroporation. In general, inserted miRNA levels increased with elevated voltage, setting the number of pulses at 5. However, increasing the number of pulses to 10 negatively affected inserted miRNA levels compared with the 5 pulses setup; inserted miRNA levels were comparable to iEVs. The 850V, 5 pulses setup presented the best results compared to both simple incubation and other setups, so we decided to continue with this setting. This protocol was implemented for EV enrichment with miR-199a-3p mimic, a known inducer of CM proliferation, which was meant to be delivered by the EVs to increase its levels by 5 order of magnitudes relative to its basal levels and achieve a fivefold increase compared with simple incubation (Figure 2B).

Figure 2.

Electroporation optimization for EV loading with miRNA. (A and B) Relative expression levels of cel-miR-39 (A) and miR-199a-3p (B) in EVs following application of different electroporation settings, compared with naïve EVs (nEVs) and EVs incubated with free miRNA (iEVs), determined by RT-PCR. Data represent mean ± SEM (n = 3) *P < 0.05, **P < 0.01, ***P < 0.001, **** P < 0.0001; Tukey’s test, one-way ANOVA. (C–F) Electroporation effect on EVs. (C) Representative particle concentration (left) and normalized concentration (right) size distributions for iEVs and EVs after electroporation in 850 V and 5 pulses, measured by NTA. (D) Representative cryo-transmission electron micrographs of EVs after electroporation at 850V (left) and incubation with miRNA (right); scale bar: 200 nm. (E) Size distribution of electroporated EVs (EP-EVs, blue), iEVs (orange) and nEVs (green), measured according to cryo-transmission electron microscopy.

Loading efficiency was also determined, calculated to be 21.0 ± 1.3% (mean ± SEM, n = 4).

EV concentration (yield) and morphology were also examined by NTA (Figure 2C–E). The 850 V, 5 pulses settings presented a slight decrease in EV yield (Figure 2C, left) when compared with simply incubated EVs, evaluated by the relative AUC (0.95 ± 0.10, mean ± SEM, n = 8). The size distribution was barely altered, except possible EVs fusion, evident by the shift in EV mode size (Figure 2C, right). Since electroporation also resulted in a high loading efficacy, EVs obtained by these settings were further evaluated by electron microscopy (Figure 2D). Cryo-TEM imaging of EVs post-electroporation also verified the findings obtained by NTA, as size distribution and mode size of EVs were barely altered compared to iEVs (Figure 2E). Regarding particle morphology, it appears that electroporation did not change the shape of EVs or affect their membrane integrity (Figure 2D), meaning that engineered EVs maintain their stability.

Overall, the optimal setting for loading selective miRNA into MΦs-derived EVs of 850V, 5 pulses was selected for the rest of the experiments, considering its high efficacy with minor compensation in EV yield.

3.3. EVs uptake by neonatal rat cardiomyocytes

Macrophage-derived EVs were selected for miRNA delivery due to the assumption that this type of cell communicates 324with a variety of cardiovascular cells^[47-50]. To validate this assumption, it is essential to show that MΦ-derived EVs are internalized by their target cells. Furthermore, EV uptake kinetics must be investigated to evaluate the time frame in which delivered miRNA affects recipient cells, specifically CM.

Optimal EV concentration and the exposure period in which EVs uptake and cellular accumulation is detectable were determined. NRCMs were treated with CFSE-labeled EVs for 2, 12 and 24 h, at concentrations of 4 × 10⁹ and 8 × 10⁹ EVs/mL, according to previous reports^[22,37]. At the lower concentration, EV uptake was observed only after 24 h (Figure 3A, top right). Higher concentration led to EV internalization, which was detectable after 12 h, but EV accumulation continued through the first 24 h (Figure 3A, bottom left and right). Quantification of EV uptake specifically by CM following 24 h of incubation showed that EVs were present in 58 ± 4% of TNT-immunostained NRCMs. Confocal imaging also confirmed that EVs were indeed internalized by NRCMs (Figure 3B and 3C).

Figure 3.

Neonatal rat cardiomyocytes uptake MΦs-EVs. (A) Optimization of CFSE-labeled MΦs-EVs (green) uptake by NRCM, stained for F-actin (red) and nuclei (blue). Cells were incubated with 4 × 10⁹ EVs/ml or 8 × 10⁹ EVs/ml for 12 h and 24 h. EV-positive cells are indicated by white arrows. Scale bar: 50 µm. (B) Representative images of CFSE-labeled MΦ-EVs found inside NRCMs. Cells were incubated for 24 h without EVs (top panel) or with 6 × 10⁹ EVs/mL (bottom panel). Scale bar: 50 µm. (C) High-magnification images of XY (top) and reconstructed YZ (bottom) planes, 24 h after adding MΦs-EVs (i) and filtered CFSE dye (ii), showing internalization of the labeled EVs into NRCM. Nuclei (blue); Cardiac troponin (red) and EVs (green). Scale bar: 10 µm.

These results demonstrate the applicability of MΦ-derived EVs as a drug delivery vehicle for NRCMs. Taking into consideration the time frame in which miRNA is functional (24–28 h after transfection^[51,52]), a timetable of 48–72 h from EV transfection was established to assess EV treatment efficacy.

3.4. Engineered EVs loaded with miR-199a-3p induce NRCM proliferation

After confirming successful MΦs-EV internalization by NRCM, we further assessed the effect of EV treatment on cardiac regeneration-related cellular processes, such as CM proliferation.

NRCMs were incubated for 24 h with either EVs loaded with miR-199a-3p mimic or EVs loaded with cel-miR-39 mimic. NRCMs presented limited proliferative potential up to 7 days post-natal^[1,53], thus non-treated NRCMs were used as control for basal levels of CM proliferation. After 24 h of incubation, EVs were removed, and NRCM proliferation was examined 24–48 h from EV transfection, using Ki67 staining for cell proliferation (Figure 4A). Twenty-four hours after transfection, significant differences were observed in the ratio of Ki67⁺ NRCM to all NRCM between the miR-199a-3p loaded group and the controls. Approximately 15% of CM were found to be in active cell-cycle stages, compared with 6%–7% of NRCMs in the control group (Figure 4B). A functional effect was observed after an additional 24 h, where the group treated with miR-199a-3p-loaded EVs exhibited an increase in proliferating NRCM ratio (36% Ki67⁺ CM), while the controls were significantly lower (27%). This is an indication that as soon as 48 h post-incubation, miR-199a-3p starts to affect signaling pathways related to cell proliferation.

Figure 4.

Electroporated EVs induce NRCM proliferation and cytokinesis. (A) Representative images of Ki67+ (red) and cTnT+ (green) NRCM (white arrows) to demonstrate increased Ki67 staining 48 h after exposure to miR-199a-3p-electroporated EVs. Scale bar: 50 μm. (B) Quantification of Ki67⁺ cTnT+ NRCM 24 and 48 h from exposure to EVs. Data are mean ± SEM., n = 3–4, *p < 0.05, Tukey’s multiple comparisons test, two-way ANOVA. Quantification was based on counting of Ki67⁺ cTnT+ co-stained cells relative to total cTnT⁺ cells. (C) Representative images of Aurora kinase B⁺ (red) cTnT⁺ (red) NRCM (white arrows) to demonstrate increased cytokinesis 48 h after exposure to miR-199a-3p-electroporated EVs. Scale bar: 50 μm. (B) Quantification of midbodies⁺ cTnT⁺ NRCM 48 h from exposure to EVs. Data are mean ± SEM., n = 3–4, *p < 0.05, Tukey’s multiple comparisons test, one-way ANOVA. Quantification was based on counting of midbodies+ cTnT⁺ co-stained cells relative to total cTnT+ cells.

To further establish the claim that EVs enriched with miR-199a-3p are capable of inducing NRCM proliferation, Aurora B kinase midbodies were quantified 48 h post-transfection, indicating the occurrence of cytokinesis (Figure 4C). While the controls exhibited comparable ratios of midbodies⁺ CM (7.6 ± 0.4% and 8.7 ± 0.5 for non-treated and cel-miR-39-loaded EVs-treated groups, respectively), miR-199a-3p-loaded EVs presented a twofold increase in midbodies⁺ CM ratio (16.7 ± 1.5%). This is a clear indication that miR-199a-3p-loaded EVs induce not only cell-cycle re-entry, but also promote cell mitosis.

3253.5. Engineered EVs present proangiogenic potential

Sufficient blood and nutrient supply to the engineered cardiac patch is essential for long-term survival of residing cells after implementation. In this aspect, EV-mediated communication between MΦs and ECs in the cardiovascular system was vastly investigated, indicating EVs capability to induce EC migration and organization into tubular networks^[35,54,55]. Therefore, it was speculated whether MΦ-EVs could potentially promote cardiac patch vascularization post-printing, thus contributing to cell survival in the CP. The internalization of the MΦ-EVs was examined using a HUVECs model, evaluated following 24 h incubation. Confocal microscopy imaging indicated substantial EV uptake (Figure 5A), confirmed by flow cytometric analysis, showing 94.7 ± 0.4% (mean ± SEM, n = 3) of the cells positive for EV uptake (Figure 5B and 5C). The potential of the MΦ-EVs delivery system to promote angiogenesis was examined by cultivating HUVECs on Matrigel to form tubular networks. HUVECs were mixed with media containing either naïve or miR-199a-3p-engineered EV, and then cultured for 18 h before evaluated for tube formation. Media supplemented with VEGF, a known proangiogenic growth factor, were used for positive control, while HUVECs supplemented with PBS were used for negative control (Figure 5D).

Figure 5.

Engineered MΦs-EVs present pro-angiogenic potential. (A–C) CFSE-labeled MΦs-EVs (green) uptake by HUVECs. (A) Representative images of CFSE-labeled MΦs-EVs found inside HUVECs (white arrows), incubated for 24 h with engineered EVs. Nuclei (blue); F-Actin (red) and EVs (green). Scale bar: 50 µm. (B) Flow cytometric analysis of EV uptake by HUVEC cells (green), compared with non-treated cells (gray) and filtered CFSE dye (orange). (C) Quantification of cellular uptake, according to flow cytometry. (D–F) HUVECs tube formation evaluation. (D) Representative fluorescently labeled HUVECs tube formation following 24 h with engineered EVs (right), naïve EVs (middle) or without treatment (left). (E) Quantification of total vessel length, normalized to positive control. (F) Quantification of the number of junctions. Data are mean ± SEM, n = 3, *P < 0.05, Tukey’s multiple comparisons test, one-way ANOVA.

The EVs-treated groups exhibited improvement in relative total vessel length compared with the negative control (Figure 5E). Moreover, EVs-treated cells also produced a significantly higher amount of junctions per field, indicating a higher number of tubes formed, compared with non-treated cells (Figure 5F). These results show that naïve EVs, secreted by activated MΦs, present intrinsic proangiogenic potential. More importantly, the engineering of MΦ-EVs did not seem to disrupt the proangiogenic function of naïve MΦ-EVs. This is another testament for the benefits of the improved electroporation settings, allowing addition of signaling miRNAs without jeopardizing the EVs natural function. Overall, the engineered MΦ-EVs system was shown to potentially improve cell survival of the whole cardiac patch by contribution to angiogenesis-related processes.

3263.6. Engineered EVs improve CM survival within 3D-bioprinted constructs

A major burden in extrusion-based 3D bioprinting fabrication of a sustainable and functional cardiac patch is the viability of residing CMs post-printing. Therefore, it was speculated that induction of cardiac regeneration-related processes (i.e., cell proliferation and prevention of apoptosis) would improve cell survival following 3D bioprinting. To initiate these processes within the CP-residing cells, miR-199a-3p-engineered EVs were incorporated within the alginate-based, cell-laden bioink (Figure 6A). Bioink without EVs was used to fabricate the CP for the control group. The cell-laden bioink was used to fabricate a large, cylindrical shaped 3D-CP (2 mm high, 1 cm in diameter; Figure 6B, i), exhibiting a printability value of 1.01 ± 0.01. The resulting 3D CP had a Young’s modulus value of 4.2 ± 0.4 kPa 1-day post-printing and was decreasing with time (2.0 ± 0.1 kPa by day 5). Since cells were mixed within the bioink solution, cells were evenly distributed within the entire 3D construct (Figure 6B, ii–iii). According to cryogenic-SEM (Figure S3 ^{(899.4KB, pdf)} in Supplementary File), EVs maintained their sphere-like morphology post-printing.

Figure 6.

Engineered EVs improve cell survival within 3D-bioprinted cardiac patches (CPs). (A) Schematic description of the 3D bioprinting procedure. (B) 3D-bioprinted CPs. (i) Images of the 3D-bioprinted CP (1 cm in diameter, 2 mm high). (B, ii–iii) Confocal images of XY (ii) and reconstructed YZ (iii) planes, 24 h post-3D bioprinting of fluorescently labeled cells. (C and D) Expression levels of the cardiac-specific proteins inside the CPs, after 5 days in culture. (C) Flow cytometric analysis of cTnT⁺ (top) and sarcomeric α-Actinin⁺ (bottom) of NRCM co-printed with engineered EVs (green) or without EVs (red), compared with secondary antibody control (gray). (D) Relative mean fluorescent intensity (MFI) of cardiac-specific proteins. (E and F) Cell viability analysis within the 3D-bioprinted CPs, following 5 days in culture. (E) Relative specific metabolic activity of CP residing cells. (F) DNA content in the CPs relative to day 1 post-printing, quantified by Hoechst. (G) Flow cytometric analysis of relative cellular viability within the 3D-bioprinted CPs, following 5 days in culture. (H) Quantification of activated caspase 3⁺ α-Actinin⁺ NRCM, 5 days post-printing, shows decreased apoptosis among CM co-printed with engineered EVs (mean ± SEM, n = 3).

For evaluation of the CM and exclusion of non-myocytes populations, the cardiac-specific protein markers cTnT and sarcomeric α-actinin were measured 1 and 5 days post-3D bioprinting of the CP (Figure 6C). The expression levels of cardiac proteins inside the patch were decreased over 5 days in culture. Such decrease was previously observed in RGD-modified alginate, 3D cultures of NRCM, suggesting cell-matrix interactions play a key role in the preservation of CM phenotype^[56]. Furthermore, neonatal cardiac cells isolation consists also of non-myocyte populations with higher proliferative capacity in the short term and better adhesive capacity in RGD-modified alginate. Nevertheless, the cardiac-specific proteins profile changed within EV-containing CP, 327exhibiting a more moderate decrease in expression level compared with CP with no EVs (Figure 6D). Macrophage-derived EVs were documented to also influence cardiac fibroblasts (approx. 40% of the isolated culture), inhibiting their proliferative capacity^[57], suggesting the EVs attenuate the shift in cellular composition inside the presented CP.

To assess cell viability, the metabolic activity of the cells within the 3D-printed CP was measured over 5 days in culture. The incorporation of EVs within the bioink had an immediate effect on cellular viability, as EV-containing bioinks produced CPs with twofold greater specific metabolic activity only 1 day after printing, compared to bioink that did not include EVs (2.3 ± 0.2 fold-change in specific metabolic activity). Over 5 days in culture, CPs containing the EVs demonstrated better recovery post-printing, presenting a threefold increase in relative specific metabolic activity, compared to CPs that did not include any EVs (Figure 6E). The EVs-containing CPs also exhibited higher DNA content following 5 days in culture (Figure 6F), indicating possible, however limited, cell proliferation inside the CP.

The effect of EV addition on cellular viability was not limited to the increase in metabolic activity. Live–dead staining analysis of cells from dissociated CPs showed that cellular viability was maintained when EVs were added to the bioink, while cells from CPs without EV exhibited a decrease in cell viability over 5 days in culture (Figure 6G).

Nevertheless, cellular viability analyses are not specific to CMs and therefore provide a limited insight regarding the mechanism behind EVs’ beneficial effect following 3D bioprinting. Considering the increased viability, accompanied with increased DNA content and a general decrease in cardiac-specific proteins expression profile, it was speculated that EVs may improve CM survival following 3D printing. To specifically target this effect in CM, flow-cytometric analysis was performed to evaluate the ratio of apoptotic CMs, staining for activated Caspase-3, a cellular marker for cell apoptosis. Following 5 days in culture, the ratio of apoptotic CM was significantly lower in CPs printed with EV-containing bioink in comparison to bioink without EVs (Figure 6H), suggesting that EVs contribute to cellular viability mainly through attenuation of CM death post-printing.

Yet, there are still concerns regarding possible negative effects of inherent EV cargo on recipient cells outside the 3D-bioprinted patch. However, since the engineered EVs were mixed directly with the cells prior to addition to the bioink and considering EVs are rapidly taken by recipient 328cells^[58], it is speculated that in the proposed system, the EVs mostly affect the bioink residing cells. However, further investigation of the effect of engineered EV released from the patch is required to eliminate these concerns.

4. Conclusion

Herein, we described the successful engineering of a macrophage-derived EV delivery system, capable of affecting multiple processes related to cardiac regeneration, including CM proliferation and cell death attenuation, while also presenting angiogenic potential. The feasibility of miRNA cargo loading, while maintaining the EVs membrane integrity, morphology and functionality, was demonstrated. These attributes have dramatic influence upon the applicability of EV-based delivery systems. EV manipulation generally reduces EV yield^[46,59], affecting treatment efficiency and cost. Moreover, EV stability might affect their uptake and functionality upon delivery. The proposed EV delivery system exhibited capability to harness new properties while maintaining their innate beneficial traits; MΦ-EVs were also shown to affect ECs in a way that could not only promote cell survival within the CP, but also stimulate pre-vascularization if ECs are incorporated within the patch.

Currently, most efforts to improve 3D bioprinting results are focused on changes in the bioink mechanical properties and adjustments of printing parameters^[60-62]. Here, the inclusion of the proposed engineered EVs within a cell-laden bioink contributed to multiple aspects of cellular viability, including CM survival, following 3D bioprinting. Cell vitality is a major burden particularly in engineering of CPs, where high concentrations of cells are required. Proliferation of CM within CPs is considered to play a key role affecting the success of CP viability and integration in vivo^[63]. To the best of our knowledge, this is the first evidence on the successful induction of cardioprotection-related processes in CM post-3D bioprinting. The obtained results could also be harnessed to improve the outcomes of 3D bioprinting for fabrication of more viable complex cellular constructs. Furthermore, the engineered EV delivery system recapitulates multiple regenerative traits, including angiogenesis. These could possibly provide the CP with capability to integrate better with the host after implementation in vivo.

Funding

The work was supported by the Jordan Baruch Stem Cell Fund: 87351721.

Conflict of interest

The authors declare no conflict of interests.

Author contributions

Conceptualization: Assaf Bar, Smadar Cohen

Formal analysis: Assaf Bar

Investigation: Assaf Bar, Olga Kryukov

Methodology: Assaf Bar

Supervision: Smadar Cohen, Sharon Etzion

Writing – original draft: Assaf Bar

Writing – review & editing: Assaf Bar, Sharon Etzion, Smadar Cohen

Ethics approval and consent to participate

The study was carried out in strict accordance with the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. All animal studies reported in this study were approved by the institutional ethics committee of Ben-Gurion University of the Negev, Israel (Protocol IL-72-09-2020(A)).

Consent for publication

Not applicable.

Availability of data

Not applicable.