Fibrin-Enriched Cardiac Extracellular Matrix Hydrogel Promotes In Vitro Angiogenesis

Abstract

Angiogenesis is essential for cardiac repair after myocardial infarction. Promoting angiogenesis has been demonstrated as an effective approach for myocardial infarction treatment. Several different strategies for inducing myocardial angiogenesis have been explored, including exogenous delivery of angiogenic genes, proteins, microRNAs, cells, and extracellular vesicles. Various types of injectable hydrogels have been investigated for cardiac tissue repair. One of the most promising injectable hydrogels in cardiac regeneration is a cardiac extracellular matrix hydrogel that is derived from decellularized porcine myocardium. It can be delivered minimally invasively via transendocardial delivery. The safety and efficacy of cardiac extracellular matrix hydrogels have been shown in small and large animal myocardial infarction models as well as clinical trials. The main mechanisms underlying the therapeutic benefits of cardiac extracellular matrix hydrogels have been elucidated and involved in the modulation of the immune response, downregulation of pathways related to heart failure progression and fibrosis, upregulation of genes important for cardiac muscle contraction, and enhancing cardiomyocyte differentiation and maturation from stem cells. However, no potent capillary network formation induced by cardiac extracellular matrix hydrogels has been reported. In this study, we tested the feasibility of incorporating a fibrin matrix into cardiac extracellular matrix hydrogels to improve the angiogenic properties of the hydrogel. Our in vitro results demonstrate that fibrin-enriched cardiac extracellular matrix hydrogels can induce robust endothelial cell tube formation from human umbilical vein endothelial cells and promote the sprouting of human mesenchymal stem cell spheroids. The obtained information from this study is very critical toward the future in vivo evaluation of fibrin-enriched cardiac extracellular matrix hydrogels in promoting myocardial angiogenesis.

Graphical Abstract

1. INTRODUCTION

Promoting angiogenesis is one of the most effective strategies for improving the recovery of ventricular function after myocardial infarction (MI).^1-3 The root cause of MI is the reduced blood flow to the infarcted region, which restricts oxygen supply to the cardiomyocytes and ultimately leads to myocardial necrosis. Angiogenesis can improve revascularization, rescue dying cardiomyocytes, and reduce scarring and adverse left ventricular remodeling.^4-11 Furthermore, locally enhanced angiogenesis is a vital requirement for the long-term survival of therapeutic cells transplanted to an ischemic heart.^4,12,13 Several different strategies for inducing myocardial angiogenesis have been explored, including exogenous delivery of angiogenic genes, proteins, microRNAs, cells, and extracellular vesicles.^14-19 Successful preclinical studies and promising results of early clinical trials have demonstrated the great potential of myocardial angiogenesis for MI treatment.^20-23

Injectable hydrogels have become a powerful tool to be exploited in minimally invasive, catheter-based heart procedures. Various types of injectable hydrogels have been investigated for cardiac tissue repair.^24-26 For example, methacrylic anhydride (MA)–hyaluronic acid (HA) hydrogel has been used to deliver exosomes derived from human mesenchymal stem cells (MSCs) to the post-MI porcine heart through intrapericardial injection using a catheter.²⁷ Algisyl-LVR, a commercially available acellular alginate hydrogel, has been tested clinically in patients with MI and showed preservation of cardiac function after injection.²⁸ One of the most promising injectable hydrogels in cardiac regeneration is a cardiac extracellular matrix (cECM) hydrogel that is derived from decellularized porcine myocardium.^24,29,30 It can be delivered minimally invasively via transendocardial delivery.³¹ The safety and efficacy of cECM have been tested using small and large animal MI models.³² The outcomes of the recent phase 1 clinical trial support the safety and feasibility of injecting cECM hydrogels into post-MI patients and suggest the potential efficacy in chronic MI patients.³³ The main mechanisms underlying the therapeutic benefits of cECM hydrogels have been elucidated and involved in the modulation of the immune response, downregulation of pathways related to heart failure progression and fibrosis, and upregulation of genes important for cardiac muscle contraction.³⁴ In vitro studies also show that cECM hydrogels could enhance the differentiation and maturation of cardiomyocytes derived from pluripotent stem cells.^35,36 However, to date, there is no research evidence to suggest that cECM has a potent capability to induce capillary network formation.

In the present study, we aim to develop a proangiogenic cECM hydrogel for myocardial angiogenesis applications. We hypothesized that the incorporation of a fibrin matrix into cECM hydrogels would improve the angiogenic properties of the hydrogel. The role the fibrin matrix plays in stimulating and regulating angiogenesis has been extensively studied.^37-39 A large range of in vitro angiogenesis models have utilized the fibrin matrix as a culture system. When cultured on top or within a fibrin matrix, endothelial cells would rapidly form interconnecting capillary networks.^40,41 Here, we fabricated a hybrid hydrogel (Fn-cECM) by blending cECM with the fibrin matrix (Fn). We characterized the material properties of Fn-cECM hydrogels and compared them with those of cECM hydrogels. Using human umbilical vein endothelial cells (HUVECs) and human mesenchymal stem cells (hMSCs), we tested whether Fn-cECM hydrogels could induce in vitro angiogenesis without the addition of soluble factors. The obtained results are very critical to further optimize Fn-cECM hydrogels and evaluate their efficacy in promoting angiogenesis in MI models.

2. MATERIALS AND METHODS

2.1. Materials.

Human mesenchymal stem cells, human umbilical vein endothelial cells, EGM-2 BulletKit, 0.25% trypsin for hMSCs, and dissociation reagents kit for HUVECs (HBS, 0.25% trypsin, and TNS) were purchased from Lonza (Walkersville, MD). DMEM high in glucose with L-glutamine, fetal bovine serum, and penicillin–streptomycin were acquired from Gibco (Carlsbad, CA). Sodium dodecyl sulfate (SDS) was obtained from Bio-Rad (Hercules, CA). Pepsin, collagenase type-I, fibrinogen, tris-buffer saline, sodium hydroxide, thrombin, and calcium chloride were purchased from Sigma-Aldrich (St. Louis, MO). Dulbecco’s phosphate-buffered saline (DPBS) was obtained from Corning Incorporated (Bedford, MA). AggreWell400 24-well plates, antiadherence rinsing solution, and reversible cell strainer (37 μm) were obtained from StemCell Technologies (Vancouver, Canada). ELISA quantification kit and VEGF-A were purchased from R&D Systems (Minneapolis, MN). Paraformaldehyde (4%), Triton X-100, and goat serum were purchased from Thermo Fisher Scientific (Florence, KY). Anti-CD31, anti-vWF, and Alexa Fluor 488 and 647 were obtained from Abcam (Cambridge, MA). Calcein AM, Hoechst, and DAPI were acquired from Invitrogen (Waltham, MA).

2.2. Fabrication of Cardiac Extracellular Matrix Hydrogels.

Cardiac extracellular matrix (cECM) hydrogels were produced following a previously established protocol.^30,42 Briefly, fresh adult porcine hearts were obtained from a local USDA-approved butcher (Fischer’s Meat Market and Grocery, Muenster, TX). The myocardium was isolated from the fresh whole heart after removing excessive vessels, fat, and connective tissues. The tissues were then sliced into 1 mm thick pieces and rinsed thoroughly with deionized (DI) water to remove blood, followed by decellularization with 1% SDS for 4 days with solution replaced with a fresh one every day.

The decellularized myocardial tissue was washed with excessive DI water overnight to completely remove SDS residue and then lyophilized. The lyophilized tissue was milled into a fine powder under liquid nitrogen and then enzymatically digested in 0.01 M hydrogen chloride (HCl) containing 1 mg/mL pepsin at room temperature for 24 h under constant stirring. After the powder was completely solubilized, 0.1 M sodium hydroxide (NaOH) and 10× PBS were added to adjust the pH to 7.4. The neutralized solution was diluted to desired concentration using 1× PBS and incubated at 37 °C to form cECM hydrogels.

2.3. Fabrication of Fibrin–Cardiac Extracellular Matrix Hybrid Hydrogels.

Fibrin–cardiac extracellular matrix hybrid (Fn-cECM) hydrogels were fabricated by blending the fibrin matrix (Fn) into cECM during hydrogel formation. Specifically, fibrinogen solution (100 mg/mL in tris-buffer saline) and thrombin (4 U/mL in 40 mM calcium chloride solution) were added to cECM pregel solution. The mixture was incubated at 37 °C to form Fn-cECM hydrogels. The final concentrations of fibrin (Fn) and cECM in the Fn-cECM hydrogel were 25 and 6.25 mg/mL, respectively.

2.4. Hydrogel Morphology.

The cardiac extracellular matrix (cECM), fibrin (Fn), and fibrin–cardiac extracellular matrix hybrid (Fn-cECM) were fractured after being frozen in liquid nitrogen and then lyophilized. The cross sections were sputter-coated with gold/platinum (Au/Pt) and then imaged with a scanning electron microscope (SEM, Hitachi, S-4800, Japan).

2.5. Turbidimetric Gelation Kinetics.

The gelation kinetics of cECM, Fn, and Fn-cECM hydrogels was studied turbidimetrically⁴³ using a Synergy H1 Hybrid microplate reader (BioTek) preheated to 37 °C. For each hydrogel, 100 μL was added in a 96-well plate in triplicate, and the absorbance was recorded at 405 nm every 2 mins over 60 mins. Normalized absorbance (NA) was calculated using the equation NA = (A − A₀)/(A_max − A₀), where A is the absorbance at a given time, A₀ is the initial absorbance, and A_max is the maximum absorbance.

2.6. Swelling Ratio and In Vitro Degradation.

The swelling ratio was evaluated following a previously published protocol.⁴⁴ Briefly, cECM (12.5 mg/mL) and Fn-cECM hydrogels were formed in 1 cm diameter steel rings and treated with 1× DPBS for 24 h at 37 °C. The swollen weight (W_s) was obtained, and then the hydrogels were air-dried at 40 °C for 24 h to determine the dry weight (W_d). The percentage swelling was calculated using the equation (W_s − W_d)/W_d × 100. For studying enzymatic degradation, sample hydrogels were treated with collagenase type-I (250 U/mL) dissolved in 1× DPBS containing 0.9 mM CaCl₂ as previously described,^45-47 whereas control hydrogels were treated with 1× DPBS alone. Wet hydrogel weights of the sample (W_s) and control (W_c) were measured after 5 and 24 h of incubation at 37 °C. The percentage of remaining weight was calculated using the equation (W_s/W_c) × 100%. n = 3 for the two types of hydrogels at each time point.

2.7. Rheometric Analysis.

The mechanical properties of both cECM and Fn-cECM hydrogels were determined using a parallel plate rheometer (ARES RFS III). The pregel solutions were added into 1 cm diameter steel rings and incubated overnight at 37 °C. The following day, the hydrogels were subjected to mechanical testing, and the gap distance between the two parallel plates was maintained at 0.8 mm for all of the hydrogels. Dynamic frequency sweep tests were performed over a frequency range of 0.1 to 100 rad/s within the linear viscoelastic strain region of 1.5%, which was determined experimentally with a dynamic strain sweep test at a constant frequency of 0.5 rad/s. The storage modulus (G′) determined from the dynamic frequency tests was reported over the range of 0.1–5 rad/s frequency. n = 5 for each type of hydrogel.

2.8. Culture of HUVECs and hMSCs.

Human umbilical vein endothelial cells (HUVECs) were cultured using an EGM-2 BulletKit supplemented with 1% penicillin–streptomycin (P/S) following the protocol provided by the manufacturer. Human mesenchymal stem cells (hMSCs) were cultured in DMEM supplemented with 10% fetal bovine serum (FBS) and 1% P/S at a seeding density of ~3,000 cells/cm². The cells were maintained in a humidified cell culture incubator at 37 °C and 5% CO₂, and the medium was replaced with a fresh one every other day for HUVECs and every 3 days for hMSCs. All experiments were performed using HUVECs at passages 2–4 and hMSCs at passage 5.

2.9. Cell Spheroid Formation.

hMSC spheroids were formed using AggreWell400 following the manufacturer’s protocol. Briefly, each well of the AggreWell400 plate was rinsed with 500 μL of antiadherence rinsing solution at 1300 g for 5 min and then washed with 2 mL of warm DMEM basal medium. After prepping the plate, 0.6 × 10⁶ cells/mL was added to each well (500 cells/microwell) with an additional 1 mL of medium, followed by centrifugation at 100 g for 3 min to evenly distribute the cells in all of the microwells. The cells were incubated at 37 °C for 5 days with 75% media changes performed every other day to form spheroids. After incubation, the spheroids were collected using a cell strainer.

2.10. In Vitro Angiogenesis Assay.

In vitro angiogenesis was performed using either cells or cell spheroids on two different substrates: cECM hydrogels and Fn-cECM hydrogels. First, a thin layer of either cECM or Fn-ECM hydrogels was formed inside of a 12-well plate by adding 19 μl (5 μL/cm²) of pregel solution to each well that was evenly spread with the flat rubber end of the piston from a 1 mL BD syringe. Then, the cells or cell spheroids were seeded onto the thin layer of hydrogels at specific density (HUVECs: 1 × 10⁵ cells/well; hMSCs: 3,000 cells/well; and hMSCs spheroids: 50 spheroids/well). After seeding, the cells and spheroids were cultured inside a cell culture incubator (Fisher Scientific Isotemp 3532 CO₂ Incubator) at 37 °C with 5% CO₂. Angiogenesis analysis was performed by staining the cells or cell spheroids using Calcein AM (2 μg/mL) and then analyzing the images by ImageJ software⁴⁸ with an angiogenesis analyzer plugin.⁴⁹

2.11. Expression of Endothelial Cell Markers.

Immunofluorescence staining was used to confirm the expression of endothelial cell markers CD31, vWF, and VE-Cadherin. The cells or cell spheroids were fixed with 4% paraformaldehyde in 1× PBS solution at room temperature for 15 min, followed by permeabilization with 0.1% Triton X-100 in 1× DPBS for 15 min. They were then blocked with 10% goat serum for 1 h at room temperature. After the permeabilization and blocking steps, the cells or cell spheroids were incubated with primary antibodies CD31 (1:50), vWF (1:200), and VE-Cadherin (1:50) at 4 °C overnight. At the end of incubation, the samples were thoroughly washed with 1× DPBS and incubated with DAPI (12 μM/mL) and goat-derived secondary antibodies conjugated with Alexa Fluor 488 (1:500 for HUVECs and 1:100 for hMSCs spheroids), Alexa Fluor 647 (1:100 for HUVECs), and Alexa Fluor 594 (1:200 for hMSCs spheroids). HUVECs were treated with the secondary antibody for 2 h at room temperature, whereas hMSC spheroids were treated for 5 h. Next, the washing step was repeated, followed by fluorescence imaging with an inverted AxioVision A1 microscope (Carl Zeiss) for HUVECs and a Ti-E inverted microscope (Nikon A1 confocal) for imaging hMSC spheroids.

2.12. VEGF Secretion of hMSCs on Hydrogels.

The VEGF secreted by hMSCs seeded on the different types of hydrogels was quantified with an ELISA kit by following the manufacturer’s protocol. For the quantification, about 150 μL of the hydrogel solutions was added into a 12-well plate and incubated at 37 °C overnight. The following day, 20,000 cells/mL was seeded in each well with an additional 1 mL medium and incubated at 37 °C. The medium was collected at the end of each time point, after which 2 mL of 250 U/mL collagenase type-1 was added to the wells and incubated at 37 °C for 24 h. The medium collected was centrifuged at 600 g for 5 min to discard any residual cells. The following day, the dissolved hydrogel solution was centrifuged and filtered with a 0.2 μm filter to remove debris. All of the samples (n = 3) collected over 21 days were tested for the amount of VEGF secreted using ELISA.

2.13. Statistical Analysis.

All of the data were reported as means ± SD with 3–10 independent samples. The statistical analysis was performed using GraphPad Prism 9. Comparison of two groups was performed with the unpaired, two-tailed Student’s t-test, while for multiple groups, two-tailed two-way ANOVA was performed with Tukey’s post hoc test. The data were considered statistically significant for a p-value less than 0.05.

3. RESULTS

3.1. Morphology and Gelation Time of Fn-cECM Hydrogels.

The Fn-cECM hydrogel fabrication process was determined by tuning the concentrations of fibrinogen, thrombin, and cECM solutions during the blending process. Various concentrations of fibrinogen (from 12.5 to 100 mg/mL), thrombin (1, 2, and 4 U/mL), and cECM solution (from 1 to 12.5 mg/mL) were tested, and the gelation times of the formed hydrogels were measured. For our chosen combination of solution concentrations, the gelation time was ~30 min, which is in the range of catheter-deliverable hydrogels.^50-52 Compared with the cECM hydrogel that looks slightly translucent at the same concentration, the Fn-cECM hydrogel looks opaque with well-rounded edges (Figure 1A). SEM images showed that cECM hydrogels have a nanofibrous network with some sheets and that fibrin hydrogels have a general fibrous structure (see the Supporting Information). The Fn-cECM hydrogel also has a combination of nanofibers and sheets that appear much similar to the cECM hydrogel (Figure 1A). The gelation kinetic data demonstrate that Fn-cECM solution starts transitioning into a gel state faster than cECM solution (Figure 1B). The cECM hydrogel gelation curve has a typical sigmoidal shape, and the gelation started after a lag phase (T_lag = 13.33 ± 1.15 min), whereas the gelation kinetic curve of Fn-cECM does not have any lag phase (T_lag = 0) (Figure 1C). The time to reach 50% gelation (T_1/2) for cECM hydrogels was 17.67 ± 1.15 min and 12.67 ± 0.58 min for Fn-cECM hydrogels.

Figure 1.

Hydrogel morphology and gelation properties. (A) Macroscopic view of cardiac extracellular matrix (cECM) and hybrid (Fn-cECM) hydrogels with their respective scanning electron microscope (SEM) images. (B) Gelation kinetics of the hydrogels. (C) T_lag and T_1/2 of the hydrogels. Scale bar: 10 μm. n = 3. **p < 0.01, ****p < 0.0001.

3.2. Mechanical Properties, In Vitro Swelling, and Degradation of Fn-cECM Hydrogels.

The storage moduli (G′) and complex moduli (G*) of Fn-cECM hydrogels were measured and compared with cECM hydrogels. Both G′ and G* of Fn-cECM hydrogels were significantly higher than those of cECM hydrogels (897 ± 110 Pa vs 138 ± 27 Pa for G′; 915 ± 111 Pa vs 140 ± 27 Pa for G*) (Figure 2A,B). The swelling behavior of the hydrogels was studied in vitro. Figure 2C shows that the swelling ratio of Fn-cECM (3545 ± 285%) was significantly lower than that of cECM (5152 ± 94%). The transition from the dry state into hydrogels increased the mass of Fn-cECM by over 35 times and that of cECM by over 50 times. The enzymatic degradation of Fn-cECM and cECM hydrogels was examined. As shown in Figure 2D, there was no significant difference between the percentage of weight remaining for cECM and Fn-cECM hydrogels after 5 h immersion in collagenase solution. However, after 24 h, only 28 ± 6.27% of Fn-cECM hydrogels remained, which is significantly less than that of cECM hydrogels.

Figure 2.

Hydrogel characterization. (A) Storage moduli (G′) of cECM and Fn-cECM hydrogels. (B) Complex moduli (G*) of cECM and Fn-cECM hydrogels. n = 5. As G′ ≫ G″, the value of G* is similar to G′.(C) Swelling ratios of cECM and Fn-cECM hydrogels. (D) Degradation of cECM and Fn-cECM hydrogels in collagenase type-I. n = 3. **p < 0.01, ****p < 0.0001.

3.3. Fn-cECM Hydrogel Induces Endothelial Cell Tube Formation.

To test the proangiogenic capacity of Fn-cECM, we performed a tube formation assay. After 16 h of seeding HUVECs onto Fn-cECM hydrogels, extensive tubelike structures formed and linked with each other to form networks (Figure 3A). Strong expression of endothelial cell markers CD31 and VWF are detected (Figure 3B,C). HUVECs also attached to cECM hydrogels and expressed endothelial cell markers after seeding, but only a small percentage of cells could form vascular networks (Figure 3D-F). Image analysis results confirm that Fn-cECM induces a higher level of endothelial cell tube formation evidenced by forming a higher number of nodes, tubes, total tube length, meshes, mean mesh size, and total mesh area when compared with cECM hydrogels (Figure 3G). Similarly, HUVECs also formed networks on fibrin (Fn) hydrogels; however, they were not as prominent as those on Fn-cECM hydrogels (see the Supporting Information).

Figure 3.

Endothelial cell tube formation assay. (A, D) Vascular network formation and expression of (B, E) CD31 of HUVECs on Fn-cECM (A, B) and cECM (D, E) hydrogels. (C, F) Expression of vWF for HUVECs seeded on Fn-cECM (C) and cECM (F) hydrogels. Scale bar: 200 μm in A, D and 100 μm in B, C, E, and F. (G) Quantification of angiogenesis parameters using ImageJ software. n = 7; *p < 0.05, ****p < 0.0001.

3.4. Fn-cECM Hydrogel Interacts with hMSCs.

Similar to the effects on HUVECs, Fn-cECM hydrogels also induce more robust tube network formation of seeded hMSCs compared with cECM hydrogels (Figure 4A,B). The difference between the number of nodes, tubes, total tube length, meshes, and total mesh area derived from the networks formed on the two types of hydrogels is still significant (Figure 4C). However, less fold changes of these parameters are observed when compared with seeding with hMSCs instead of HUVECs on Fn-cECM and cECM hydrogels. For example, the average tube number of HUVECs increased 4-fold when cultured on Fn-cECM hydrogels other than cECM hydrogels, while it only increased 1.5-fold for hMSCs. There is no significant difference in the mean mesh size on the two types of hydrogels. In addition, no detectable expressions of CD31 and VWF were found on the cells forming networks on Fn-cECM hydrogels. We also measured VEGF secreted from the hMSCs seeded on the two types of hydrogels for up to 21 days (Figure 4D). It is noticeable that beginning from day 14, more VEGF was secreted by the hMSCs seeded on Fn-cECM hydrogel.

Figure 4.

Vascular network formation and VEGF secretion of hMSCs seeded on the hydrogels. (A, B) Calcein AM staining of hMSCs seeded on cECM and Fn-cECM hydrogels. Scale bar: 200 μm. (C) Quantification of angiogenesis parameters using ImageJ software. n = 7. (D) VEGF secreted by hMSCs seeded on cECM and Fn-cECM hydrogels quantified over a period of 3 weeks using ELISA. n = 3. ***p < 0.001, ****p < 0.0001.

3.5. Fn-cECM Hydrogel Promotes Sprouting of hMSC Spheroids.

Since we did not detect the expressions of CD31 and vWF for hMSCs seeded on Fn-cECM, we examined whether a three-dimensional (3D) culture would affect the response of hMSCs to Fn-cECM hydrogels. We performed an in vitro spheroid sprouting assay to test the influence of Fn-cECM hydrogels on 3D-cultured hMSCs. After 24 h of seeding hMSC spheroids on either cECM or Fn-cECM hydrogels, we observed the formation of sprout structures on both hydrogel types with a distinct pattern (Figure 5A,B). The tip cells of spheroids on cECM hydrogels formed short sprouts with weak interactions, while on Fn-cECM hydrogels, the tip cells formed well-defined sprouts with long capillary-like structures. Further quantitative assessment confirmed that hMSC spheroids cultured on Fn-cECM hydrogels had more sprouts coming out of the spheroids (Figure 5C). The average and cumulative sprouting lengths of the sprouts grown on Fn-cECM hydrogels were also significantly longer (Figure 5D,E). The spheroid area, which is defined as the area covered by the core of the spheroid for both hydrogel types, did not show a statistically significant difference (Figure 5F). However, the spheroid sprouting area, which is the total area covered by sprouts only, was significantly higher for Fn-cECM hydrogels (Figure 5G).

Figure 5.

Spheroid sprouting assay for hMSCs seeded on hydrogels. (A, B) Spheroid sprouting from Calcein-AM-stained hMSCs seeded on cECM and Fn-cECM hydrogels. Scale bar: 200 μm. (C–F) Quantification of spheroid sprouting using ImageJ software. n = 10. *p < 0.05, ****p < 0.0001.

3.6. Fn-cECM Hydrogel Enhances Endothelial Differentiation of hMSC Spheroids.

To verify whether the sprouting structures we observed from the in vitro spheroid sprouting assay are angiogenic sprouting, we stained the samples with endothelial cell markers CD31 and VE-Cadherin. As shown in Figure 6A,B, we detected CD31⁺ and VE-Cadherin⁺ endothelial cell populations in hMSC spheroids cultured in hMSC cell culture medium. Most of the VE-Cadherin-expressing cells were located near the edge of the spheroids. There are more cells expressing CD31 than VE-Cadherin within the spheroids when they are cultured in the medium. When cultured on cECM hydrogels and Fn-cECM hydrogels, more cells not only within spheroids but also in sprouting structures express CD31 (Figure 6C,E). The VE-Cadherin⁺ endothelial cells dramatically increased in the center area of the spheroids when cultured on Fn-cECM hydrogels (Figure 6B,D,F).

Figure 6.

Immunofluorescence staining of hMSC spheroids with endothelial cell markers CD31 (A, C, and E) and VE-Cadherin (B, D, and F) under different culture conditions. (A, B) hMSC spheroids cultured in cell medium. (C, D) hMSC spheroids cultured on cECM hydrogels. (E, F) hMSC spheroids cultured on Fn-cECM hydrogels. Scale bar: 100 μm.

4. DISCUSSION

The motivation of this study is to develop an injectable hydrogel for cardiac tissue repair by enhancing the benefits of hydrogels derived from decellularized cardiac ECM tissue with the angiogenic properties of fibrin. Both cECM and fibrin hydrogels are very promising biomaterials that provide numerous advantages in functioning as tissue engineering scaffolds and cell delivery platforms for treating injured myocardium.^53-56 Both are natural materials, fully degradable, and favor cell interactions.^57-59 In addition, each of them has unique features enabling their cardiac applications. The most appealing features of cECM hydrogels are as follows: (1) retaining cardiac tissue-specific ECM composition; (2) serving as a reservoir for anchored cytokines and growth factors; and (3) demonstrating safety in minimally invasive intramyocardial delivery through a catheter in a clinical setting.^60,61 Unlike the cECM hydrogel, which is an emerging biomaterial, fibrin is one of the classical and well-known biomaterials that has been widely used for many applications.^62-65 One well-proven benefit of fibrin hydrogels is that they can promote angiogenesis (see the Supporting Information Figure S2). It has been reported that fibrin hydrogels led to endothelial assembly into microvessels and angiogenic sprouting both in vitro and in vivo.^66,67 In this study, we demonstrate the feasibility of physically blending cECM and fibrin to form an injectable hydrogel with potential for cardiac applications. It needs to be noted that another research group also investigated developing a hybrid of cECM and fibrin, but the reported fabrication process requires additional crosslinkers, e.g., transglutaminase (TG), to achieve the desired mechanical properties. Furthermore, the research focus is to investigate how stiffness and ECM composition of the cECM and fibrin hybrid affect cardiovascular differentiation of cardiac progenitor cells,⁶⁸ while ours aims to optimize hydrogel properties to promote angiogenesis.

Our obtained results suggest that the incorporation of a fibrin matrix into cECM hydrogels has multifaceted effects on hydrogel material properties. First, it enables the tuning of hydrogel gelation to be more favorable for our intended cardiac applications. cECM and Fn hydrogels have distinct gelation curves. The gelling behavior of Fn-cECM hydrogels follows the same pattern as Fn hydrogels (see the Supporting Information Figure S1) with accelerated gelation compared with cECM hydrogels. The Fn-cECM hydrogel we developed has the gelation kinetics that enables sufficient time to mix pregel solutions and pass the solution mixture through a catheter and timely transition to gel after delivery. Second, the incorporation of a fibrin matrix into cECM hydrogels increases mechanical stiffness and reduces the swelling of the hydrogel, which are beneficial to provide mechanical support for damaged cardiac tissue and avoid heart wall disruption after hydrogel injection. Many studies have demonstrated that the mechanical properties of substrate would affect cell behaviors. For example, endothelial cell spreading and extension formation increase with matrix stiffness.⁶⁹ Our study also confirmed that Fn-cECM hydrogels induced higher levels of tube network formation due to their higher mechanical strength compared to cECM hydrogels.

Third, we noticed that Fn-cECM hydrogels degraded significantly faster than cECM hydrogels when treated with collagenase solution for 24 h. Although the reason that causes the rapid enzymatic degradation is unclear, it can be a potential drawback in using Fn-cECM hydrogels in cardiac treatment. We will examine the in vivo degradation rate of Fn-cECM hydrogels in our future animal studies and further modify it by adding MMP inhibitors such as doxycycline when necessary.⁷⁰ Last but most importantly, the incorporation of a fibrin matrix into cECM hydrogels enhanced the proangiogenic properties of the hydrogel. Our results demonstrate that Fn-cECM hydrogels promote rapid and robust vascular network formation of HUVECs, which have not been observed in cECM hydrogels in vitro.

Our results prove that blending Fn could be a simple and effective strategy to enhance the angiogenic properties of existing hydrogels. This strategy has been explored by several research groups with other types of hydrogels.^71-75 For example, a collagen/fibrin mixed construct containing HUVECs has been shown to induce lumen formation 7 days after being implanted subcutaneously in mice.⁷⁶ Blending fibrin in a composite hydrogel containing alginate and iron nanoparticles has been reported to enhance the attachment of endothelial cells and promote strong endothelization.⁷⁷ With the increasing interest in utilizing organ-specific ECM hydrogels for disease treatment and the critical role angiogenesis plays in tissue repair, we expect the approach in this study can be readily adopted by many researchers to benefit a broader scientific community.

We also evaluated the response of MSCs to Fn-cECM hydrogels to test whether the proangiogenic feature of our hydrogel could induce endothelial differentiation of MSCs. MSCs are one of the most popular cell types used in cardiac cell therapy.^78-80 Many completed and ongoing clinical trials have been conducted to investigate the efficacy of MSCs in MI treatment.^81-88 The therapeutic effects of MSCs include their ability to differentiate into cardiovascular cells, immunomodulatory properties, antifibrotic activity, and ability to undergo neovasculogenesis.^89-92 In this study, the distinct effects of Fn-cECM on MSCs and MSC spheroids are identified. Fn-cECM could cause morphological alteration of MSCs toward vascular tube formation and VEGF secretion but is insufficient in directing them to express endothelial cell-specific markers. However, when it acts on MSC spheroids, it could not only enhance angiogenic sprouting but also upregulate the expression of endothelial cell-specific markers of cells in MSC spheroids. We observe that culturing MSCs in 3D as spheroid triggers their differentiation toward endothelial cells. The synergy between 3D culture and Fn-cECM hydrogels facilitates the endothelial differentiation process and promotes sprouting angiogenesis. These findings suggest the potential use of Fn-cECM hydrogels to deliver MSC spheroids to increase MSC functionalization for enhanced therapeutic applications.

Although the developed Fn-cECM hydrogel demonstrated enhanced material properties and improved angiogenic capacity, there are some issues that could potentially affect its successful application in myocardial infarction treatment. The Fn-cECM hydrogel is injectable (see the Supporting Information), but whether it is compatible with catheter delivery in a clinical setting is unknown. The Fn-cECM hydrogel is degradable. Its in vivo degradation rate would affect the therapeutic outcomes and should be carefully examined. In this study, we used a 2D angiogenesis assay by seeding the cells and cell spheroids on the hydrogels to test the angiogenic properties of Fn-cECM hydrogel. Although the 2D angiogenesis assay is widely used by biomaterials researchers and has many advantages, the cells seeded on the hydrogel and encapsulated inside the hydrogel may respond differently. In our future studies, we will perform a 3D angiogenesis assay to further investigate the effects of Fn-cECM hydrogel on encapsulated cells and conduct in vivo study to investigate the feasibility and efficacy of using the developed Fn-cECM hydrogel to promote myocardial angiogenesis.

5. CONCLUSIONS

In this study, we presented the development of a proangiogenic cECM hydrogel. We demonstrated the feasibility of incorporating a fibrin matrix into cECM hydrogels by blending. We have found that our developed Fn-cECM hydrogel has accelerated gelation and improved mechanical properties. We confirmed that Fn-cECM hydrogels promote in vitro angiogenesis through a vascular tube formation assay. Additionally, we found that Fn-cECM hydrogels increase the secretion of VEGF of MSCs and enhance the angiogenic sprouting of MSC spheroids. The approach we exploited in this study is a simple and effective strategy to enhance the angiogenic properties of existing hydrogels.