Article pubs.acs.org/journal/abseba
Fibroblast Growth Factor‑1 Released from a Heparin Coacervate Improves Cardiac Function in a Mouse Myocardial Infarction Model Zhouguang Wang,†,‡ Daniel W. Long,†,§ Yan Huang,†,‡,§ Sinan Khor,⊥ Xiaokun Li,‡ Xiao Jian,*,‡ and Yadong Wang*,† †
Department of Bioengineering, Swanson School of Engineering, University of Pittsburgh, Pittsburgh, Pennsylvania 15261, United States ‡ School of Pharmacy, Key Laboratory of Biotechnology and Pharmaceutical Engineering, Wenzhou Medical University, Wenzhou 325035, China ⊥ Department of Molecular Pharmacology, Albert Einstein College of Medicine, Bronx, New York 10461, United States S Supporting Information *
ABSTRACT: Emerging evidence supports the beneficial effect of fibroblast growth factor-1 (FGF1) on heart diseases, but its application has been hindered by the short half-life and limited bioactivity of the free protein. We designed an injectable coacervate to facilitate robust growth factor delivery, which would both protect and increase the bioactivity of growth factors. In this study, a model for acute myocardial infarction was established in mice, and the cardioprotective effect of the FGF1 coacervate was investigated. Echocardiographic results showed that the FGF1 coacervate inhibited ventricular dilation and preserved cardiac contractibility more than the free FGF1 and the saline control within the 6-week duration of the experiments. Histological examination revealed that the FGF1 coacervate reduced inflammation and fibrosis post-MI, significantly increased the proliferation of endothelial and mural cells, and resulted in stable arterioles and capillaries. Furthermore, the FGF1 coacervate improved the proliferation of cardiac stem cells 6 weeks post-MI. However, free FGF1, dosed identically, did not show significant difference from saline treatment. Thus, one injection of FGF1 coacervate was sufficient to attenuate the injury caused by MI, and the results were significantly better than those obtained from an equal dose of free FGF1. KEYWORDS: controlled release, myocardial infarction, fibroblast growth factor-1, heparin, coacervate
1. INTRODUCTION Despite decades of research, cardiovascular disease remains the largest cause of death as well as a major contributor to disability, affecting 5.8 million people in the United States with an estimated annual healthcare cost of $300 billion.1 Within the myocardial infarction (MI) zone, insufficient blood supply to a region triggers both cell death and subsequent pathological remodeling, which eventually results in heart failure.2 There has been a considerable amount of research interest in developing novel therapies, such as gene therapy, stem cell therapy, and direct administration of pro-angiogenic growth factors.3 Although preclinical studies and initial clinical trials supported the beneficial effects of pro-angiogenic factors,4,5 patients failed to show appreciable improvement in double-blinded clinical trials.6−8 These negative findings may have resulted from a dearth of knowledge regarding how blood vessels form as well as the body’s endogenous responses to ischemia as well as growth factor selection and/or the timing that growth factors reach the target zone. Moreover, the extent by directly administered growth factors are beneficial is restricted by their short half-lives after injection; enzymatic deactivation and © XXXX American Chemical Society
proteolytic degradation occur within minutes of treatment. In this study, an injectable biocompatible and biodegradable coacervate is used to control the release of heparin-binding growth factors and offer an effective method of extending fibroblast growth factor 1 (FGF1) bioactivity. We have developed a controlled delivery system that utilizes the charge interaction between a heparin, a natural polyanion, and poly(ethylene argininylaspartate diglyceride) (PEAD), a biodegradable polycation, which together form a complex coacervate. The coacervate is a phase separation of liquids where the polycation and polyanion forms a neutral complex and separates from the surrounding aqueous environment, encapsulating the heparin-binding proteins and protecting them from degradation. This heparin-based coacervate delivery platform functions to both protect and release heparin-binding growth factors, including nerve growth factor,9 fibroblast Special Issue: Tissue Engineering Received: August 31, 2016 Accepted: March 30, 2017 Published: March 30, 2017 A
DOI: 10.1021/acsbiomaterials.6b00509 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX
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ACS Biomaterials Science & Engineering growth factor-2 (FGF2),9 heparin-binding epidermal growth factor-like growth factor,10 and stromal cell-derived factor-1α.11 Additionally, this system is retained within the heart for up to one month in vivo after injection.12 In our previous study, we proved that Sonic Hedgehog (Shh) delivered by the coacervate combined with a degradable hydrogel is cardio-protective and promotes heart function and angiogenesis in rodents after MI.13,14 We also utilized fibrin gel and the coacervate in order to sequentially deliver vascular endothelial growth factor (VEGF) and platelet-derived growth factor (PDGF) to rat ischemic hearts. Within 1 week, VEGF was released, and PDGF release was sustained a minimum of 3 weeks in vitro. Sequentially releasing VEGF and PDGF significantly enhanced heart function, which was assessed by measuring cardiac contractility through fractional area change (FAC), in an acute MI model in rats. We also demonstrated enhanced vascularization and survival of cardiomyocytes as well as reduced fibrosis, inflammation, left ventricle (LV) wall thinning, scar expansion, and fibrosis post-MI.15 FGF1 is a member of the FGF family and has many functions. FGF1 is synthesized by many different cell types, including cardiomyocytes, endothelial cells, and fibroblasts.16 FGF1 has shown therapeutic potential in burns and wound healing.17 Previous studies have reported that in acute ischemia and reperfusion, FGFs exert cardioprotective effects by inducing an ischemic preconditioning (IPC)-like state.18−20 In the heart, FGF1 with its receptor FGFR1 is vital in the regulation of cardiac morphogenesis, arteriogenesis, and angiogenesis.21 Furthermore, FGF1 greatly enhances cardiac remodeling after MI through the induction of cardiomyocyte proliferation.22 By enhancing angiogenesis and reducing injury from ischemia-reperfusion, FGF1 would be expected to effectively treat cardiac ischemia. Indeed, constitutive overexpression of cardiac-specific FGF1 can delay myocardial infarct formation in vivo.20 However, one major obstacle of growth factor therapies in the treatment of heart disease is their short half-lives in vivo.23 Additionally, FGF1 in its free form possesses low levels of bioactivity but exhibits a much higher bioactivity when combined with heparin.24 Because of the use of heparin in our delivery system, we therefore aim to extend the half-life and improve the bioactivity of FGF1, investigating its potential to reduce cardiac scar burden and improve heart function postMI. In this study, we formed the FGF1 coacervate and researched its bioactivity on endothelial and cardiac stem cells in vitro. Its therapeutic efficacy was evaluated in a murine MI model. Notably, the histological examination revealed that the FGF1 coacervate reduced inflammation and fibrosis post-MI, significantly increased the proliferation of endothelial and mural cells, and resulted in stable vasculature. Finally, we observed significantly improved cardiac function due to the controlled delivery of FGF1 to infarcted hearts.
suspended in saline containing 500 ng of FGF1 was centrifuged at 12 100 × g for 10 min and the pellet stored at 37 °C. On days 0, 1, 4, 7, 10, 14, 21, and 28, the supernatant was aspirated and stored at −80 °C, and 200 μL of fresh saline was added to cover the pellet. The amount of released FGF1 in three separate fractions per time point was determined by ELISA (R&D Systems, Minneapolis, MN). Coacervate size was determined by dynamic light scattering as previously described.9 Heparin and PEAD were separately dissolved in DI water then combined at a 5:1 PEAD:heparin mass ratio in 1 mL total solution. Coacervate droplet size was then immediately measured using the Zetasizer Nano ZS (Malvern, Westborough, WA). 2.2. Measurement of Cell Proliferation and Migration in Vitro. Human umbilical vein endothelial cells (HUVEC) were purchased from ATCC (Manassas, VA) and cultured in EBM-2 media (Lonza, Basel, Switzerland) supplemented with 2.5% fetal bovine serum to simulate nutrient-deprived conditions. Human Sca1+/ckit+ cardiac stem cells (hCSC) were purchased from Celprogen Inc. (Torrance, CA) and plated on extracellular expansion matrixcoated plates (Celprogen Inc.). To simulate nutrient deprivation in hCSCs, hCSC media with serum (Celprogen Inc.) was diluted 1:4 with basal DMEM media (25% hCSC-CM).HUVECs and hCSCs were each seeded at 2 × 103 cells per well in a 96-well plate for proliferation assays and incubated in 100 μL of nutrient-deprived media overnight to allow cells to attach. .The following day, all cells were washed with DMEM, and 200 μL nutrient-deprived media containing no supplements, delivery vehicle, 50 ng/mL free FGF1, or 50 ng/mL FGF1 coacervate were added to each well to determine their effect on cell proliferation (n = 4 per group). This dose of FGF1 is based off previous in vitro studies utilizing FGF1.26 The plates were then incubated for 72 h at 37 °C. After washing all wells, CellTiter 96 AQueous One Solution Cell Proliferation Assay (MTS) reagent (Promega, Madison, WI, USA) in DMEM was added. The plate was incubated in 5% CO2 at 37 °C for 3 h, at which point the absorbance at 490 nm (with reference at 650 nm) was read with the Infinite 200 PRO plate reader (Tecan, Switzerland). The effect of FGF1 coacervate on the migration of hCSC and HUVEC was measured by transwell chemotaxis. Nutrient-deprived media containing no supplements, delivery vehicle, 50 ng/mL of free FGF1, or 50 ng/mL of FGF1 coacervate were loaded into bottom wells (n = 4 per group). hCSC and HUVEC were seeded in 24-well transwell inserts with 8 μm pore size (Millipore, Billerica, MA) at a density of 10 000 cells/cm2. After incubation at 37 °C for 12 h, nonmigrated cells were removed with cotton swabs. Migrated cells were fixed in methanol for 10 min and stained with Quant-iT PicoGreen dsDNA reagent (P7581; Thermo Fisher Scientific, Waltham, MA, USA). Fluorescent images were captured by Nikon Eclipse Ti fluorescence microscope equipped with NIS-Elements AR imaging software (both from Nikon, Tokyo, Japan). The number of migrated cells was quantified and averaged from 3 independent images taken in 3 different areas per sample per group (n = 4). The cell number of each group was individually normalized to the average number of cells in the basal media control group. 2.3. Mouse Acute MI Model and Intramyocardial Injection. Male Balb/cJ mice (Jackson Laboratory, Bar Harbor, ME) at 9−12 weeks old were used and cared for in compliance with the Institutional Animal Care and Use Committee of the University of Pittsburgh. A total of 39 mice were used in this study, 13 for each of three treatments. At 2 weeks, 4−5 mice were chosen from each group at random and sacrificed for 2-week analysis. The remaining 8−9 in each group were sacrificed 6 weeks post-MI. MI and intramyocardial injections were performed as we previously reported.12,27,28 Briefly, MI was induced by ligation of the left coronary artery. Five min after the induction of MI, a total volume of 35 μL saline, free FGF1 (500 ng of FGF1) or FGF1 coacervate (500 μg of PEAD, 100 μg of heparin and 500 ng of FGF1) was injected across three sites of the ischemic myocardium (one at the center and two at the border zone of the infarct). The FGF1 dose was chosen based on previous studies utilizing this coacervate system with other growth factors in a mouse MI model.12,27 FGF1-free coacervate was not tested as it has shown to have no effect in cardiac repair in our previous studies.15,27 The
2. MATERIALS AND METHODS 2.1. FGF1 Coacervate Characterization and Release ProfilePEAD and the Complex Coacervate were Prepared as Previously Described.9,25 PEAD and heparin were each dissolved separately in DI water at concentrations of 10 mg/mL. Heparin and FGF1 were first combined, then PEAD was added to form the coacervate. Following th addition of PEAD, the solution went from clear to turbid, indicating the coacervate had formed. The final mass ratios of PEAD:heparin:FGF1 were 500:100:1. The release profile of the FGF1 coacervate was determined in vitro as previously described.25 Briefly, 200 μL of FGF1 coacervate B
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Figure 1. Coacervate controls the release of FGF1 in a steady fashion. (A) Chemical structure of the PEAD polycation. (B, C) Analysis of coacervate droplet size with polydispersity index (PDI). (D) Confocal fluorescent imaging of fluorescein-labeled FGF1 shows spherical morphology with loading specifically within the coacervate droplet. Bar = 10 μm. (E) The release profile of FGF1 coacervate in vitro for 4 weeks as measured by ELISA. Data are presented as percent cumulative release (normalized to the original load). Error bars indicate means ± SD. surgeon performing the surgical procedures and injections was blinded to the treatment received by each mouse. 2.4. Echocardiography. Echocardiography was repeatedly performed by a blinded investigator before surgery and at 2 and 6 weeks postinfarction to assess cardiac function. Briefly, the heart and respiratory rates were continuously monitored while the body temperature was maintained at 37 °C by a hot pad. Echocardiographic parameters were measured using a high-frequency linear probe (MS400, 30 MHz) connected to a high-resolution ultrasound imaging system (Vevo 2100; FUJIFILM VisualSonics, Toronto, Ontario, Canada). End-systolic area (ESA) and end-diastolic area (EDA) were measured from short-axis images of the LV by B-mode. FAC was calculated as [(EDA-ESA)/EDA]*100%.this has been validated as an accurate prediction of cardiac contractility.29 LV ejection fraction (LVEF) was also calculated using echocardiography. The mice that were sacrificed for histological analysis prior to 6 weeks postinjection were not included in the echocardiographic study. 2.5. Histological Analysis. At 2 and 6 weeks postinfarction, the mice were sacrificed and the hearts were harvested following the established methods.30 The harvested hearts were frozen in OCT compound for staining. Specimens were serially sectioned at a thickness of 8 μm from apex to the ligation level (approximately 0.5 mm in length). For comparison of the ventricular wall thickness in the infarct zone, wall thickness was measured at the midinfarct point of
each ventricle on H&E stained slides using NIH ImageJ software. Seven hearts for each treatment were used for this quantification. For observation of fibrosis, Masson’s trichrome kit (IMEB, San Marcos, CA) was used to stain collagen fibers. Twelve sections from each treatment group were imaged to examine scar expansion through the ventricle wall. Inflammatory mast cells are known regulators of matrix metalloproteinase activity following MI.31 To quantify their infiltration, we performed toluidine blue staining, and mast cell density was measured in two fields of view within the border zone of each heart. All wall thickness, fibrosis, and mast cell measurements were taken from mice sacrificed at the 6 week time point. 2.6. Immunofluorescent Staining. For evaluation of inflammation, a rat antimouse CD68 (Abcam, Cambridge, MA) primary antibody was used, followed by an antirat IgG secondary antibody (Invitrogen, Carlsbad, CA). For detection of endothelial cells, a rat antimouse CD31 (BD Biosciences, San Jose, CA) and the same antirat IgG secondary antibody was used. For α-SMA staining, an FITCconjugated anti-α-SMA monoclonal antibody (Sigma-Aldrich, St. Louis, MO) was utilized. To examine murine CSCs, we first incubated sections overnight at 4 °C with rat antimouse CD117/c-kit (Cedarlane Laboratories, Burlington, NC). For the detection of cardiomyocytes, sections were incubated overnight at 4 °C with mouse anticardiac troponin T (cTnT) primary antibody (Abcam), followed by goat antimouse Alexa 488 IgG at RT for 1 h. To detect proliferating cells, C
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Figure 2. Bioactivity of FGF1 coacervate on hCSC and HUVEC proliferation and migration. (A) Representative immunofluorescent images showing migration of hCSC and HUVEC in response to different treatments through a transwell chemotaxis assay. (B, C) Quantification of proliferation data showed that FGF1 coacervate significantly increased proliferation relative to both basal media and free FGF1 in both hCSC and HUVEC. Free FGF1 also increased proliferation of HUVEC relative to basal media but not hCSC. (D-E) Quantification of migration data showed enhanced chemotaxis of HUVEC and hCSC in both the free FGF1 and FGF1 coacervate groups. However, the same dose of FGF1 delivered by the coacervate had greater chemotactic effects compared to the both basal media and the free FGF1 group in both HUVEC and hCSC (n = 4 per group; data normalized to the respective basal media). *p < 0.05, **p < 0.01 compared to basal media; #p < 0.05 compared to free FGF1. All quantitative data represent means ± SD. after the first staining for one of the cell lineage markers above, a second overnight incubation was performed at 4 °C with rabbit antimammalian Ki67 primary antibody (Abcam), followed by donkey antirabbit Alexa 488 IgG at RT for 1 h. The nuclei were stained with DAPI at RT for 10 min. Immunofluorescent images were taken using a Nikon Eclipse Ti fluorescence microscope equipped with NISElements AR imaging software (both from Nikon). For quantification, four to six sections from different hearts were used for each group. Measures of inflammation, angiogenesis, and progenitor cell populations were taken as cells per mm2 area; cTnT staining was measured by quantifying the fractional area of cTnT+ within the microscopic field. 2.7. Statistical Analysis. All data are presented as the mean ± standard deviation (SD). Significant differences between groups were analyzed by Student’s t test (two groups), one-way ANOVA (multiple groups), or two-way repeated ANOVA, and p ≤ 0.05 was considered significantly different. Statistical analyses were performed with GraphPad Prism 5.0 (GraphPad Software, La Jolla, CA, USA).
these positive charges on PEAD and the negative charges carried by the sulfates on heparin allow coacervate formation. FGF1 has a high-affinity binding site for heparin. Collectively, a ternary complex self-assembles: [PEAD:heparin:FGF1]. We fluorescently labeled FGF1 (DyLight 594, red), which showed that the coacervate droplets had an average size of 526.2 ± 106.4 nm (Figure 1 B-D). To measure the controlled release of proteins in vitro, the amount of FGF1 released from the coacervate was measured by ELISA at 0, 1, 4, 7, 10, 14, 21, and 28 days. As shown in our result, less than 10% FGF1 was detectable in the supernatant after centrifugation (day 0 of release assay), and therefore, the loading efficiency was greater than 90%. After the initial release, FGF1 coacervate released approximately 15.1 ± 2.5% during the first 24 h. The total release of FGF1 was estimated to be 79.8 ± 6.3% over the 28day duration (Figure 1E). These results showed that FGF1 was released in a sustained manner without an initial burst at an approximately linear 2.31% day−1. 3.2. FGF1 Coacervate Induces Endothelial Cell and Cardiac Stem Cell Chemotaxis and Proliferation. The effect of FGF1 coacervate on HUVEC and hCSC proliferation was investigated in vitro. Cells were seeded on tissue culture-
3. RESULTS 3.1. Characterization of FGF1 Coacervate. As shown in the chemical structure of PEAD, each repeating unit contains two functional groups with a positive charge: an ammonium and guanidinium group (Figure 1A). The interactions between D
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Figure 3. FGF1 coacervate mitigates MI-associated injury. (A) cTnT staining (red) showed few viable cardiomyocytes in the saline and free FGF1 groups, while the FGF1 coacervate group showed a larger area of viable cardiac muscle in the infarct zone. Scale bars = 50 μm. (B) Quantitative analysis shows a significantly higher density of viable cardiomyocytes in hearts receiving FGF1 coacervate relative to saline or free FGF1. (C) H&E staining showed that MI caused wall thinning and ventricular dilation in the saline control group. The higher magnification micrographs revealed damaged cardiac myofibers surrounded by scar tissue. Similar morphology was observed in the free FGF1 groups. FGF1 coacervate reduced the infarct area and partially preserved the normal tissue structure in the infarct zone. Scale bars = 1 mm (top) and 50 μm (bottom) (D-E) Quantitative analysis showed significantly increased ventricular wall thickness and reduced infarct area in the FGF1 coacervate group compared with the saline and free FGF1 groups. **P < 0.01, compared with saline, #p < 0.05 compared with free FGF1. Data represent means ± SD.
HUVECs and hCSCs in both the free FGF1 (50 ng/mL) and FGF1 (50 ng/mL) coacervate groups relative to basal media (p < 0.05 for free FGF1, p < 0.01 for FGF1 coacervate) (Figure 2A, D, E). However, an identical dose of FGF1 released by the coacervate exhibited greater chemotactic effects compared to the free FGF1 group in both HUVECs and hCSCs (p < 0.05) (panels D and E). The vehicle group demonstrated no effect on cell migration compared with the basal media group (p > 0.05). Thus,, these results indicate that FGF1 released by the coacervate is highly bioactive and stimulates the proliferation and migration of HUVECs and hCSCs in vitro. 3.3. FGF1 Coacervate Protects Cardiac Structure and Improves Cell Survival Post-MI. To further elucidate how FGF1 coacervate affects the structure and function of the heart, we performed cTnT immunofluorescent staining in each group to examine the survival of cardiomyocytes (Figure 3A). At 6 weeks post-MI, the saline and free FGF1 groups both showed significant reductions in cardiomyocyte (cTnT+) population in the infarct region. The FGF1 coacervate group, however, showed a significantly higher density of cTnT+ cells than the
treated polystyrene using nutrient-deprived media for the duration of the experiment in order to simulate nutrient starvation following coronary artery blockage. A fixed load of 50 ng/mL FGF1 was selected based on previous studies utilizing growth factors for in vitro assays.26 DMEM basal medium served as the negative control. The treatment of HUVEC with 50 ng/mL of free FGF1 or FGF1 coacervate led to an increase in HUVEC proliferation, which was statistically significant (p < 0.05) (Figure 2C). However, FGF1 coacervate could also significantly increase HUVEC proliferation in comparison to free FGF1 (p < 0.05) and saline (p < 0.01) (Figure 2C). Consistent with these results, the hCSC proliferation test also showed that the FGF1 coacervate significantly increased hCSC proliferation compared to free FGF1 (p < 0.05) and saline (p < 0.01) (Figure 2B). We did not observe any significant difference between the no-treatment control and the vehicle group in these two cell types (p > 0.05) (Figure 2A−C). We evaluated chemotaxis on HUVECs and hCSCs induced by FGF1 coacervate using a transwell assay. Fluorescent images of transwell insert membranes showed enhanced chemotaxis of E
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Figure 4. FGF1 coacervate reduces inflammation and fibrosis post-MI. (A) Fibrosis in the border zone at 6 weeks as examined by Masson’s trichrome staining. The saline control had many collagen fibers deposited along the inner wall. FGF1 coacervate effectively prevented propagation of fibrosis. Scale bars =1 mm for top row and 50 μm for bottom 2 rows. The second and third row are representative images showing collagen deposition in the infarct and border zones at higher magnification. (B) Distribution of macrophages in the infarct and border zone at 2 weeks as indicated by CD68 staining. Scale bars = 50 μm. (C) Quantitative analysis showed significantly reduced CD68+ cells in the FGF1 coacervate group compared with the saline and free FGF1 groups. *P < 0.05, **P < 0.01, compared with saline, #p < 0.05 compared with free FGF1. Data presented as means ± SD.
4A). Fibrotic tissue was observed up to the border zone in the saline control group, as shown by dense collagen deposition along the border zone’s ventricular wall (Figure 4A). In the free FGF1 group, we observed similar fibrosis in this region. In contrast, images from the FGF1 coacervate group had reduced collagen deposition and scar formation in the infarct area in comparison to the saline and free FGF1 groups. FGF1 coacervate visibly reduced the amount of collagen in the border zone, indicating its efficacy at reducing fibrosis. Consequently, the tissue would be more compliant and should enhance cardiac contractility after MI, as shown below. In addition to cardiac fibrosis, MI could also trigger systemic and local inflammation in the infarcted zone. In our study, we detected phagocytic cells within the infarct area utilizing a panmacrophage marker, CD68, at 2 weeks postinfarction. An earlier 2-week time point was selected for this analysis due to the large inflammatory response typically seen in the early acute phase of MI. Healthy myocardium has very few CD68+ cells; on the contrary, a significant number of CD68+ cells were present in the infarct area at 2 weeks postinfarction (Figure 4B). Macrophage density was not affected by free FGF1 treatment, although significantly fewer CD68+ cells were observed in the FGF1 coacervate group (Figure 4B). When analyzed, there was no significant difference in CD68+ cell density between the saline and free FGF1 groups (p > 0.05) (Figure 4C). However, administration of FGF1 coacervate greatly decreased the number of CD68+ cells in the infarct area (p < 0.01 relative to saline, p < 0.05 relative to free FGF1)
saline (p < 0.01) and free FGF1 (p < 0.05) groups, which suggests that FGF1 coacervate is more effective at preserving the cardomyocytes (Figure 3B). To evaluate the effect of FGF1 coacervate on cardiac structure, the infarct regions of the hearts were examined 6 weeks post-MI using H&E-stained sections (Figure 3C). In the saline control, the ventricle wall where the infarct occurred was very thin. The ventricle was drastically dilated compared to a normal heart and healthy tissue was replaced by granulation and scar tissues. Free FGF1 could reduce the infarct area, but most fibers in this region were damaged, and the tissue architecture was not drastically different from the saline group. In contrast, FGF1 coacervate was capable of preventing these damaging effects on the infarcted tissue as well as preserving normal ventricular size. As a result (Figure 3E), the FGF1 coacervate group showed significantly increased LV wall thickness in the infarct zone (274.8 ± 64.8 μm) compared to the saline (104.7 ± 31.8 μm, p < 0.01) and free FGF1 groups (153.4 ± 37.4 μm, p < 0.05). Overall, FGF1 coacervate reduced the infarct size (Figure 3D), prevented ventricular dilation and preserved cardiac fiber morphology. 3.4. FGF1 Coacervate Reduces Fibrosis and Inhibits Chronic Infiltration of Phagocytic Cells in the Infarcted Myocardium. To examine the effect of FGF1 coacervate on cardiac fibrosis, we performed Masson’s trichrome staining to observe collagen deposition in the infarcted myocardium 6 weeks post-MI. Consistent with the literature, there was significant deposition in the infarct zone post-MI (Figure F
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Figure 5. FGF1 coacervate achieves stable vasculature in the infarcted myocardium. (A, B) CD31 staining for vascular endothelial cells in the infarct zone. At 6 weeks, significantly fewer CD31 positive cells were detected in the saline control group compared to the FGF1 coacervate group. FGF1 coacervate induced angiogenesis and achieved a significantly higher vessel density than saline and free FGF1. (C, D) CD31 & KI67 staining for proliferation of endothelial cells in the infarct zone. Analysis showed significantly higher numbers of proliferating CD31+ cells in the FGF1 coacervate group relative to both saline and free FGF1 groups. (E-F) CD31 & α-SMA costainingto mark endothelial and mural cells in the infarct zone, which could further detect the formation of stable vessels including arterioles. FGF1 coacervate exhibited significantly higher numbers of SMA + cells relative to saline or free FGF1 groups. **P < 0.01, compared to saline, #p < 0.05, ##p < 0.01 compared to free FGF1. Data represent means ± SD.
(Figure 4C). In summary, these results showed that FGF1 coacervate could reduce macrophage infiltration post-MI in the infarcted tissue. 3.5. FGF1 Coacervate Promotes Long-Term Revascularization. MI causes an ischemic environment, and vascularization to restore blood flow to the infarcted region is very important for both tissue regeneration and a functional recovery. To investigate mature and stable vasculature formation in the infarct zone, immunofluorescent staining was performed using the endothelial cell CD31 marker (mostly located in microvasculature/capillaries) and α-SMA marker for vascular smooth muscle cells, which surround large blood vessels, at 6 weeks post-MI. In the infarct region (Figure 5A), there were very few CD31+ cells in the saline group, and the free FGF1 treatment group had higher CD31+ endothelial cell density compared to the saline control. However, the saline control was not different from the free FGF1 group after statistically analysis (p > 0.05). In contrast, the FGF1
coacervate treatment showed a much greater increase in CD31+ endothelial cell density than the saline (p < 0.01) and free FGF1 groups (p > 0.05) (Figure 5B). To further detect endothelial cell proliferation 6 weeks postinfarction, the number of CD31+/Ki67+ cells in the ischemic myocardium were measured. Immunostaining showed that FGF1 coacervatetreated hearts had many more CD31+/Ki67+ proliferating endothelial cells than the free FGF1 (p < 0.05) and saline (p < 0.01) control groups at the infarct areas (Figure 5C−D). Thi suggests that the FGF1 coacervate helped stimulate neovessel formation within the infarct and border zones. To further detect the stable vessel formation including arterioles, we costained CD31 and α-SMA to mark endothelial and mural cells together. As shown in our result, we could detect very few α-SMA+ cells after 6 weeks in the saline control (Figure 5E). The saline control had 42.67 ± 12.01 α-SMA+ cells per mm2 and had no significant difference from the free FGF1 group at 78.01 ± 15.87 per mm2 (p > 0.05). However, G
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Figure 6. FGF1 coacervate increases cardiac precursor cell proliferation. (A−C) Dual immunofluorescent detection and quantification of CD117+/ Ki67+ proliferating cardiac stem cells in infarct zone 6 weeks post-MI. **P < 0.01 compared with saline, #p < 0.05 compared with free FGF1. Data represent means ± SD.
Figure 7. Echocardiographic assessment suggests that FGF1 coacervate improves cardiac function. (A) Both at 2 and 6 weeks, the FGF1 coacervate group had the lowest EDA compared to the other groups, which was significant at 6 weeks. (B) Pairwise comparisons revealed that FGF1 coacervate treatment achieved a significantly lower ESA compared to saline and free FGF1 at 6 weeks. (C) FGF1 coacervate significantly increased FAC postMI over saline or free FGF1 treatment at 6 weeks. (D) FGF1 coacervate significantly increased EF post-MI relatively to saline at 6 weeks. (E) Representative M-mode 2-D echocardiography at 6 weeks post-MI showed the changes of cardiac contractility in each group. (F) Representative Bmode 2-D echocardiography at 6 weeks post-MI showed the changes in cardiac contractility in each group. *P < 0.05, **P < 0.01 compared with saline, #p < 0.05 compared with free FGF1. Data represent means ± SD.
H
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21.25 ± 5.61%, 25.2 ± 3.8% for the free FGF1 group, and 37.88 ± 4.27% for the FGF1 coacervate group (Figure 7C). Additionally, EDA and ESA values were significantly reduced in FGF1 coacervate group relative to saline (ESA p < 0.01, EDA p < 0.05) and free FGF1 (ESA and EDA p < 0.05). Consistent with FAC measurements, LVEF was greatly increased in the FGF1 coacervate group relative to saline(Figure 7D).Taken together, these data demonstrate that the FGF1 coacervate can potently improve cardiac function 6 weeks post-MI by improving contractility and preventing ventricular dilation.
the FGF1 coacervate treatment group had 163.1 ± 30.12 αSMA+ cells per mm2, which was much greater than the saline control (p < 0.01) and free FGF1 (p < 0.01) groups (Figure 5F). In summary, free FGF1 had no significant effect in promoting angiogenesis compared with the saline group. Using coacervate delivery, the same dose of FGF1 formed substantially more blood vessels in the injured myocardium. 3.6. FGF1 Coacervate Increases Proliferation of Cardiac Precursor Cell Populations in Vivo. To assess the effect of FGF1 coacervate on cardiac precursor cell density within the infarct area, we measured the number of c-kit+ cells in the myocardium. Quantitative analysis showed that while free FGF1 does not contain high numbers of c-kit+ cells relative to saline (p > 0.05), FGF1 coacervate shows a significantly higher density of c-kit+ cells as compared to saline (p < 0.01) and free FGF1 (p < 0.05) (Figure 6B). We also investigated the effect of FGF1 coacervate on cardiac precursor cell proliferation by measuring the amount of c-kit+/Ki67+ cells in the ischemic myocardium. Immunohistochemistry conducted 6 weeks postinfarction showed that FGF1 coacervate-treated hearts had much higher numbers of c-kit +/Ki67+ proliferating CSCs than the free FGF1 (p < 0.05) and saline control (p < 0.01) groups at the infarct areas (c-kit: stem cell growth factor receptor or CD117; Ki67: a cellular proliferation marker) (Figure 6A,C). Mast cells are also known to express this marker, so we performed toluidine blue staining to examine the effect of FGF1 on their infiltration. Staining at 6 weeks post-MI showed insignificant numbers of mast cells present in the infarct and border zone, regardless of treatment group (Figure S1). Therefore, c-kit+ cells were predominantly progenitor cells rather than inflammatory mast cells. This result supports the postulation that FGF1 coacervate promotes cardiac precursor cell survival and proliferation in hearts after infarction that may in part be responsible for augmented cardiac function. 3.7. Intramyocardial Delivery of FGF1 Coacervate Improved post-MI Cardiac Function. To assess the therapeutic efficacy of FGF1 coacervate on cardiac function M-mode and B-mode 2-D echocardiography was used at 2 weeks and 6 weeks post-MI to monitor the changes in cardiac contractility (Figure 7E, F). As shown in our results (Figure 7A, B), EDA and ESA of the saline group increased to 18.99 ± 5.8 mm2 and 13.45 ± 6.04 mm2, respectively at 2 weeks post-MI. Both are significantly higher than healthy animals (p < 0.05), confirming that MI led to ventricular dilation. The ESA and EDA values were similar in the saline, free FGF1 and FGF1 coacervate groups, which suggests that the difference in ventricular dilation between the groups at 2 weeks post-MI is not significant (p > 0.05). FAC revealed that myocardial contractility in the saline control was lower than the normal value (from 52.82 ± 4.36% to 29.29 ± 4.876%) (Figure 7C). The free FGF1 groups (36.36 ± 6.32%) showed no significant difference in comparison to the saline group (p > 0.05). The FAC of the FGF1 coacervate was higher than saline and free FGF1 at 2 weeks (42.95 ± 4.06%), although this difference was not statistically significant (p > 0.05). A similar trend was seen in LVEF measurements (Figure 7D). At 6 weeks, the FAC declined slightly for the saline, free FGF1 and FGF1 coacervate groups, however the FGF1 coacervate group could maintain improved cardiac function with the highest FAC of all three groups (p < 0.01 versus saline, p < 0.05 versus free FGF1), which was consistent with the result at 2 weeks. The FAC values of the saline group were
4. DISCUSSION Protein therapy using proangiogenic factors that are capable of promoting cardiac repair as well as regeneration have been widely investigated.32,33 To promote angiogenesis to revascularize ischemic myocardium, proangiogenic factors have been used with success in preclinical MI models.34,35 However, in clinical trials, these methods of treatment have been ineffective with generally disappointing results. For example, FGF1, VEGF, granulocyte macrophage colony stimulating factor, FGF-2, hepatocyte growth factor, and neuregulin-1 therapies could not significantly improve revascularization and myocardial function in Phase I and II clinical trials on a consistent basis, despite being tolerable and reasonably safe at the different doses used.3,33,36,37 One of the major obstacles of protein therapy utilizing exogenous growth factors or cytokines is the short half-life of the naked protein. In addition, systemically delivered growth factors have variable bioavailability in the target tissue largely due to the availability of local vasculature. These drawbacks have led to frequent administration of high growth factor doses in order to achieve therapeutic efficacy, but a systemic high quantity of the protein is potentially toxic. For example, a double-blind clinical trial showed that a high dose (50 ng/kg/min) of VEGF administered by intracoronary infusions in patients with myocardial ischemia can induce nitric oxide-mediated hypotension.38 Thus, to better improve the local bioavailability and efficacy of exogenous growth factors and decrease the dosage required for ischemic injury, a suitable controlled release system for consistent, localized delivery is urgently needed. Currently, there are many different types of vehicles for the controlled release of growth factors such as hydrogels, microand nanoparticles, and affinity-based delivery systems.35,39 However, these systems suffer from different shortcomings including large initial burst release, low protein-loading efficiency, reduced bioactivity of cargo proteins due to the use of organic solvents, and high cost. Traditional hydrogels keep the protein within the aqueous phase where hydrolysis can rapidly degrade them. Unfortunately, these gels are characterized by a burst release of protein due to the swelling nature of hydrogels after implantation and incomplete loading of growth factor within the hydrogel matrix.40,41 Thus, they are usually modified to better facilitate protein delivery. Micro- and nanoparticles are another commonly used vehicle; methods that fabricate micro- or nanoparticles require the use of organic solvents, which can denature the protein. The release of proteins from micro/nanoparticle-based systems follow firstorder release kinetics, which is difficult to alter for various applications.40,42 Thus, for efficient protein delivery, a vehicle that generates a high affinity between the vehicle and the delivered protein is warranted to increase the biological effect of growth factors. I
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cells in the FGF1 coacervate group, consistent with our in vitro findings using HUVECs and hCSCs. We also saw reduced myocardial fibrosis, which helps to mitigate the loss in contractile function observed in the control groups.47 Moreover, cardiomyocyte survival as a means of preserving contractile function, was greatly improved using our FGF1 coacervate delivery platform. In addition, FGF1is capable of activating cardioprotective signaling pathways in cardiomyocytes.48 Maintaining viable cardiac muscle is vital to better improve cardiac function post-MI, which has been shown in studies that attempt to stimulate the proliferation of cardiomyocytes, inhibit apoptosis, and stimulate the recruitment of cardiac progenitor cells to the heart.49−52 Moreover, out study demonstrated that FGF1 coacervate was capable of reducing macrophage infiltration in the infarcted region 2 weeks after MI. This reduction could be linked to indirect FGF1 inhibition of pro-inflammatory cytokines. Another possibility is that the improved angiogenesis and better preservedcardiac muscle we observed could reducee tissue damage, which would in turn be able to reduce inflammation. Previous studies suggest similar trends using FGF1 in other applications, ultimately showing reduced presence of CD68+ and inflammatory M1 macrophages.53 Thus, these benefits are manifested on a functional level through the improvements in cardiac contractility seen 6 weeks post-MI. Overall, FGF1 coacervate exhibited higher therapeutic potential when compared with the saline and free FGF1 groups. Large preclinical animal models could help to validate this therapeutic approach and eventually pave the way for implementation of this controlled delivery approach in ischemic heart disease patients. Additional studies to optimize FGF1 dosing is necessary to maximize the efficacy of this therapy. In addition, future investigated is warranted to better delineate FGF1 coacervate could serve as a treatment of other ischemic conditions such as myocardial reperfusion injuries and peripheral artery disease. However, delivering a single growth factor still demonstrates limited therapeutic effect. In our delivery system, heparin had a high affinity to a wide range of proteins, so it is possible that multiple proteins could be delivered using the coacervate generating a synergistic effect for stronger MI therapy.
In this study, we used a complex coacervate that composed of heparin and a polycation PEAD as the delivery vehicle. Heparin, the most negatively charged natural polymer in the body, can bind to more than 400 proteins and peptides such as extracellular matrix (ECM) proteins as well as growth factors and cytokines that are very important biologically. contains basic amino acid residues such as lysine and arginine in the heparin-binding domain of many such proteins are vital to the intermolecular interactions and downstream signaling.43 In our coacervate, electrostatic interactions in the complex immobilize heparin noncovalently, which can guarantee that its natural bioactivity will be preserved. Preservation of this bioactivity has been shown previously.44 The polycation PEAD is designed specifically for protein delivery. Heparin facilitates the interaction between FGF1 and its receptor, and the coacervate phase separation isolates the protein from its aqueous environment, preventing its rapid degradation through hydrolysis or proteolysis.25,44 It is a biodegradable polyester that exhibits minimal cytotoxicity regardless of its cationic charge.25 With regard to cardiac repair, Shh delivery with our coacervate is both cardioprotective and capable of stimulating healthy vascularization and heart function in rodents post-MI. Additionally, FGF-2 coacervate was capable of stimulating vascular stromal cell recruitment and long-term angiogenesis in the infarct’s border zone post-MI.27 FGF2 coacervate could also reduce peri-infarct inflammation and fibrosis, most likely by replenishing a functional vasculature. In our present study, we investigated whether FGF1 coacervate had potential cardioprotective effects. FGF1 was chosen due to its heparin-dependent bioactivity changes. Previous studies show that FGF1 activity is greatly increased when able to form a stable complex with heparin, which facilitates its interactions with receptors.24 There was an almost evenly homogeneous incorporation and distribution of FGF1 within coacervate droplets. Our delivery system not only had high loading efficiency for FGF1 (greater than 90%) but also exhibited low initial releases of approximately 15.1 ± 2.5% during the first 24 h with a relatively linear trend of FGF1 release for the following 28 days (approximately linear at 10 ng day−1). The in vitro assays indicated that FGF1 coacervate greatly improved the proliferation of HUVECs and hCSCs compared with free FGF1. The increase in cell proliferation in the presence of FGF1 is consistent with other studies,45,46 and our results indicate a further increase in FGF1 activity when used in the coacervate. In addition to the proliferation assay, we performed transwell assays to evaluate the chemotaxis of HUVEC and hCSC induced by the FGF1 coacervate. We found that FGF1 coacervate had greater chemotactic effects compared to the free FGF1 group in both HUVEC and hCSC. These results are evidence that FGF1 released from the coacervate is highly bioactive and capable of stimulating both proliferation and migration of HUVECs and hCSCs in vitro. In our animal study, we used a mouse MI model and demonstrated that the FGF1 coacervate system stimulated angiogenesis through the robust formation of mature and functional blood vessels in the infarct region. The the number of CD31 and α-SMA positive vessels increased significantly, which indicated the formation of new stable and mature vasculature. Additionally, CD31+/Ki67+ staining showed that endothelial cells continued to proliferate in the FGF1 coacervate group at 6 weeks post-MI. Conversely, we observed a large increase in the proliferative capacity of cardiac precursor
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CONCLUSION
Taken together, our present findings demonstrated that controlled release of FGF1 with the coacervate triggered both formation and mature stabilization of neovasculature, and it could therefore better imrpove cardiac function post-MI in a mouse model. The improvement of cardiac function was observed at 2 weeks and reached a higher level at 6 weeks postMI. This delivery system significantly improved myocardial function throughout the heart during post-MI remodeling by promoting angiogenesis, increased the formation of mature vasculature, promoting cardiomyocyte survival, inducing cardiac precursor cell proliferation, and decreasing collagen deposition and inflammation in the infarct zone. Large animal models could be used as a next step to validate the viability of the coacervate as a therapeutic approach in ischemic heart disease patients. J
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(6) Voors, A. A.; Belonje, A. M.; Zijlstra, F.; Hillege, H. L.; Anker, S. D.; Slart, R. H.; Tio, R. A.; van ’t Hof, A.; Jukema, J. W.; Peels, H. O.; Henriques, J. P.; Ten Berg, J. M.; Vos, J.; van Gilst, W. H.; van Veldhuisen, D. J. A single dose of erythropoietin in ST-elevation myocardial infarction. Eur. Heart J. 2010, 31 (21), 2593−2600. (7) Henry, T. D.; Annex, B. H.; McKendall, G. R.; Azrin, M. A.; Lopez, J. J.; Giordano, F. J.; Shah, P. K.; Willerson, J. T.; Benza, R. L.; Berman, D. S.; Gibson, C. M.; Bajamonde, A.; Rundle, A. C.; Fine, J.; McCluskey, E. R.; Investigators, V. The VIVA trial: Vascular endothelial growth factor in Ischemia for Vascular Angiogenesis. Circulation 2003, 107 (10), 1359−1365. (8) Simons, M.; Annex, B. H.; Laham, R. J.; Kleiman, N.; Henry, T.; Dauerman, H.; Udelson, J. E.; Gervino, E. V.; Pike, M.; Whitehouse, M. J.; Moon, T.; Chronos, N. A. Pharmacological treatment of coronary artery disease with recombinant fibroblast growth factor-2: double-blind, randomized, controlled clinical trial. Circulation 2002, 105 (7), 788−793. (9) Chu, H.; Johnson, N. R.; Mason, N. S.; Wang, Y. A [polycation:heparin] complex releases growth factors with enhanced bioactivity. J. Controlled Release 2011, 150 (2), 157−63. (10) Johnson, N. R.; Wang, Y. Controlled delivery of heparin-binding EGF-like growth factor yields fast and comprehensive wound healing. J. Controlled Release 2013, 166 (2), 124−9. (11) Lee, K. W.; Johnson, N. R.; Gao, J.; Wang, Y. Human progenitor cell recruitment via SDF-1alpha coacervate-laden PGS vascular grafts. Biomaterials 2013, 34 (38), 9877−85. (12) Chen, W. C.; Lee, B. G.; Park, D. W.; Kim, K.; Chu, H.; Kim, K.; Huard, J.; Wang, Y. Controlled dual delivery of fibroblast growth factor-2 and Interleukin-10 by heparin-based coacervate synergistically enhances ischemic heart repair. Biomaterials 2015, 72, 138−51. (13) Johnson, N. R.; Kruger, M.; Goetsch, K. P.; Zilla, P.; Bezuidenhout, D.; Wang, Y. D.; Davies, N. H. Coacervate Delivery of Growth Factors Combined with a Degradable Hydrogel Preserves Heart Function after Myocardial Infarction. ACS Biomater. Sci. Eng. 2015, 1 (9), 753−759. (14) Johnson, N. R.; Wang, Y. Controlled delivery of sonic hedgehog morphogen and its potential for cardiac repair. PLoS One 2013, 8 (5), e63075. (15) Awada, H. K.; Johnson, N. R.; Wang, Y. Sequential delivery of angiogenic growth factors improves revascularization and heart function after myocardial infarction. J. Controlled Release 2015, 207, 7−17. (16) Battegay, E. J. Angiogenesis: mechanistic insights, neovascular diseases, and therapeutic prospects. J. Mol. Med. 1995, 73 (7), 333− 346. (17) Bing, M.; Da-Sheng, Ch.; Zhao-Fan, X.; Dao-Feng, B.; Wei, L.; Zhi-Fang, C.; Qiang, W.; Jia, H.; Jia-Ke, C.; Chuan-An, S.; Yong-Hua, S.; Guo-An, Z.; Xiao-Hua, H. Randomized, multicenter, double-blind, and placebo-controlled trial using topical recombinant human acidic fibroblast growth factor for deep partial-thickness burns and skin graft donor site. Wound Repair Regen. 2007, 15 (6), 795−799. (18) Cuevas, P.; Reimers, D.; Carceller, F.; Martinez-Coso, V.; Redondo-Horcajo, M.; Saenz de Tejada, I.; Gimenez-Gallego, G. Fibroblast growth factor-1 prevents myocardial apoptosis triggered by ischemia reperfusion injury. Eur. J. Med. Res. 1997, 2 (11), 465−468. (19) Htun, P.; Ito, W. D.; Hoefer, I. E.; Schaper, J.; Schaper, W. Intramyocardial infusion of FGF-1 mimics ischemic preconditioning in pig myocardium. J. Mol. Cell. Cardiol. 1998, 30 (4), 867−77. (20) Buehler, A.; Martire, A.; Strohm, C.; Wolfram, S.; Fernandez, B.; Palmen, M.; Wehrens, X. H.; Doevendans, P. A.; Franz, W. M.; Schaper, W.; Zimmermann, R. Angiogenesis-independent cardioprotection in FGF-1 transgenic mice. Cardiovasc. Res. 2002, 55 (4), 768− 777. (21) Fernandez, B.; Buehler, A.; Wolfram, S.; Kostin, S.; Espanion, G.; Franz, W. M.; Niemann, H.; Doevendans, P. A.; Schaper, W.; Zimmermann, R. Transgenic myocardial overexpression of fibroblast growth factor-1 increases coronary artery density and branching. Circ. Res. 2000, 87 (3), 207−13.
ASSOCIATED CONTENT
S Supporting Information *
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsbiomaterials.6b00509. Toluidine blue staining of heart sections shows mast cells (if present) in deep violet; representative images of heart sections show that few if any mast cells are present in the infarct and border zone, regardless of treatment, indicating that FGF1 coacervate increases the presence of c-kit+ cells independent of inflammatory mast cell infiltration (PDF)
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AUTHOR INFORMATION
Corresponding Authors
*E-mail:
[email protected]. Tel.:+86-577-85773087. Fax: +86-577-85773087. *E-mail:
[email protected]. Tel: 1-412-624-7196. ORCID
Yadong Wang: 0000-0003-2067-382X Author Contributions §
D.W.L. and Y.H. contributed equally.
Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This work was supported by the American Heart Association (Grant 12EIA9020016 to Y.W.), the National Natural Science Foundation of China (81302775 to Z.W.), Science and Technology Project of Zhejiang Province (LY17H090017 to Z.W.).
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REFERENCES
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