Comparable Decellularization of Fetal and Adult Cardiac Tissue Explants as 3D-like Platforms for In Vitro Studies

This method can help address important questions in the cardiovascular field about the impact the extracellular matrix has during different physiological and developmental stages of the heart. The main advantage of this technique is that it allows a comparative analysis between fetal and adult extracellular matrices by applying a similar decellularization method to both types of tissue. So this method have been successful applied to other organs such as surgical resections from cancer patients facilitating the study of the extracellular matrix in different contexts.

Begin by briefly rinsing the fetal and adult mouse hearts with ice-cold PBS. Then place the tissues under a dissecting microscope and use straight small scissors to remove the atria, right ventricle, and the septa from the adult mouse hearts. Transfer the left ventricle to a new Petri dish and divide the left ventricle free wall into two millimeter thick longitudinal strips.

The use of adult mouse left ventricle explants of a homogeneous size is essential for a consistent decellularization efficiency. Use the scalpel and serrated tweezers with curved tip to remove the papillary muscle before using a two by two millimeter grid to further mince the strips into smaller tissue fragments. When all of the tissue has been minced, briefly rinse the pieces with enough PBS to cover the explants.

And transfer the fetal hearts and pieces of adult heart tissue into individual 1.5 milliliter microcentrifuge tubes containing 250 microliters of optimal cutting temperature or OCT compound per tube. Then, freeze the cardiac tissue samples in dry ice cooled isopentane for storage at negative 80 degrees Celsius. To decellularize the cryo-preserved tissues place the frozen samples in a Petri dish containing enough PBS to completely cover the explants.

After the OCT has melted, submerge the samples in a new Petri dish of PBS and wash the samples three times on a shaker for 10 to 15 minutes at 60 RPM per wash. After the last wash, add one milliliter of working hypotonic buffer to each well of a sterile 24-well tissue culture plate and use fine forceps to transfer one fragment to each well. Incubate the plate at 25 degrees Celsius for 18 hours, at 165 RPM followed by three one-hour washes in one milliliter of PBS per well, per wash.

After the last wash, add one milliliter of freshly prepared sodium dodecyl sulfate or SDS solution to each well for a 24-hour incubation at 25 degrees Celsius. The next day, rinse the samples with three 20 minute washes in one milliliter of hypotonic wash buffer per wash followed by the addition of one milliliter of DNase treatment solution to each well for a three-hour incubation, at 37 degrees Celsius. At the end of the incubation, the decellularized samples should lose their gelatin-like consistency.

Then wash the tissue fragments three times with one milliliter of PBS for 20 minutes per wash followed by an overnight wash in one milliliter of fresh PBS per well at room temperature and 60 RPM. To assess the removal of cellular remnants from the decellularized tissue, the next morning fix the samples in one milliliter of freshly prepared 10%formula neutral buffer supplemented with 0.03%of aqueous eosin per well for 2.5 to three hours at room temperature. At the end of the incubation, wash the samples in one milliliter of PBS per well.

And use a disposable vinyl specimen mold to encapsulate the decellularized fragments in histology processing gel according to the manufacturer’s specifications. Next, transfer the encapsulated fragments to a biopsy processing embedding cassette. And process the samples for paraffin embedding through successive 30-minute incubations in ascending concentrations of ethanol, isoparaffinic aliphatic hydrocarbon solution and two 56 degrees Celsius paraffin emergens.

After the second paraffin emergent, mount samples in a paraffin block and use a microtome to obtain three-micrometer thick sections. Then verify the decellularization efficiency by staining the sections with haematoxylin and eosin and Masson’s trichrome stains according to standard protocols. After the overnight PBS wash with shaking, add 500 microliters of freshly prepared DPBS, supplementd with 2.5 micrograms per milliliter of amphotericin B 1%antibiotic solution to each scaffold.

And store the samples for one to seven days at four degrees Celsius. Before seating the cells, replace the DPBS solution with 500 microliters of cell basal medium supplemented with antibiotics. And incubate the scaffolds for one hour at 37 degrees Celsius.

At the end of the incubation, use a microscope to add a four microliter drop of DPBS to the center of one well of a 96-well plate per scaffold. And use thin straight tweezers to transfer one decellularized scaffold to the top of each drop. Remove any extra DPBS by aspiration and confirm that the scaffold is not folded or wrinkled.

When the scaffolds have dried at the edges, slowly seat a single cell suspension of the cells of interest on the top of each scaffold. Embryonic day 18 decellularized tissues are characterized by a highly translucent structure while adult explants exhibit a translucent to white appearance. Hematoxylin and eosin and Masson’s trichome stains confirm an efficient cell removal by the presence porous mesh.

The nuclear material is reduced by approximately 99.8%after decellularization, compared to non-manipulated tissues which is essential for avoiding triggering an undesired inflammatory response upon implantation. The cell viability within seated scaffolds can be monitored during in-vitro culture by calcein staining. Terminal analysis of the cell repopulation and distribution across scaffolds, can also be performed after paraffin processing as observed in these representative images of a central bioscaffold section.

So this technique paved the way for researchers in the exploration of the bioactive properties of the extracellular matrix from fetal and adult mouse hearts but also of other tissues.