A New Cardiac Decellularized Extracellular Matrix (dECM)-Based Hydrogel: From Its Development with a Standardized Myocardial Decellularization Procedure to In Vitro Model Applications

TL;DR

A new hydrogel made from pig heart tissue is developed to better mimic the heart's environment for studying diseases and testing drugs.

Contribution

A novel cardiac dECM-based hydrogel is developed using a standardized decellularization procedure with reproducible properties and biocompatibility.

Findings

The dECM hydrogel preserved extracellular matrix proteins and effectively removed cellular components.

The hydrogel supported high cell viability and proliferation in both 2.5D and 3D in vitro models.

It is a promising platform for cardiac tissue engineering and cardiotoxicity screening.

Abstract

Cardiovascular diseases remain the leading cause of mortality worldwide, underscoring the urgent need for reliable in vitro models that recapitulate the complexity of the native myocardium. Conventional two-dimensional (2D) cultures lack structural and biochemical complexity, whereas in vivo models are costly, raise ethical concerns, and have poor translational potential. In this study, we developed a novel hydrogel scaffold derived from decellularized porcine ventricular myocardium (dECM). A newly optimized decellularization strategy effectively removed cellular and nuclear components while preserving essential extracellular matrix proteins. The dECM-based hydrogel exhibited reproducible self-crosslinking, gelation kinetics, and stability. Cytocompatibility assays using human bone marrow-derived mesenchymal stem cells demonstrated excellent viability and proliferation upon contact with the biomaterial. Multidimensional hydrogel applications (2.5D and 3D) in vitro revealed higher cell densities than those observed under 2D conditions. Moreover, using human umbilical vein endothelial cells, the dECM-based hydrogel proved to be a valid tool for fabricating cardiovascular in vitro models. As such, this cardiac dECM-based hydrogel is a structurally preserved, biocompatible platform that supports both short- and long-term cell culture. The scaffold has the potential to serve promising applications in cardiac tissue engineering, disease modeling, and cardiotoxicity screening by offering a closer mimicry of the native myocardial environment.

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1. Introduction

Cardiovascular disease is the leading cause of death worldwide [1]. It encompasses a wide range of disorders, including coronary heart disease (such as heart attack and angina), cerebrovascular disease (such as stroke), heart failure, and heart rhythm disorders. Traditionally, basic research in cardiovascular modeling relied primarily on two-dimensional (2D) in vitro and in vivo experiments. The 2D standard cell culture is easy to establish and is cost-effective [2], and the widespread use of induced pluripotent stem cells has provided an accessible source of cardiomyocytes [3]. However, advancements in cellular biology have highlighted the limitations of traditional 2D monolayer cultures in studying disease and functionality [4]. Indeed, such a 2D modeling approach cannot mimic the complex interactions between cells and the extracellular matrix (ECM) [5]. Conversely, in vivo models are expensive, time-consuming, and ethically challenging [6]. More importantly, they may not accurately replicate human cardiac disease due to physiological differences across species [7], particularly in terms of cardiac electrophysiology and pharmacological responses. Therefore, the generated outcomes may not be comparable with clinical data, risking being uninformative or not predictive for the real in-human scenario [8].

Three-dimensional (3D) disease models can accurately replicate the intricacy and environment of native tissues, providing a more relevant platform for investigating the causes of diseases, evaluating drugs, and developing therapeutic agents [9,10]. In fact, in native tissues, cell behavior is strongly influenced by the surrounding dynamic microenvironment—namely, the ECM—which transmits instructions and information through biochemical and biophysical signaling. The ECM also plays a crucial role in providing structural support to cells. Its composition and spatial distribution dictate its 3D architecture, depending on the types and concentrations of its protein components [11,12]. The myocardial tissue structure is elaborated and carefully tailored to maintain the heart’s continual pumping activity throughout the human lifespan. In particular, the myocardial ECM is composed of fibrous proteins (e.g., collagens, elastin), adhesive glycoproteins (e.g., laminin, fibronectin), and proteoglycans, forming a complex, alveolar 3D architecture that supports cardiomyocytes during contraction [13]. Fibrous proteins help prevent overstretching of the heart wall and contribute to force transmission during contraction. Specifically, collagen IV and V, along with small amounts of proteoglycans (agrin and perlecan) and glycoproteins (laminin and nidogen), form a basement membrane surrounding each cardiomyocyte. This structure is interconnected with the broader collagen network and elastin fibers [14]. Finally, other lamina components provide both rigidity and mechanical strength, and both impart compliance and elasticity to the ECM [15].

Recent trends in 3D cell culture systems—such as organoids and multicellular spheroids—and scaffold design for tissue engineering have emphasized the usefulness of biomimetic and tissue-specific materials to replicate the chemical and biological cues of the native microenvironment [11,12,16,17,18,19]. These systems involve encapsulating cells within an exogenous ECM or synthetic scaffold to preserve essential cell–cell and cell–ECM interactions. Hydrogels are useful biomaterials because of their biocompatibility, high swelling capacity, variable viscosity, mechanical qualities, and ability to imitate the ECM structure and function. They have the capacity to be critical scaffolding components that facilitate the biofabrication of sophisticated 3D models [20,21,22]. They may originate from either natural or synthetic sources and consist of 3D polymer networks, which can be thermally induced by physiological temperature, capable of a remarkable water retention, and closely mimicking the native ECM [23]. Finally, their hydrophilic structure could provide a controlled microenvironment that promotes cell adhesion, interaction, and proliferation, thereby making them suitable scaffold materials for disease modeling and drug screening [11,21,22].

Despite these insights, replicating the electrical, structural, and biochemical complexity of the natural myocardial ECM in vitro remains challenging. With the widespread use of decellularization techniques, decellularized ECM (dECM), a biochemical analog to the native tissue approach, is regarded as a promising candidate for engineering 3D cardiac constructs due to its functional and structural similarities to native tissue [24,25]. Hence, the use of scaffolds based on dECM—as nature’s own platform—has been explored for a variety of tissue engineering and disease modeling applications [26,27,28,29]. Cardiac dECM, with or without recellularization, could be applied both to fabricate cardiac tissue-like for implantology purposes and for in vitro modeling. Actually, these applications were primarily employed to induce regeneration and provide structural support in damaged or failing ventricular myocardium [30]. They could consist of a cardiac decellularized patch [30,31,32] or a bioprinted dECM scaffold [33,34,35], but more frequently, they were used as in situ hydrogels [19], which applied decellularized injectable biomaterials, both alone and as cell delivery vehicles. They have been studied primarily for in vivo applications, such as cardiac repair for the treatment of myocardial infarction [36,37]. Among them, decellularized myocardial matrix-injectable hydrogels—derived from porcine cardiac tissue—are especially promising [38].

However, the use of myocardial dECM as a scaffold might help to produce more reliable and tissue-like hydrogels that provide specific cues for cardiac cell growth and differentiation. In this article, we present detailed methods for fabricating well-characterized decellularized myocardial tissue to produce dECM hydrogel scaffolds that are suitable for cardiovascular in vitro studies. Several evaluations of this hydrogel, including crosslinking capacity, swelling, and main composition using immunofluorescence analysis, were performed. The cytocompatibility of this myocardial dECM hydrogel, as well as its potential to support long-term cell culture and multidimensional proof-of-concept applications, was explored for future cardiac modeling usage.

2. Results and Discussion

2.1. Myocardium Decellularization and dECM Assessment

Before and after decellularization, we performed a thorough tissue assessment of ECM components and non-ECM proteins. Native left ventricle myocardial samples (Figure 1a,c) displayed a high density of cardiomyocytes and other cells, which appeared to be removed entirely in their components, including nuclei, in decellularized counterparts (Figure 1b,d). This observation was confirmed by DAPI staining, imaged at constant exposure time (Figure 1e,f). Moreover, Masson trichrome staining revealed the presence of macrostructures, such as muscle bundles or blood vessels, and the main component appeared to be the heart muscle bundles in native tissue samples. In the decellularized LV, the tissue structure became amorphous, characterized by a long fibrous architecture, principally composed of collagens and devoid of non-ECM proteins.

To further investigate cell extraction, indirect immunofluorescence for common cardiomyocyte cell markers was performed. In the native tissue, cardiomyocyte presence was evident, both in terms of the cytoskeleton (F-actin, Figure 2a) and the sarcomere structure, and was revealed by several typical proteins, such as α-sarcomeric-actin, cardiac Troponin T (cTnT), and Myosin regulatory light chain 2 (MLC-2V) (Figure 2c,e,g). Also, the protein Connexin 43 (Cx43), a component of cardiac gap junctions, was observable. Instead, as shown in Figure 2, no positive signal was revealed for these cell markers in dECM samples (Figure 2b,d,f,h,j) when compared with native tissues.

In addition to previous analyses, residual DNA was quantified in decellularized samples using biochemical assays and compared with that in native tissue. DNA content in dECM samples (29.78 ± 18.17 ng/mg) was significantly lower than that of native tissues (615.99 ± 241.66 ng/mg) (Figure 3a). Moreover, this level was inferior to the threshold established by Crapo et al. [37] for effective decellularization.

Soluble collagen content was measured in dECM tissue samples using the Sircol assay kit and compared with that of native tissue. A significant reduction in soluble collagen in dECM (173.45 ± 68.01 μg/mg) was observed with respect to native tissue (409.9 ± 99.59 μg/mg) (Figure 3b).

Finally, by investigating the matrix structure of porcine myocardial native and decellularized tissue, it was observed that collagen I in native tissue (Figure 4a) was compact and distributed in a few bundles, in accordance with its mechanical–structural function. This pattern was preserved in dECM (Figure 4b). Collagen IV (Figure 4c) was evenly distributed in native tissue around all cardiac muscle fibers, forming a net; its high presence and spread were maintained even after decellularization (Figure 4d). Finally, laminin showed a pattern and diffusion similar to those of collagen IV, sometimes colocalized in both native (Figure 4e) and dECM (Figure 4f) samples.

Overall, the results indicate the successful production of myocardium-derived acellular scaffolds, in line with the primary aim of this study: the development of a novel, standardized decellularization strategy tailored for the myocardium that preserves its ECM while completely removing resident cells and their components. Indeed, compared to previous methods used to generate cardiac dECM-based hydrogels [39,40], ours was faster, had a higher capacity to remove non-ECM proteins, and did not require a bioreactor. The novel decellularization methodology was designed to induce osmotic shock, cell lysis, and removal via alternating hypo- or hypertonic solutions and detergents, carefully chosen to prevent adverse reactions to the ECM and its bioactivity. Building on previous experience with heart valves, pericardium, and the whole heart [41,42,43], the novel decellularization method avoids the use of protein inhibitors or toxic non-ionic detergents (such as Triton X-100) [44]. Moreover, the ionic detergent concentration was maintained at levels that did not induce ECM precipitation. This new myocardium-specific protocol starts with samples of controlled and standardized size to ensure uniform reagent penetration and reproducibility. Then, it consists of a series of osmotic shocks and a combination of detergent solutions, including sodium dodecyl sulfate (SDS), and Tergitol 15-S-9, in which tissues are immersed via dynamic diffusion. Unlike any procedure reported to date, all the decellularization solutions contained BDM (2,3-Butanedione monoxime), a myorelaxant, to increase reagent efficiency and reduce incubation time. De facto, this data confirmed that the newly optimized decellularization protocol effectively removed cellular and nuclear materials while preserving the structural and biochemical integrity of the myocardial ECM, a delicate network of basement membrane. The absence of DAPI and phalloidin signals across the dECM scaffolds (dECMs) confirmed the complete elimination of nuclei and cytoskeletal components. Moreover, the absence of immunofluorescence signals for the cardiomyocyte markers confirmed the procedure’s efficiency in removing non-ECM proteins without damaging ECM components, as evidenced by their distribution patterns. Of note, this thorough assessment is not a standard in the literature [24,28,39,40,44], where the success of decellularization is often evaluated solely by nuclear absence, without considering effects on ECM macrostructure. This makes our dECMs well-characterized. Moreover, quantitative DNA analysis further confirmed the suitability of this novel method for obtaining acellular dECM, revealing residual DNA levels below the established Crapo threshold of 50 ng/mg [37]. Removing cellular content is essential for generating novel scaffolds from native tissues to eliminate any nucleic acid source of inflammatory stimuli [45]. Another relevant characterization of this tissue concerns its soluble collagen content, which was significantly lower in decellularized tissues than in native tissues. In vivo, soluble collagen shows low immunogenicity and high affinity. However, its mechanical strength is significantly compromised, and it rapidly degrades [46]. Unlike the insoluble, structural collagen that forms a rigid matrix, soluble collagen is typically found in a precursor triple-helical structure state or as a smaller, less crosslinked molecule, which is not yet stable and represents only a small fraction of the total collagen [47]. Given its instability, finding it in dECM samples after a lengthy decellularization procedure should be considered a positive result, reflecting the mildness and safety of the manipulation. In the future, to complete the biochemical characterization, it may be useful to assess total collagen levels in native and dECM samples.

dECMs offer significant benefits due to their structural and compositional similarities to the native ECMs found in tissues and organs. Tissue decellularization is a valuable and successful process for producing biomimetic ECM scaffolds. Numerous studies have demonstrated that dECMs can stimulate the remodeling of various tissues in both animals and humans [48,49]. Thus, the preservation of ECM components is critical for maintaining scaffold mechanical strength and supporting cell adhesion, proliferation, and differentiation. Thanks to its thickness and composition, after careful biomechanical evaluation, the dECM scaffold might be used in dynamic recellularization with a bioreactor, similar to other cardiovascular scaffolds [48,49,50].

2.2. Gelation and Characterization of Cardiac dECM Hydrogel

A classic procedure to generate hydrogels from native tissues is their digestion with pepsin. Since pepsin digestion tends to break the protein bonds between ECM proteins, possibly altering them, the main components and the microarchitecture assumed by the crosslinked hydrogel were characterized using histological and indirect immunofluorescence. As shown in Figure 5a, the cardiac dECM-based hydrogel appeared to have an amorphous structure, although totally constituted of ECM proteins (Figure 5b). The analysis confirmed the absence of DNA residues (Figure 5c). Moreover, staining for collagen I and IV and laminin (Figure 5d–f) showed a high presence of these proteins and a complex network, although not tidy or organized. Finally, laminin appeared to be localized predominantly on one side of the hydrogel scaffold after crosslinking (Figure 5f).

The transformation of decellularized myocardial tissue into a hydrogel scaffold represents a significant step forward. Our aim was to characterize the biological properties of the main matrix elements, which are fundamental to bioactivity and cytocompatibility. Histological and immunofluorescence evaluations confirmed that the dECM-based hydrogel retained key ECM markers. In contrast, other works on hydrogel fabrication in the literature [28,51] focused on different characterizations of dECM hydrogel, in particular on evaluating the material’s mechanical properties through rheological testing.

The turbidimetric gelation kinetics of the dECM-based hydrogel, expressed as normalized absorbance, for different parameters are shown in Figure 6. First, we evaluated whether preservation might alter gelation kinetics. As shown in Figure 6a, both curves, generated from 25 mg/mL hydrogels derived from fresh and frozen materials, exhibited sigmoid behavior, typical of processes approaching saturation and compatible with dECM-based hydrogel crosslinking kinetics reported in the literature [28,51,52,53,54]. The curve of fresh material (blue) seemed to reach the plateau more quickly and at a slightly higher level, suggesting faster kinetics than for the frozen material curves (red). Indeed, the lag phase (t_lag_) and the time required to reach half the final turbidity (t_1/2_) were greater in the frozen materials gels than in the fresh ones. However, halfway through the crosslinking incubation time, both reached their maximum value (plateau) and showed a similar slope of the curve (k), reaching comparable maximum turbidity values. The frozen and fresh kinetic comparison showed that our hydrogels’ crosslinking properties were maintained after freezing and thawing.

Turbidimetric analysis was repeated at different hydrogel concentrations: 5 mg/mL, 12.5 mg/mL, and 25 mg/mL (Figure 6b). In terms of absolute absorbance, the higher the hydrogel concentration, the higher the turbidimetric absorbance [51]. After normalization, all different material concentrations showed a sigmoid shape with similar trends and kinetic times. The kinetic behavior of 25 mg/mL (green) and 12.5 mg/mL (blue) hydrogel curves was similar in terms of lag phase (t_lag_) and the time required to reach half the final turbidity (t_1/2_). Conversely, 5 mg/mL (red) hydrogels appeared to reach the plateau more quickly, although no significant differences were observed in the kinetic analysis. These results underlined that cardiac dECM-based hydrogel crosslinking capacity was independent of its concentration. Finally, the turbidimetric analysis was performed at different working temperatures to determine whether the material could undergo crosslinking at RT and exhibit similar kinetics. In both temperature conditions, the 5 mg/mL hydrogel showed a different curve shape and progression, excluding possible material crosslinking at RT (Figure 6c).

Since the dECM-based hydrogel might be used as a scaffold in cell culture, it was necessary to evaluate whether it undergoes macroscopic changes over time. Swelling analyses on cell-free 25 mg/mL dECM-based hydrogel molds revealed that in the first 10 min, there was a rapid increase in normalized weight, after which the curve ‘flattens’ and tends to a lower value, indicating the material reached a swelling plateau and remained stable. The saturation curve of the material was typical of swelling processes, with rapid development at the beginning, followed by a gradual slowing and eventual stabilization. This analysis confirmed that the cardiac dECM-based hydrogel structural components maintained integrity over time, enabling its application in long-term culture conditions.

As the last assessment, cytocompatibility of cardiac dECM-based hydrogel was evaluated with human bone marrow mesenchymal stem cells (hBM-MSCs). The first step towards assessing cytocompatibility was to examine the hBM-MSC capacity to be cultured on the cardiac dECM-based hydrogel. Following the MTS Cell Proliferation Assay, cardiac dECM-based hydrogel was observed as a valid growth substrate for hBM-MSCs (Figure 7a–c). Cells grown onto the cardiac dECM-based hydrogel (hydrogel condition) had a proliferation fold increase comparable or higher (day 1, 1022 ± 0.04; day 7, 1.16 ± 0.16; day 10, 1.24 ± 0.18) than the control in each time point.

The amount of LDH (Figure 7d–f) released in the hydrogel condition (0.9 ± 0.09) was comparable with the control condition in the first 24 h of culture. Likewise, at other time points (7–10 days), there were no differences in cell death between plastic and hydrogel, except when compared with the DMSO condition (cytotoxicity positive control).

The cytocompatibility of the cardiac dECM-based hydrogel was rigorously tested using hBM-MSCs. Cell proliferation assays over 1, 7, and 10 days revealed that the hydrogel provided a conducive microenvironment for cell survival and growth. Generally, cytocompatibility assays were limited to a few days of culture [55]. Instead, to test the ability of this scaffold to support long-term cultures, we continued the evaluation up to the tenth day, obtaining results comparable to those of the plastic control. At the end, both assays showed that cells exhibited a positive growth response when placed in contact with the hydrogel, suggestive of its cytocompatibility.

2.3. Cardiac dECM-Based Hydrogel In Vitro Applications

2.3.1. Cardiac dECM-Based Hydrogel Applied to an Endothelial In Vitro Model

We further evaluated the potential of applying a dECM-based hydrogel for cardiovascular in vitro modeling. Therefore, a cardiac dECM-based hydrogel coating was created to simulate its potential application in establishing an in vitro cardiovascular model using human umbilical cord endothelial cells (HUVECs), a widely adopted cell line for in vitro modeling. Moreover, HUVECs were exposed to valsartan and bosentan, drugs commonly used in hypertension therapy and known to target endothelial cells and affect their viability. The effects of different culture conditions on viability were compared using the MTS Cell Proliferation Assay. As shown in Figure 8a–c, cardiac dECM-based hydrogel coating significantly increased HUVEC viability at day 2 (1.27 ± 0.07), and a trend toward increased proliferation compared with the plastic control was maintained later.

LDH release was also assessed. As visible in Figure 8d–f, a negative correlation was revealed in HUVEC cultures between the coating and plastic control. Cells treated with drugs showed a continuous increase in LDH release. At day 3, this difference (2.02 ± 0.55) was significant compared with plastic and coating (1.02 ± 0.04). These results revealed that the dECM-based hydrogel increases HUVEC proliferation capacity in a manner comparable to drug treatment, but with lower LDH release.

Although the native architecture of the decellularized ECM was disrupted during hydrogel processing and treatment, its main components were preserved, providing a cytoprotective support for HUVECs. Thus, the cardiac dECM-based hydrogel coating can act as a tissue-specific biological substrate that simulates heart microvasculature scaffolding. Although not in hydrogel form, other studies used digested cardiac ECM-coated surfaces with similar biochemical cues presented to cells to promote cell culture and increase proliferation, viability, and adhesion [56,57].

2.3.2. Cardiac dECM-Based Hydrogel for a 2.5D In Vitro Model

The crosslinked cardiac dECM-based hydrogel was employed as a cell scaffold. This kind of culture substrate was defined as a 2.5D condition, and it was tested as another possible application of cardiac dECM-based hydrogel for in vitro modeling. With this aim, the hBM-MSCs were seeded and grew on the cardiac dECM-based 2.5D scaffold for 10 days. MTS Cell Proliferation Assay (Figure 9a–c) revealed the ability of cardiac dECM-based hydrogel to support cell adhesion and survival (Figure 9a). On day 1, similar proliferation was observed compared to the control (negligible fold increase (0.91 ± 0.04)). In the following time points (day 7, 0.72 ± 0.16; day 10, 0.71 ± 0.14), the cell vitality for the hBM-MSCs seeded onto the dECM hydrogel scaffold seemed to have a decreasing trend, showing a lower growth than cells in plastic and coating conditions, although not significantly.

The amount of LDH (Figure 9d) released in the 2.5D condition was significantly lower (0.23 ± 0.09) than in other conditions in the first 24 h of culture. At the following time points (7–10 days), no differences in cell death were appreciated between plastic (2D) and 2.5D, except when compared with the DMSO condition (cytotoxicity positive control).

It can be concluded that the 2.5D cardiac dECM-based hydrogel allowed cells to proliferate and maintain vitality throughout, without any cytotoxic effect, as evidenced by this comparison with the control. The unclear and contrasting results regarding viability could be explained by the MTS color assay’s methodology. In fact, in the literature, color viability assay evaluations were prevalently performed under coating conditions or for material contact [24,27,28,52], but not for direct seeding onto or into a scaffold. In line with our results, Dominijanni et al. [58] observed inaccuracies in cellular viability outputs across commonly used assays when comparing different hydrogel formulations in 2D cell culture. This may be likely due to the colored byproduct quenching by the hydrogel during the necessary technical incubation of the assay reagent with the tested cells. As empirical proof of this hypothetical explanation, all the seeded scaffolds appeared colored at the end of the reaction, as if they contained the reaction byproduct of the assay, and the medium to be tested appeared clearer. Therefore, a lower absorbance value for the 2.5D condition than that of the plastic could be related to the vitality assay solution absorption by the hydrogel.

2.3.3. 3D In Vitro Application

To evaluate whether a cardiac dECM-based hydrogel could be used as a 3D scaffold for in vitro viable tissue formation with hBM-MSCs, immunofluorescence of cells grown onto plastic, coating, and 3D cryosections was performed at each time point. Under the same cytocompatibility evaluation conditions, including cell seeding density, hBM-MSC-dECM-based hydrogel 3D constructs were generated by mixing 25 mg/mL of pre-hydrogel solution with hBM-MSCs, crosslinking in a well of a 96-well cell plate, and culturing for 10 days. Cell distribution was rated using histological analysis and digital counting. After counting, the number of cells populating the plate and the scaffold was normalized for each culture surface area, thus allowing an appropriate 3D vs. 2D comparison. As shown in Figure 10, the number of cells for cm^2^ was comparable between plastic (4538.9 ± 441.9 cells/cm^2^) and coating (5972.2 ± 985.6 cells/cm^2^), but with significantly higher values in 3D conditions (7224.4 ± 1621.2 cells/cm^2^) already after 1 day from the seeding. This gap in cell counting between 2D cultures, coating conditions, and 3D hydrogel construct increased on day 7 (plastic 22,416.2 ± 911.4 cells/cm^2^ vs. coating 19,363.3 ± 3414.3 cells/cm^2^ vs. 3D 60,951.3 ± 23,006.3 cells/cm^2^) and 10 (plastic 24,027.8 ± 3259.8 cells/cm^2^ vs. coating 22,311.1 ± 3790.0 cells/cm^2^ vs. 3D 62,726.9 ± 5822.9 cells/cm^2^), showing a clear and significant superiority of the 3D hydrogel formulation in supporting cell growth and viability.

The disparity between 2D CTR and coating condition increased over time, indicating that our cardiac hydrogel dECM-based not only allows cells to survive but also provides a compatible, permissive support, enabling cells to grow, proliferate, and undergo 3D colonization. All these findings are also crucial for further applications external to in vitro tissue modeling and cardiotoxicity evaluation. De facto, they suggest that the hydrogel not only mimics the native myocardial matrix but also supports the integration of exogenous cells, a fundamental requirement for tissue engineering and disease modeling. Together with easy availability, crosslinking temperature, and the possibility of varying the concentration of the pre-hydrogel solution to tune its properties, it might be a very promising tool for cardiac tissue engineering. In this sense, further characterization of the rheological properties is essential to explore the mechanical features. Indeed, viscoelasticity properties could influence cell differentiation [59,60], especially in the cardiac field [17,18,61]. Furthermore, given that dECM-based hydrogels have been used in several fields for 3D bioprinting, it might also be appropriate to evaluate combinations of our biomaterial with other ones to improve its biophysical properties [8,33,35,48].

Engineered tissues are currently used to generate in vitro 3D platforms for disease modeling, drug development, and screening of new therapeutic hypotheses [62,63,64]. The general concept is, on the one hand, to fabricate constructs using cells and bioscaffolds with a microenvironment and structure similar to the tissue or organ of interest, and on the other hand, to recreate stimuli and conditions that simulate the physio/pathological condition of interest. It is mandatory to rely on cardiac models that closely resemble human myocardium, one of the main tissues affected by cardiac injury and disease [64,65]. To achieve this fundamental goal, coupling cardiac dECM scaffolds with human cardiomyocytes might be a suitable, nature-inspired approach to generate in vitro clinically relevant replicas of the normal and pathological myocardium.

3. Conclusions

In conclusion, this study successfully developed a novel in vitro alternative for the fabrication of 3D-engineered cardiac tissue, i.e., a highly cytocompatible myocardium-specific dECM hydrogel scaffold. Excellent decellularization efficiency, ECM preservation, and material cytocompatibility are promising features for the future manufacture of a viable and functional myocardial tissue replica. By bridging the gap between natural tissue models and engineered scaffolds, this dECM hydrogel serves as a transformative tool for cardiac tissue engineering, disease modeling, and cardiotoxic drug screening or prediction.

The findings of this study highlight significant advancements in biomaterial availability, decellularization efficiency, and hydrogel cytocompatibility, with promising translational implications for in vitro cardiac modeling, cardiotoxicity testing, and myocardial regeneration. However, dECM materials alone can often be insufficient to suit the varying requirements of physiological or pathological microenvironments. Recently, composite materials comprising biomaterials and dECM have emerged as a solution, considerably improving the biological functions and clinical utility of dECM [33,63,66]. Therefore, in the future, to implement the model fabrication and the material properties, the cardiac dECM-based pre-hydrogel could be used to enrich other natural and/or synthetic compounds (e.g., chitosan, alginate, polylactic acid, polyethylene glycol, etc.), ECM proteins (e.g., collagen I, fibronectin, hyaluronic acid, etc.), and materials (e.g., carbon nanotubes/fiber and gold particles), or it could be used alone as a cardiac dECM bioink. By leveraging bioprinting technology, the scaffold could be tailored to different shapes and thicknesses and combined with human cardiac cells to create a faithful, reproducible, clinically relevant cardiac tissue-like model. Finally, in vivo application as an in situ cardiac hydrogel for heart healing might be envisaged, provided that stringent specifications for the certification of the material’s origin and manufacturing quality are met [24,29,44]. Its generation with non-toxic chemical reagents, the self-crosslinking ability at 37 °C, and the efficient cell homing are reassuring features for this scope.

4. Materials and Methods

4.1. Porcine Myocardium Tissue Isolation

Adult porcine hearts were collected at a local slaughterhouse (F.lli Guerriero S.r.l., Villafranca Padovana, Italy) (Figure 11a) as a discard of the food chain of market-weight pigs (Large White, ~110 kg; 6 months old). The left ventricle was isolated with a surgical steel scalpel (Swann-Morton Ltd., Sheffield, UK). Standardized 800 μm thick tissue slices (Figure 11b) with a 1 cm^2^ area were obtained using a vibratome (Leica Biosystems Richmond Inc, Richmond, IL, USA) and maintained in a physiological solution (0.9% sodium chloride (NaCl), Merck, Darmstadt, Germany) until the beginning of the decellularization process or stored at −80 °C as native tissue to be utilized as a comparison control for dECM scaffolds.

4.2. Porcine Myocardium Tissue Decellularization

Previously isolated myocardium tissue specimens were washed in sterile deionized water (dH_2_O) for 1 h (Figure 11c). Then, they were treated with hypertonic solution (0.5 M NaCl) enriched with a muscle relaxant (20 mM 2,3-Butanedione monoxime (BDM), Merck) for 2 h and 30 min. Samples were rinsed with sterile dH_2_O for 30 min and placed in an anionic detergent solution (SDS 1% (Merck), 0.5 M NaCl, and 20 mM BDM) for 40 h, with refreshment after 20 h. The myocardial specimens were then rinsed in sterile dH_2_O for 4 h, refreshed each h, and incubated with a non-ionic detergent solution (0.1% TERGITOL^™^ (Merck), in dH_2_O) overnight (12–18 h). The specimens were washed for the last time in dH2O for 4 h. All steps were performed at room temperature, and in each one, the diffusion through the samples was facilitated by magnetic stirring (M2-A ARGOlab, Milan, Italy) at 700 rpm.

An enzymatic treatment with aspecific endonucleases (Benzonase^®^ Nuclease, Merck, 45 U/mg, in 20 mM Tris (hydroxymethyl)aminomethane (Tris, pH 8, Roche Diagnostics, Basel, Switzerland; 2 mM MgCl_2_, Merck)) was also performed for 48 h at 56 °C to eliminate residual nucleic acids. An enzymatic activity refresh was carried out at 24 h. After a 1 h wash with sterile dH_2_O, the samples were stored at 4 °C in phosphate-buffered saline (PBS, Merck) until freezing.

4.3. Decellularization Assessment

4.3.1. Histological and Immunohistochemical Assessment

Before freezing, native tissue and dECM samples were treated with 4% paraformaldehyde (PFA, Merck) for 15 min at room temperature. Samples were washed in PBS and incubated at 4 °C first for 1 h in 10% sucrose solution and then overnight in 30% sucrose solution before being embedded in OCT compound (O.C.T. compound, Bio-Optica, Milan, Italy) using nitrogen vapors. Frozen samples were then sectioned at 5 μm with a cryostat (CM1850 UV, Leica Biosystems, Nussloch, Germany) and collected on microscope slides (Adhesive plus, Bio-Optica). Cryosections were maintained at −20 °C until further processing for staining.

Histological and histochemical analyses were carried out using the Hematoxylin and Eosin staining kit (Rapid Frozen Section) and the Masson trichrome staining kit (both from Bio-Optica) following the manufacturer’s instructions. All the information about immunofluorescence staining procedures and antibodies was reported in Table S1.

A Leica DMRE microscope (Leica Biosystems, Wetzlar, Germany) equipped with a Leica 541 517 HC camera was used for observation and image collection. The proprietary Leica Application Suite X_3.10.0.28982 software (Leica Microsystems) was utilized for image elaboration.

All decellularization experiments were repeated across a minimum of 4 donors, and at least 3 samples per experiment were analyzed to ensure that decellularization was achieved and that the sample pool yielded consistently.

4.3.2. Quantitative Assessment of DNA Content

After lyophilizing and weighing samples, the DNeasy Blood & Tissue Kit (Qiagen; Hilden, Germany) was used to extract residual DNA from both native and decellularized myocardial LV specimens (n = 7). All DNA extracts were quantified using Nanodrop (ND-1000 Spectrophotometer, Thermo Fisher Scientific, Milan, Italy). The final concentration, expressed in ng/mg, was obtained by normalizing this value for tissue weight and compared to the maximum threshold of 50 ng/mg dry tissue, defined by Crapo et al. for effective decellularization [30].

4.3.3. Quantitative Assessment of Collagen Content

After the freezing step in liquid nitrogen vapor for 10 min and overnight lyophilization (LIO-5P DIGITAL, 5PASCAL, Padua, Italy), cardiac dECM was pulverized using a mortar and pestle [31] (Figure 11d–f). Soluble collagen was extracted from cardiac dECM powder using 0.1 mg/mL pepsin (Merck) in 0.5 M acetic acid (Merck) overnight at 4 °C. The amount of pepsin-soluble collagen was determined in each sample using the Sircol Soluble Collagen assay (Biocolor, Ltd., Belfast, UK), according to the manufacturer’s instructions, and normalized to the weight of lyophilized cardiac dECM powder. Soluble collagen was quantified in dECM originating from 4 donors in duplicate.

4.4. dECM Hydrogel Fabrication and Characterization

4.4.1. dECM Digestion and Crosslinking

Based on the Fercana GR et al. method [32], dECM powder was digested in a solution of 0.01 M HCl containing 1 mg/mL pepsin from porcine gastric mucosa (~2000–2300 U/mg, Merck) at a final concentration of 35 mg/mL (Figure 11g). Optimal digestion was performed at pH 2 for 48 h at room temperature while stirring at 700 RPM. After pepsin inactivation (Figure 11h), the pre-hydrogel was formed by raising the pH to 7.4 with 0.1 N NaOH and 10X PBS (1/10 and 9/10 of the pre-hydrogel solution volume, respectively) at 4 °C. The pre-hydrogel solution was brought to the desired concentration of 25 mg/mL using cold (4 °C) 1X PBS, pH 7.4. It was placed at 37 °C for gelation to occur or stored at −20 °C until further use (Figure 11i).

4.4.2. Hydrogel Gelation Kinetics

Turbidimetric hydrogel gelation kinetics were determined for porcine cardiac dECM-based hydrogel (25 mg/mL). Samples were tested both when fresh and after thawing from frozen storage to assess any variation induced by cryopreservation. Hydrogel optical density readings were performed at 405 nm on 100 μL aliquots of n = 6 samples in triplicate in a 96-well culture plate (Corning, Germany), with measurements taken every 2 min for up to 1 h using a spectrophotometer (Multiskan FC, Thermo Fisher).

Normalized absorbance (NA) was determined by the following equation [32]:

$$[eqn]$$

where ‘A’ represents the absorbance reading at a particular time point, ‘A_0_’ represents the initial absorbance, and ‘A_max_’ represents the maximum absorbance. Additional metrics of ECM gelation determination include the time required for 50% gelation, defined as ‘t_½_’, and the lag phase ‘t_lag_’, determined via extrapolation of the linear portion of the normalized absorbance curve.

4.4.3. Hydrogel Swelling Analysis

Cardiac dECM-based hydrogel scaffolds were generated by pipetting 100 μL of pre-hydrogel solution at 25 mg/mL into each well of a sterile 96-well culture plate under a sterile hood. The solution was left to crosslink at 37 °C for 1 h in a cell incubator (Eppendorf CellXpert C170i, Hamburg, Germany), resulting in a transition from a highly viscous liquid to a soft solid form.

Cell-free cardiac dECM-based hydrogel scaffolds were assayed for equilibrium swelling analysis at both 12.5 mg/mL and 25 mg/mL. After polymerization, the blocks were weighed. Then, they were immersed in prewarmed 1X PBS (37 °C) and weighed at different time points (0, 1, 5, 10, 15, 30, 60, 120, 180, 1440, 2880, and 4320 min). Hydrogel swelling measurements were repeated six times per point.

4.4.4. Hydrogel Cytocompatibility

The hBM-MSC cell line, provided by PromoCell^®^ (cryopreserved, PromoCell^®^, Heidelberg, Germany), was employed to evaluate cardiac dECM-based hydrogel cytocompatibility and the feasibility of using it to develop in vitro models. The cell line was cultured on plastic tissue culture dishes (Corning) at a cell density of 5 × 10^3^ cells/cm^2^ in a cell incubator (Eppendorf CellXpert C170i) at 37 °C and 5% CO_2_ in complete culture medium (minimum essential medium eagle (MEMα), 20% fetal bovine serum (FBS), 1% v/v L-glutamine solution, 1% v/v penicillin-streptomycin (all from Merck)). The medium was changed every two days, upon 75–80% confluence, and then hBM-MSC was split with Trypsin/EDTA solution (ScienCell Research Laboratories, Faraday Ave Carlsbad, CA, USA).

Wells of a 96-well cell plate were pre-treated with cardiac dECM-based hydrogel without crosslinking it. Cytocompatibility analyses were performed for several days (1, 7, and 10 days) in duplicate four times (n = 4) for each condition and time point using vitality and cytotoxicity assays.

hBM-MSC viability was assessed through the CellTiter 96^®^ Aqueous One Solution Cell Proliferation Assay (Promega, Milan, Italy) following the manufacturer’s instructions. At the end of the reaction, absorbance at 450 nm was measured with a spectrophotometer (Multiskan FC, Thermo Fisher). Cells cultured on plastic were used as a reference, while DMSO was treated as a negative control. The proliferation index was expressed as the fold increase based on plastic culture for each time point.

Cytotoxicity reactions to the cardiac dECM-based hydrogel were investigated by measuring the amount of cytoplasmic LDH released into the medium through the TOX7 assay kit (Merck) following the manufacturer’s instructions. Post-reaction, the absorbance at 490 nm was measured with a spectrophotometer (VICTOR X, PerkinElmer, Milan, Italy). Cells cultured on standard plastic were used as a reference, while cells exposed to 0.1% DMSO were considered as a cytotoxic/positive control. The LDH release index was expressed as a fold increase.

4.5. Cardiac dECM-Based Hydrogel Scaffold Applications

4.5.1. dECM-Based Hydrogel Cardiovascular In Vitro Model

The HUVEC cell line, provided by PromoCell^®^ (cryopreserved, PromoCell^®^), was employed to evaluate the cardiac dECM-based hydrogel’s feasibility of using it to develop in vitro cardiovascular models. The cell line was cultured on plastic tissue culture dishes (Corning) (Figure 12a) at a cell density of 5 × 10^3^ cells/cm^2^ in a cell incubator (Eppendorf CellXpert C170i) at 37 °C and 5% CO_2_ in complete culture medium (EBM^®^-2 Basal Medium supplemented with 2% FBS, 4 μL/mL hFGF-B, 1 μL/mL VEGF, 1 μL/mL R3-IGF-1, 1 μL/mL Ascorbic Acid, 1 μL/mL hEGF, 1 μL/mL GA-1000, 1 μL/mL Heparin, and 0.4 μL/mL Hydrocortisone (all from Lonza, Milan, Italy)). The medium was changed every two days, upon 85–90% confluence, and the splitting was performed with Trypsin/EDTA solution (ScienCell Research Laboratories).

HUVECs were seeded onto plastic (CTR) and cardiac dECM-based hydrogel coating (coating) (Figure 12b) under culture conditions at 5 × 10^3^ cells/cm^2^. Serum starvation in fresh culture medium containing 1% FBS was performed for 24 h before drug treatment to minimize exposure to serum-derived growth factors, which can influence cellular responses to treatment. Drug stimulation was performed with valsartan (PHR1315, Merck) 10^−5^ M and bosentan (Roche, Switzerland) 10^−7^ M [33] and added to cell cultures simultaneously (TR and coating + TR). Cell conditioning was carried out for 24, 48, and 72 h and tested for vitality and cytotoxic effect.

4.5.2. dECM-Based Hydrogel 2.5D Culture

Cardiac dECM-based hydrogel at 25 mg/mL was pipetted and crosslinked in wells of a 96-well cell plate to fabricate 2.5D scaffolds (Figure 12c). Afterwards, three washes were performed with sterile 1X PBS. The cell culture medium was added for 1 h at 37 °C to equilibrate the scaffold until further cell seeding. Cardiac dECM-based hydrogel scaffolds were seeded using the drop technique at the same density as the plastic (2D) control. Cell attachment to the hydrogel surface was allowed at 37 °C for 1 h. Afterward, the culture medium was added to each well to reach the final desired volume. Cell culture was carried out for 1, 7, and 10 days and tested for vitality and cytotoxic effect.

4.5.3. In Vitro 3D Tissue-Like Construct

To realize a 3D construct, cardiac dECM-based pre-hydrogel was mixed with 5 × 106 hBM-MSCs/mL (Figure 12d). A volume of 100 μL of the mixture was seeded for each well of a 96-well cell plate and incubated for 1 h at 37 °C to crosslink. Then, the constructs were moved to a 12-well cell plate for the long culture and stopped at days 1, 7, and 10 for histological analysis. Cells cultured on plastic and 2.5D coating were used as a culture control.

Cell Vitality and Proliferation

Cells grown in a 96-well cell plate (plastic and coating) and a cellularized cardiac dECM-based hydrogel 3D scaffold condition were fixed with 4% PFA (Merk) for 15 min at room temperature for the following cell counting examinations. Samples were washed three times with 1X PBS. The same immunofluorescence procedure applied for dECM analysis, previously described in the decellularization assessment chapter, was also used for these samples, which were stained with phalloidin-Atto 488 (Merck) and DAPI-containing medium (FluoroshieldTM with DAPI, Merck) for nuclear counterstain. Anti-collagen IV immunofluorescence was performed on cryosections of cellularized hydrogels for cardiac dECM visualization and area normalization. The Leica 541 517 HC camera and Leica Application Suite X software were used to acquire images. ImageJ software v1.53t was used to elaborate images and cell counting. For each condition and time point, n = 5 samples and n = 5 images were analyzed. The total number of cells was normalized to the growth area and expressed as cells per cm^2^.

4.6. Statistics

All statistical analyses and graphs were carried out utilizing GraphPad Prism 8.0.2 software (Dotmatics Luma, Boston, MA, USA).

Statistical analysis of quantitative assessment of DNA content was carried out utilizing an unpaired t-test, and a p < 0.05 was considered statistically significant. Statistical analysis of the quantitative assessment of collagen content was performed using an unpaired t-test, and p < 0.05 was considered statistically significant. The sigmoidal graphs of hydrogel gelation kinetics were analyzed using Mann–Whitney t-tests, and statistical significance was determined using the Holm–Sidak method with alpha = 0.05. Hydrogel swelling normalized weight was graphed and analyzed with the Friedman one-way ANOVA test, and statistical significance was determined using Dunn’s method, with alpha = 0.05.

The cytocompatibility results were normalized to the control condition for each time point, put in a graph together, and compared with one-way ANOVA. Multiple comparisons within each row were performed using Bonferroni’s method, and p < 0.05 was considered statistically significant.

The vitality and cytotoxicity results of cardiovascular and 2.5D models were normalized to the control condition for each time point, graphed together, and compared with one-way ANOVA. Multiple comparisons within each row were performed using Bonferroni’s method, and p < 0.05 was considered statistically significant.

Cell proliferation data of an in vitro 3D tissue-like construct was compared using one-way ANOVA. Multiple comparisons within each row were performed using Bonferroni’s method, and p < 0.05 was considered statistically significant.