2.2. hCPC Viability, Differentiation, and Proliferation within Bioprinted Cardiac Patches

Evaluating the viability of cells within the cardiac patches is critical to ensure live cells that can participate in producing important pro-reparative paracrine factors.^[12,15] Evaluation of cell viability directly is also critical within bioprinted scaffolds, particularly because bioprinting has been shown to reduce cell viability in printed constructs due to high shear stresses on the cells from small diameter needle tips, such as the tips used in this study.^[42,43] In addition, cells grown in thick 3D structures can suffer death due to lack of nutrient diffusion, particularly at the center of the structures, producing a necrotic core.^[42,45] As shown in Figure 5, hCPCs within printed cardiac patches were stained to determine the total number of dead (read) to live (green) cells for either GelMA (Figure 5A) or GelMA-cECM (Figure 5B). Cell viability was quantified by measuring the number of live and dead cells at different locations and heights within the cardiac patch at days 1, 3, and 6 after formation and showed high viability, from 70–80% live cells on average seen in Figure 5C. There was no significant difference between groups or time points when comparing the percent of viable cells. Throughout all structures, there was no necrotic core or reduction of cell viability, indicating that nutrient diffusion was likely not a factor. The cell viability overall was most likely not impacted significantly by the printing methodology, or if there were effects to the cells due to the printing, the degree of cell damage was mitigated by the material being an effective environment for cell growth and nutrient diffusion coupled with printing of aligned fibers which may be beneficial to cell function.

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Figure 5.

hCPC functionality within printed patches. A) Characteristic live/dead fluorescence image of hCPCs in GelMA patches, with live cells marked green (Calcien AM) and dead cells marked red (EtD) at 1 day after formation. B) Characteristic live/dead fluorescence image of hCPCs in GelMA-cECM patches at 1 day after formation. C) Viability of hCPCs in printed patches at 1, 3, and 6 days. D) Proliferation of hCPCs in printed patches at 1 and 7 days, where absorbance intensity is normalized to the measured absorbance of hCPCs in GelMA patches in all experiments. E) Fold change gene expression over hCPCs in GelMA patches for Cx43, GATA4, MEF2C, MYH7, VE-Cad, CD31, FLT-1, and ACTA-2 at day 3. F) Fold change gene expression over hCPCs in GelMA patches for Cx43, GATA4, MEF2C, MYH7, VE-Cad, CD31, FLT-1, and ACTA-2 at day 7. * = p-value < 0.05, ** = p-value < 0.005, given by ANOVA with Tukey’s post-test, n = 3–6 for all samples at all timepoints.

Proliferation and differentiation are additional parameters that are important in characterizing functionality of hCPCs in bioprinted patches. hCPC-laden patches were grown in culture media supplemented with EdU for 3 and 7 days, and absorbance intensity from GelMA-cECM patches was normalized to values measured from GelMA patches. While there was no difference between the proliferation of hCPCs in GelMA and GelMA-cECM patches after 3 days, as seen in Figure 5D, hCPCs in GelMA-cECM patches had reduced proliferation compared to GelMA patches after 7 days. Similarly, hCPC-laden patches were grown for 3 and 7 days, and the fold change in genetic expression of key cardiac, endothelial, and smooth muscle genes from hCPCs in GelMA-cECM patches compared to hCPCs in GelMA patches was assessed through polymerase chain reaction (PCR). Analysis of gene expression of cardiac transcription factors GATA4 and myocyte enhancement factor 2C (MEF2C) and cardiac-specific proteins connexin 43 (Cx43) and β-myosin heavy chain (MYH7), endothelial cell markers vascular endothelial cadherin (VE-Cad), platelet endothelial cell adhesion molecule (CD31), and vascular endothelial growth factor receptor 1 (FLT-1), and smooth muscle marker α-smooth muscle actin (ACTA-2) was evaluated at 3 and 7 days, as seen in Figures 5E and 5F. hCPCs in GelMA-cECM patches showed enhanced cardiac differentiation through increased expression of MEF2C, Cx43, and MYH7, and decreased expression of GATA4, an early differentiation marker, indicating that the hCPCs in GelMA-cECM patches were moving towards later differentiation then hCPCs in GelMA patches. hCPCs in GelMA-cECM patches also showed increased expression of endothelial marker CD31 at day 3, although there was no difference in expression of endothelial markers VE-Cad and FLT-1 or smooth muscle marker ACTA-2. At day 7, expression of all cardiac and endothelial markers was increased in GelMA-cECM patches, with higher fold-change values than day 3. ACTA-2 remained unchanged between groups at both days. Thus, we conclude that incorporation of cECM into patches improved both cardiac and endothelial differentiation of hCPCs, while not influencing smooth muscle differentiation. The enhanced differentiation of hCPCs in cECM incorporated patches at day 7 mirrors the proliferation trends seen in Figure 5D, as stem cells most often show reduced proliferation with increased committed. These assessments also reaffirm the results measured for hCPCs in 2D culture, where cECM improved differentiation of CPCs compared to cells grown on collagen-based materials.^[16] Regardless, it is clear that while hCPCs remained viable in printed patches, the inclusion of cECM improved differentiation and reduced proliferation of hCPCs, which in turn may improve paracrine potential of hCPC-laden GelMA-cECM patches.

2.3. Pro-Angiogenic Potential of hCPC/cECM Cardiac Patches

Many studies now attribute the true benefit of cell therapy to be release of paracrine factors.^[15,46–50] To evaluate paracrine release, we grew cell-laden patches in treatment media for up to 7 days and collected the conditioned media every 2 days. We then performed a tube formation assay using human umbilical vein endothelial cells (HUVECs) cultured on Matrigel with conditioned media. HUVECs grown in either non-conditioned treatment media or endothelial cell growth media with supplemented growth factors showed similar values for total tube length formed, so non-conditioned treatment media was used as positive controls. As shown in Figure 6, HUVECs formed tube-like structures when cultured in conditioned media taken from both cell-laden GelMA (Figure 6A) and GelMA-cECM (Figure 6B) patches. When comparing the angiogenic potential of cell-free patches, seen in Figure 6C, there was no difference between GelMA and GelMA-cECM groups. In contrast, the angiogenic potential of media collected from cell-laden GelMA-cECM patches was significantly higher than media from GelMA patches alone at day 3, while both groups showed improved angiogenic potential at day 7 compared to day 3. While GelMA-cECM was superior at both time points, both groups showed an increase in angiogenic potential over time. While there are many other parameters that conditioned media may alter, angiogenesis may be one of the most important for improving cardiac function. Additionally, we may be underestimating the effects as some growth factors released may interact with the GelMA and/or cECM and prevent release in to the conditioned media, as studies have shown that growth factors, such as heparin binding growth factor and hepatocyte growth factor, bind to cECM and are released gradually.^[51,52]

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Figure 6.

Angiogenic potential of cardiac patches. Characteristic HUVEC tube formation after 6 hours when grown with conditioned media collected at day 7 from cell-laden A) GelMA and B) GelMA-cECM patches. Total HUVEC tube length normalized to positive controls for C) cell-free and D) cell-laden patches. * = p-value < 0.05, given by ANOVA with Tukey’s post-test, n = 3-6 for all samples at all timepoints.

In developing effective hydrogels for soluble factor release, it is key to generate materials that are stiff enough to allow for scaffold stability, but do not have too dense a network that limits cellular functions and release of signaling factors.^[44] Systems that balance these parameters have been developed around natural or synthetic materials, many of which prove effective in releasing reparative factors into damaged tissues, whether through encapsulation of regenerative cells or the factors themselves.^[46–49] The hCPC-laden GelMA-cECM patches developed here also allowed for generation of a solid patch with enhanced factor release. These results are similar to other materials that have been developed, but with incorporation of cardiac specific cells and matrix that may release cardiac-specific paracrine factors. This cardiac specificity in the patch design may be most beneficial in repairing the damaged myocardium, as opposed to using non-tissue specific biomaterials such as GelMA.^[19,46,50]

2.4. Mechanical Characterization of Bioprinted Patches

Mechanical properties of biomaterials play a critical role in modulating cellular function. Stem cells are more viable, proliferative, and produce more effective regenerative outcomes when grown in hydrogels that match the properties of native tissue.^[12,53] This is also true as well for hCPCs, where cells perform more effectively when grown in materials that match the mechanical modulus of native myocardium from 5–15 kPa.^[54] A material modulus that more closely matches the myocardium also ensures there is limited mechanical mismatch between the hydrogel and the heart, which can otherwise cause problems such as dissection, buckling, or immune responses.^[29,55] While GelMA and GelMA-cECM patches are evaluated in this study, we also evaluated the use of modifying the mechanical properties of the patches by adding acrylate groups using N-succinimidyl acrylate, as employed in previous studies.^[33] While the modification increased stiffness over GelMA and GelMA-cECM groups, the patches more readily degraded compared to both groups and did not alter the hCPC viability or paracrine function over GelMA-cECM (data not shown), so this direction was not pursued further, although the properties of the patch could potentially be modified through this method.

As seen in Figure 7A, the modulus of pure GelMA was 3000 Pa, similar to published studies, though short of the native myocardium.^[56] Incorporation of cECM significantly increased the modulus to 5000 Pa, indicating that the material properties of the GelMA-cECM patch could be tailored within physiological ranges. In addition to stiffness, we also measured swelling ratio in Figure 7B. All samples were sufficiently hydrated, with a swelling ratio between 9 – 12. There was a decrease in the swelling ratio between the GelMA and GelMA-cECM groups, which was expected as increases in stiffness suggest a tighter polymer network and result in more liquid exclusion.

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Figure 7.

Material analysis of printed patches. A) Viscoelastic storage moduli of GelMA and GelMA-cECM. B) Swelling ratio of GelMA and GelMA-cECM patches. C) Degradation of patches, measured as the sample weight compared to initial weight of the patches post-swelling. D) Degradation of cell-free materials in cell culture media, measured as the sample storage modulus compared to the initial modulus of the material post-swelling. E) Remodeling of hCPC-laden materials grown in cFB conditioned media, measured as the sample storage modulus compared to the initial modulus of the material post-swelling. * = p-value < 0.05, ** = p-value < 0.01, *** = p-value < 0.005, given by ANOVA with Tukey’s post-test, n = 3 for all samples in all subfigures.

As a cardiac patch must persist for the repair process, degradation time is a critical parameter. We evaluated the degradation of patches and materials in cell treatment media over 21 days by examining both the change in wet weight (Figure 7C) and change in stiffness (Figure 7D). When comparing weight change of printed patches or the change in stiffness of the materials, both groups remained solid and did not significantly degrade over the range of 21 days. Although the data presented describes the degradation of patches without cells, patches that incorporated cells lasted the duration of testing shown in sections 2.3 and 2.4 with no observable degradation. It is important to describe this method of degradation as only evaluating hydrolysis, as opposed to exposing the patch to conditions seen in vivo including proteases, cyclic strain, and shear stresses. To evaluate the degradation of the patches in a more physiological relevant environment, hCPC-laden materials were cultured in conditioned media harvested from cardiac fibroblasts (cFBs), which would be present in cases of ventricular remodeling and hypertrophy. The cFB conditioned media more closely captures the environment of patches in vivo due to incorporation of a complex mixture of remodeling components, compared to commonly employed incubation in collagenase I, which degrades the patches in a matter of hours and may not be as physiologically relevant. As seen in Figure 7E, hCPC-laden GelMA materials did not degrade or change mechanical modulus over the course of 7 and 21 days in cFB conditioned media. Interestingly, while hCPC-laden GelMA-cECM materials did not degrade or change mechanical modulus over the course of 7 days, by 21 days the material stiffness increased compared to the stiffness at both 1 and 7 days. The change in stiffness at 21 days is also significantly higher than hCPC-laden GelMA materials at the same time and may be due to stimulation of the hCPCs to remodel their environment. Regardless, the GelMA-cECM patches, both with and without hCPCs, do not degrade in vitro over an extended timeframe and may be suitable for extended retention in vivo.

Neonatal Human Cardiac Progenitor Cell Isolation and Culture –

The Institutional Review Board at Children’s Healthcare of Atlanta and Emory University approved the harvesting of human neonatal c-kit expressing cardiac progenitor cells (hCPCs) from atrial appendage, as previously described.^[17,58] In short, right atrial appendage tissue was obtained from pediatric patients aged 1 week or less undergoing heart surgeries due to congenital heart diseases. The atrial appendage tissue was transported using Krebs-Ringer solution, washed with HBSS, and broken down into small sections. The tissue was then enzymatically degraded using 1 mg/mL of collagenase type II at 37°C, 5% CO₂ for 30 minutes and passed through a 70 µm filter. The mixture was centrifuged at 1000g for 5 minutes to pellet the cells. The cells were combined with magnetic beads conjugated with anti-c-kit antibody, allowed to incubate for 2 hours at 37°C, followed by magnetic sorting and successive washes with cell culture media. Separated c-kit+ cells were expanded and expression of c-kit in the cell population was measured by flow cytometry to ensure they were at least 90% positive. Cells from three donors were pooled at first passage and used in all experiments described in this research. hCPCs were grown in T-75 cell culture treated dishes with culture media for expansion. Media was changed every 2–3 days until bioink preparation. Cell culture media consisted of Ham’s F-12 media supplemented with 1x Pen-Strep, 1% L-glutamine, 10% FBS, and 10 ng/mL bFGF.

Cardiac Extracellular Matrix Isolation and Characterization –

Decellularized porcine ventricular extracellular matrix (cECM) was generated and processed as previously described.^[17,16,33] Briefly, porcine ventricular tissue was separated, sectioned into small pieces, rinsed in PBS, and decellularized using a 1% solution of SDS for 4–5 days. The decellularized cECM was rinsed with water, frozen at −80°C overnight, lyophilized, and milled into a fine powder. Then, the cECM was processed into liquid form by partial digestion with pepsin (1 mg/mL) in 0.1 M HCl for two days, at a ratio of 10:1 of cECM to pepsin. The cECM was then raised to basic pH by adding 1 M NaOH and salt concentration of 1x PBS, followed by adjustment to pH of 7.4 using HCl and NaOH and diluted to a solution concentration of 8 mg/mL. The solution was aliquoted, immediately frozen at −80°C overnight, lyophilized for 24 hours, and stored at −80°C prior to use.

Rat Cardiac Fibroblast Isolation and Conditioned Media Harvesting –

Primary cardiac fibroblasts (cFB) were isolated as previously described.^[59] In short, hearts were excised from adult male rats, shredded, and subjected to digestion with 1 mg/mL trypsin in HBSS at 4°C for 6 hours. The solutions were digested with 0.8 mg/mL collagenase in HBSS at 37°C for 15 minutes and then quenched with cell culture media. The cell suspensions were passed through 100 µm filters, followed by cell pelleting and plating for 3 hours to allow for cFB adherence, and then washed to remove non-cFBs. cFBs were grown in fibroblast growth media, which is composed of DMEM supplemented with 1x Pen-Strep, 1% L-glutamine, and 10% FBS. cFB conditioned media was collected every two days from cFBs while they were grown until confluence, at which point the media across the entire culture time was combined and homogenized.

Bioink Preparation –

GelMA solutions of 14.432% were created by dissolving lyophilized GelMA in 15 mM HEPES buffer and allowed to dissolve under stirring at 60°C for 1–2 hours. GelMA solutions were frozen at −20°C until use. Stock solutions of 13 mg/mL cECM are formed by rehydrating the cECM material with HEPES buffer followed by adjusting the solution to pH 7.4 with HCl immediately prior to use. All bioinks were prepared immediately before printing. GelMA bioinks were prepared by mixing GelMA (final concentration 5% w/v), Eosin Y (100 µM), NVP (0.75% v/v), TEOA (3% v/v), and HEPES buffer (15 mM). In the case of GelMA-cECM bioinks, the HEPES in the above formulation was replaced by the cECM (final concentration 8 mg/mL) solutions. For the inclusion of cells, neonatal hCPCs (passage 6–10) were removed from cell culture plates, pelleted at 1000 rpm for 5 minutes, and mixed with 1 mL solution of the bioink solution, producing a final concentration of 3 million cells/mL.

Bioprinting and Patch Formation –

All bioink solutions, with or without cells, underwent similar printing protocols. 1 mL of bioink was deposited into sterilized 30 cc printer barrels and pushed towards the barrel head with a sterile loose fit plunger, removing any air bubbles that formed. A sterile 27-guage plastic needle tip was added to the barrel, and a cap connecting the print head to the barrel as added. The barrels were put in the low-temperature head of the bioprinter (EnvisionTEC 3D-bioplotter Developer Series), which was set to 10°C, and the bioink was allowed to polymerize for 10 minutes. After initial gelation, the printer head was calibrated and purged at 1.2 bar for 1–3 seconds to ensure free flowing and uniform filaments. Patches and grids were printed onto a glass slide platform at room temperature, using a pressure 0.7–0.8 bar and speed of 10 mm/s. Patches were 10 mm in diameter and 0.6 um thick, which printed in 3 layers, with an infill pattern of 90° grids with 0.5 mm spacing. Six patches were printed at once. Test grids were 10mm x 10mm boxes with an infill pattern of 90° grids with 1 mm spacing. CAD models of the patches and grids printed were generated using SOLIDWORKS and imported to the printing control system through the Bioplotter RP program. Following printing, both patches and test grids were exposed to white light (Braintree Scientific) at 4°C for 5 minutes to allow for radical polymerization. The patches were removed from glass slides and put in 2 mL solutions of cell culture media in 24-well plates. Patches that were being evaluated for paracrine release were put in 2 mL solutions of treatment media in 24-well plates. The treatment media consisted of Hams F-12 media supplemented with 1x Pen-Strep, 1% L-glutamine, 1x ITS, and 10 ng/mL bFGF. All patches were kept in cell culture incubators during further experiments, which allowed for cECM polymerization within 1 hour after GelMA radical polymerization. Media was changed and/or harvested every 2 days for each group. The test grids followed a similar protocol after polymerization, with the difference being that the grids were not removed from the slides and media was added directly over the grids to allow for full and complete coverage. The test grids were left in the cell culture incubator for 1 hour to allow for the cECM to fully polymerize, washed several times, and removed immediately for imaging.

Imaging Printed Test Grids –

Imaging of the printed test grids was performed at 10x magnification with an Olympus 1X71 Inverted Microscope. Bright-field images of both GelMA and GelMA-cECM grids were taken for printability comparison. For evaluating cECM homogeneity throughout the printed structures, cECM solutions were allowed to bind with AF568 at 4°C for 1 hour, which forms a strong bond to primary amines, at a concentration of 13.3 ug/mL based on modifying a previously described protocol.^[60] The cECM solution was used to form test grids as described in the printing section, followed by swelling and incubation for at least 1 hour at 37°C. Stained GelMA-cECM test grids were imaged at 10x magnification both on an Olympus 1X71 Inverted Microscope and Olympus FV1000 Confocal Microscope. Printed test grids with hCPCs were also imaged at 10x magnification with an Olympus 1X71 Inverted Microscope. To image only cells, hCPCs were incubated with the lyophilic dye DiD according to manufacture protocol. Briefly, hCPCs were tyrpsinized from cell culture dishes, counted, and pelleted. The cells were suspended at a density of 1 million cells/mL in serum-free culture media supplemented with 5 uL/mL of DiD solution (1 mg/mL stock) and mixed. hCPCs were allowed to incubate for 20 minutes at 37°C. The cells were centrifuged at 1000 rpm for 5 minutes, the supernatant was removed, and the cells were resuspended in serum free media. The wash procedure was repeated twice to remove any unbound DiD, and the cells were resuspended in the bioink solution for printing as described above. Printed test grids were again imaged at 10x on the fluorescence microscope. For image analysis of cell homogeneity throughout the printed structure, ImageJ was used to measure several line scans of fluorescence intensity along grid lines, which were then averaged to produce the Figure.

Printability Analysis –

The printability analysis implemented in this work looks at the effectiveness of the extruded filaments in the test grids to form square holes between filaments, as previously described.^[41] 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. To this end, and as previously derived and defined, printability is given as Equation 1

Where L is perimeter and A is area of a shape. A printability of 1 is equal to a perfect square, and indicates optimal gelation, and thus printing, conditions of a bioink. Bright field images of test grids were evaluated by measuring the perimeter and area of several holes in each sample and Pr was calculated using Equation 1, with 3 technical replicates and 4–6 holes per n.

Rheological Analysis –

As the printed patches were too thick to be measured on a rheometer without rupture, disk shaped hydrogels without cells were made by sandwiching 15 uL of sample solution between two glass slides separated by a thin spacer, allowed to gel at 4°C for 10 minutes, and polymerized by white light (Braintree Scientific) for 5 minutes at 4°C. The sample disks were incubated overnight in cell treatment media to undergo cECM polymerization and swelling. The storage and loss moduli of the disks were measured using dynamic oscillatory strain and frequency sweeps performed on an Anton Paar MCR 302 stress-controlled rheometer with a 9-mm diameter 2° measuring cone.^[61] The disks were loaded in the rheometer and the system was lowered to a 39 μm gap. Strain amplitude sweeps were performed at ω = 10 rad/s to determine the linear viscoelastic range of the samples. Oscillatory frequency sweeps between 0.5−30 rad/s and 2% strain were then used to measure the storage and loss moduli. Samples were measured at 1, 4, 7, 12, and 21 days for degradation analysis of cell-free samples grown in treatment media and at 1, 7, and 21 days for degradation analysis of cell-laden samples grown in cFB conditioned media. All samples had 3–6 technical replicates per n.

Swelling –

Patches were printed without cells as described above and allowed to swell for 24 hours in treatment media. The swollen patches were weighed (wet weight) and put in separate centrifuge tubes. The samples were lyophilized in a Labconvo lyophilizer for 2 days and the weight of the dried material was measured (dry weight). All samples had 3 technical replicates per n. Swelling ratio was calculated as wet weight/dry weight.

Degradation –

Patches were printed as described above and allowed to swell for at least 24 hours in treatment media. For measuring of degradation via weight change, the patches were weighed at days 1, 3, 7, 10, 15, and 21 after formation, and degradation via hydrolysis was determined as change in weight compared to original weight at day 1. For measurement of degradation via mechanics, as described in the Rheological Analysis section, mechanical measurements of the materials were taken at 1, 4, 7, 12, and 21 days for cell-free samples grown in treatment media, and at 1, 7, and 21 days for cell-laden samples grown in cFB conditioned media. Degradation was determined as the change in storage modulus (measured at 1.61 rad/s) compared to original modulus at day 1. All samples had 3–6 technical replicates per n.

Viability Analysis –

hCPC containing patches were grown in cell culture media for 1, 3, or 6 days, changing media every 2 days. Patches were removed from growth plates and placed in a 250 uL solution of 3 uL/mL Calcein AM (live) and 2 uL/mL EtD (dead) in HBSS in 48 well plates. The patches were left for 30 minutes at 37°C to incubate, followed by two washed with 1x HBSS for 5 minutes each. The patches were removed and placed on glass bottom dishes for imaging on an Olympus FV1000 Confocal Microscope. Live/dead images of the hCPCs within the patches were taken at several locations. Several areas of each patch, six patches each, were used as technical replicates to evaluate data expressed as live cells/total cells.

Tube Formation Assay –

Conditioned media from empty or hCPC containing patches were grown in treatment media was collected at days 3 and 7. The conditioned media was centrifuged at 10000g for 10 minutes to remove any cell debris or particulate matter, and the supernatant was stored at −80°C until analysis. HUVECs were grown on 0.1% w/v gelatin-coated T-75 tissue culture plates with endothelial cell growth media until assays were performed. Tube formation assays were implemented as previously described.^[59,61] In short, HUVECs were removed from culture using trypsin and added to Matrigel coated well plates at a concentration of 10000 cells/well. Conditioned media harvested from patches (200 µL) was added to the top of each well. For positive controls, non-conditioned treatment media was added. All HUVECs were allowed to grow for 6 hours. Calcein AM dye (2 mg/mL) was added to each well and cells were imaged via fluorescence to measure tube formation. Extent of total tube length formed in each well was evaluated by the Angiogenesis Analyzer for ImageJ (Gilles Carpentier), and the total tube length for each sample was normalized to the value of the positive controls.

RNA Isolation, Reverse Transcription, and Quantitative Real Time PCR –

Cell-laden patches were grown in culture media for 3 and 7 days. At each timepoint, 3 technical replicates/n were harvested, added to vials containing 1 mL Trizol to isolate RNA, and homogenized (Fisher Scientific PowerGen 500) for several minutes.^[61] The homogenized suspension was centrifuged at 15000g for 1 minute with QIAshredder filters to separate the cellular components from the gel. RNA extraction was performed according to manufacturer’s protocols. RNA quantification and purity were determined by measuring absorbance at 260 and 280 nm wavelength on a spectrophotometer (Thermo Scientific NanoDrop One), followed by running reverse transcription as previously described.^[16,17,61] Briefly, 0.5–2 µg RNA was mixed with hexamers, oligo(dT), dNTP, and RNase-free water in a final volume of 12 µL, and samples were heated to 65°C for 5 minutes to denature the RNA, followed by cooling to 25°C for 10 minutes to allow for components to anneal. Then, RNaseOUT inhibitor, M-MLV, first strand buffer, and dithiothreitol were added to solutions, heated to 37°C for 60 minutes to undergo reverse transcription, and 70°C for 15 minutes for enzyme inactivation. cDNA samples were stored at −80°C prior to further measurement. Gene expression was measured using a quantitative real time PCR system (Applied Biosystems, StepOne Plus Software). cDNA in 1:5 ratio was mixed with Power SYBER Green, RNase-free water, and target primer, heated to 95°C for 10 minutes, and allowed to run for 40 cycles, as previously described.^[16,17] Each sample was run in triplicate per primer, and ΔΔC_t method was used to obtain fold change values over GAPDH and GelMA control.^[59] The primer sequences used are seen in Table 1.

Proliferation –

Cell-laden patches were grown in culture media supplemented with 20 µM EdU for 3 and 7 days. At each timepoint, patches were harvested and cut into equal size sections to fit in 96 well plates. Click-iT EdU assay was performed according to manufacturer instructions. Briefly, cell-laden samples were fixed and incubated with Click-iT reaction cocktail. Samples were then incubated with anti-Oregon green HRP, followed by incubation in Amplex UltraRed reaction mixture. The reaction was stopped after 15 minutes, absorbance of each well was measured, and absorbance of the blank was subtracted from each sample. Absorbance from GelMA-cECM patches was normalized to the absorbance of GelMA patches for each n.

Rat Surgery and Imaging –

All animal experiments were performed with the approval of the Institutional Animal Care and Use Committee of Emory University. GelMA-cECM patches used for in vivo experiments were created in the same way as described in the Bioprinting section, with the difference being that AF568 dye was allowed to bind to cECM before patch formation, similar to the methodology described in the Imaging Printed Test Grids section. Sprague-Dawley Rats (~250g in weight) were anesthetized with 2% isoflurane, intubated, and placed on the ventilator (Hallowell Emc Microvent 1). Following thoracotomy, the pericardial sac was then very carefully exposed and pulled back. The patch was then gently placed over the right ventricle of the heart, ensuring that there was no folding of the patch. Patches were left without further attachment, tucked underneath the pericardium, or attached the ventricle using a single suture. After 7 and 14 days, rats were sacrificed and hearts were excised. Hearts were imaged using an Odyssey CLx (Li-Cor) for both patch fluorescence and heart background fluorescence, with an acquisition area of 20.07 mm x 20.07 mm.

Immunohistological Analysis –

Excised hearts were fixed with 4% PFA, sectioned, and mounted onto coverslips. Tissue sections were incubated overnight with IsolectinB4-FITC in blocking buffer (3% BSA, 0.1% Triton X-100, 1x PBS) at 4°C. Sections were then incubated with DAPI and mounted. Tissue sections were imaged on an Olympus FV1000 Confocal Microscope.

This study was supported by the Betkowsi Family Research Fund to Michael E. Davis and the NHLBI (R01HL113468 to Karen L. Christman). Donald Bejleri was supported by the Cell and Tissue Engineering NIH Biotechnology Training Grant (T32 GM008433).