RGDfK-Peptide Modified Alginate Scaffold for Cell Transplantation and Cardiac Neovascularization

Abstract

Cell implantation for tissue repair is a promising new therapeutic strategy. Although direct injection of cells into tissue is appealing, cell viability and retention are not very good. Cell engraftment and survival following implantation are dependent on a sufficient supply of oxygen and nutrients through functional microcirculation as well as a suitable local microenvironment for implanted cells. In this study, we describe the development of a porous, biocompatible, three-dimensional (3D) alginate scaffold covalently modified with the synthetic cyclic RGDfK (Arg-Gly-Asp-D-Phe-Lys) peptide. Cyclic RGDfK peptide is protease resistant, highly stable in aqueous solutions, and has high affinity for cellular integrins. Cyclic RGDfK-modified alginate scaffolds were generated using a novel silicone sheet sandwich technique in combination with freeze-gelation, resulting in highly porous nonimmunogenic scaffolds that promoted both human and rodent cell survival in vitro, and neoangiogenesis in vivo. Two months following implantation in abdominal rectus muscles in rats, cyclic RGDfK-modified scaffolds were fully populated by host cells, especially microvasculature without an overt immune response or fibrosis, whereas unmodified control scaffolds did not show cell ingrowth. Importantly, modified scaffolds that were seeded with human mesenchymal precursor cells and were patched to the epicardial surface of infarcted myocardium induced myocardial neoangiogenesis and significantly improved cardiac function. In summary, purified cyclic RGDfK peptide-modified 3D alginate scaffolds are biocompatible and nonimmunogenic, enhance cell viability, promote angiogenesis, and may be used as a means to deliver cells to myocardial infarct areas to improve neovascularization and cardiac function.

Introduction

Cell therapy is emerging as a promising strategy for tissue repair in a variety of diseases, including heart disease (reviewed in Pavo et al.¹). Cells can be (1) delivered directly into damaged tissue by injection² at orthotopic (heart and liver) or heterotopic sites (parathyroids and islets), (2) administered intravenously,³ in combination with biocompatible materials,⁴ or (3) prepared for implantation following in vitro culture, for example by using cell sheet monolayers⁵ or biodegradable scaffolds.⁶

Direct cell delivery by injection is an attractive method due to its minimally invasive character. This approach, however, has been frequently hampered by a lack of long-term cell survival due to insufficient cell oxygenation, lack of sufficient nutrients, and inadequacy of a suitable microenvironment, which usually consists of neighboring cells and the extracellular matrix (ECM), or stem-cell niche.^7,8 It is also suspected that delivery of cellular inoculate by needle injection may lead to an uneven cell distribution or formation of cell “islands” within tissue, further complicating both nutrient and oxygen delivery to individual cells, thereby interfering with therapeutic effects.⁹

Numerous experimental tissue engineering approaches to tissue repair and regeneration have been previously attempted with mixed success. These have included for example, grafting of monolayered or multilayered cell sheets cultured ex vivo⁵ or cell-seeded scaffolds¹⁰ in animal models of tissue degenerative diseases. Clinically, a bioengineered stem cell-seeded trachea was successfully implanted in a patient with tracheal cancer.¹¹ The application of various tissue engineering methods generally led to more efficient and better repair of damaged tissue when compared to direct needle injection-based therapies.¹²

Biomaterial carriers have a variety of biological properties, which are either beneficial or potentially harmful to viable cells. Several have been previously used in (pre)clinical applications. Some of these include, for example, PGA¹³ and collagen and hyaluronic acid.¹⁴ We chose alginate as a basic biomaterial because it has several advantages over those previously mentioned: it is plant derived, and therefore avoids the risk of pathogen (i.e., prion) transfer, it is biologically inert when sufficiently purified, and readily available in large quantities. Purified unmodified alginate has already been approved previously for several clinical applications, especially for wound dressing.¹⁵

Recent studies showed that intracoronary alginate injection was safe and prevented left ventricular (LV) enlargement after myocardial infarction (MI) in swine,¹⁶ and clinical trials are being conducted to investigate effects of an aqueous mixture of sodium alginate and calcium gluconate for the prevention of remodeling of the ventricle and congestive heart failure after acute MI (ClinicalTrials.gov No. NCT01226563). Another advantage of alginate is its carboxyl groups, which permit covalent (i.e., irreversible) modifications with biologically active peptides that contain free amino groups, by carbodiimide crosslinking.

We previously developed an alginate purification protocol that results in a highly purified, biocompatible poly-mannuronic acid containing alginate.¹⁷ Alginate purification for cell transplantation application is of paramount importance for survival of transplanted cells, since raw alginate is contaminated with mitogens such as proteins, DNA, RNA, and lipopolysaccharides.

To enhance the biologically functional properties of alginate, we hypothesized that modification of purified alginate with covalently attached cyclic RGDfK (Arg-Gly-Asp-D-Phe-Lys) peptides will lead to more effective cell survival compared to controls, since cyclic RGDfK peptides are protease resistant and stable in aqueous solution.¹⁸ RGD is an abundant peptide sequence found in the ECM as part of vitronectin, fibronectin, and collagen.¹⁹ The RGD peptide sequence binds to cell surface integrins α_vβ₃ and α₅β₁, where it signals into the cell, thereby promoting cell adhesion and formation of cell viability factors.²⁰

More specifically, cyclic RGDfK is a synthetic RGD peptide that has been shown to enhance cell adhesion and cell survival after covalent immobilization on two-dimensional (2D) surfaces.²¹ Therefore, we hypothesized that a purified three-dimensional (3D) alginate scaffold dressed with cyclic RGDfK peptide would enhance cell retention and survival within the scaffold.

Experimental Section

Materials

Solutions were prepared using purified water from an in-house Hydro Picopure (Millipore, Temecula, CA) purification system. Molecular biology grade ddH₂O (No. SH30529.02), EDC (1-Ethyl-3-[3-dimethylaminopropyl] carbodiimide hydrochloride, No. 22980), sulfo-NHS (N-hydroxysulfosuccinimide, No. 24510), hydroxylamine (No. 26103), MES buffer (No. 28390), dialysis membranes (3.5 kDa membrane, SnakeSkin, No. 88244), EDTA (No. 17892), and TrypLE Select (No. 12604013) were purchased from Thermo Scientific (Rockford, IL). All glassware was cleaned and detoxified by 24-h treatment with 0.5% hypochlorite followed by washing and baking for at least 120 min at 240°C. Pyrex^® glass plates (100 × 15 mm) were purchased from Corning (New York, NY; No. 3160-101).

Low molecular weight (MW) sodium alginate (No. A0682) obtained from brown algae consisting of straight-chain, hydrophilic, colloidal polyuronic acid, which is composed primarily of anhydro-β-d-mannuronic acid residues with 1→4 linkage, calcium chloride (No. C2661), bovine serum albumin (BSA; cell culture grade, No. A9418), collagenase (≥125 collagen digestion U/mg, No. C5138), gelatin type A (No. G9136), and DNase I (grade II, from bovine pancreas, No. 10104159001) were purchased from Sigma-Aldrich (St. Louis, MO).

Cyclic RGDfK peptide (No. PCI-3661-PI) and cyclic RADfK (negative control peptide; No. PCI-3883-PI) were purchased from Peptides International, Inc. (Louisville, KY). Nonreinforced vulcanized gloss/gloss silicone sheets (hardness 40 durometer, thickness 750 μm) were purchased from Specialty Manufacturing, Inc. (Saginaw, MI). Ten millimeters Acu-Punch skin punchers were purchased from Acuderm, Inc. (Fort Lauderdale, FL). Alpha minimum essential medium (α-MEM), fetal calf serum (FCS), l-glutamine (GlutaMax^®), and recombinant human vascular endothelial growth factor (VEGF-A, No. PHG0145) were purchased from Invitrogen (Carlsbad, CA). Human platelet-derived growth factor-bb (PDGF-BB No. GF018) was purchased from Chemicon/Millipore (Billerica, MA).

Human mesenchymal precursor cells (hMPCs) were a kind gift of Mesoblast, Ltd. (New York, NY). Primocin was purchased from InvivoGen (San Diego, CA). Osteogenic, chondrogenic, and adipogenic MSC differentiation kits (Nos. PT-3002, PT-3003, PT-3004) were purchased from Lonza (Walkersville, MD). WST-1 was purchased from Roche Applied Science (Indianapolis, IN). Factor VIII-related antigen monoclonal antibodies (mAb; No. A0082) were purchased from Dako (Carpinteria, CA). Rat α-smooth muscle actin (αSMA) mAb was purchased from Biogenex (Fremont, CA). mAb against CD163/ED-2, a rat macrophage marker, was purchased from Santa Cruz Biotechnology (Dallas, TX). The Vectastain ABC kit was purchased from Vector Laboratories (Burlingame, CA).

Sodium alginate purification

We previously described our alginate purification protocol in detail.¹⁷ In brief, 1.5% sodium alginate solution was treated with activated charcoal before serial polyvinylidene difluoride (PVDF) membrane filtration in 10 mM phosphate buffer at pH 5.5, followed by dialysis and ethanol precipitation. Resulting precipitate was freeze-dried and redissolved in molecular biology grade ddH₂O at 2% w/w before further processing.

Study 1: ex vivo studies

Cyclic RGDfK peptide conjugation, fabrication, and characterization of cyclic RGDfK-modified alginate scaffolds

We applied carbodiimide chemistry based on EDC and sulfo-NHS in MES buffer using a modified protocol as previously described^22,23 (Fig. 1a). Cyclic RGD peptide RGDfK and negative control peptide cyclic RADfK were covalently attached to alginate according to protocol with slight modifications. Cyclic RADfK peptide differs from cyclic RGDfK peptide, in that one glycine (G) is replaced with alanine (A), abolishing its effects through lack of binding capacity to integrin receptors on cell surfaces.²⁴

FIG. 1.

Schematic reaction scheme of covalent coupling of alginate -COOH groups (a.A) to cyclic RGDfK peptide -NH₃⁺ groups (a.B) to yield cyclic RGDfK-coupled alginate (a.C). Sulfo-NHS intermediary activation of alginate was used to increase efficiency of coupling reaction (adapted from Hermanson²³). hMPC adhesion to tissue culture plates and subsequent differentiation into osteoblasts, adipocytes, and mature chondrocytes was shown by Alizarin Red (b), Oil Red O (c), and Alcian Blue (d) stains, respectively. Cyclic RGDfK incorporation was evaluated using 2D cell adhesion assays with hMPCs. Unmodified alginate showed no cell adhesion (e), with rounded cells on the alginate surface. Modified alginate showed cell adhesion with a nerve cell-like phenotype (f) comparable to tissue culture plate-adherent hMPCs (d). 2D, two dimensional; hMPC, human mesenchymal precursor cell; NHS, N-hydroxysulfosuccinimide; RGDfK, Arg-Gly-Asp-D-Phe-Lys.

Briefly, 8.82, 17.65, and 35.30 mM peptide (equivalent to 5, 10, and 20 mg cyclic RGDfK) per gram alginate (estimated MW alginate = 200 D/mannuronic acid residue) in a 1% solution was incubated for 20 h at room temperature with EDC and sulfo-NHS to achieve theoretical activation of 10% of total mannuronic acid monomers. The reaction was subsequently quenched with hydroxylamine and the solution was dialyzed for 5 days against decreasing NaCl concentrations; it was then lyophilized at 0.2 torr, redissolved in molecular biology grade ddH₂O at 2% w/w, and sterile filtered.

Cell adhesion to 2D cyclic RGDfK-modified alginate scaffolds

To determine covalent cyclic RGDfK modification, solid 2D scaffolds were produced as previously described. In brief, cyclic RGDfK-modified alginate was placed in cell culture insert top wells and allowed to solidify with 100 mM CaCl₂ in bottom wells at 4°C overnight. Following solidification, scaffolds were washed thrice in molecular biology grade ddH₂O to remove excess CaCl₂ and placed in a full medium consisting of α-MEM supplemented with 10% FCS, 0.5% BSA, 0.1 μM ascorbic acid, 0.05 μM 2-mercaptoethanol, and 0.2% Primocin. For each batch of cyclic RGDfK-modified alginate, 50,000 hMPCs were seeded on 2D scaffold surface and allowed to adhere for 24–48 h in a humidified incubator at 37°C, 5% CO₂. The degree of adhesion was compared to that found in controls using unmodified alginate scaffolds (Fig. 1e, f).

Fabrication of 3D freeze-gelled cyclic RGDfK alginate scaffolds

Rectangular strips (15 × 40 mm) of silicone sheeting (0.75 mm thickness) were cut. Three circular wells with a diameter of 10 mm were punched in each strip (Fig. 2a.i). Circular silicone sheets (90 mm diameter) (Fig. 2a.ii) were cut out and transferred to sterile glass petri dishes (100 × 15 mm) (Fig. 2a.iii). Punched silicone strips were placed on silicone sheets, while assuring that no air remained between sheets. Subsequently, 56 μL cyclic RGDfK alginate solution per well was cast in the circular wells and carefully layered with a third rectangular silicone strip (15 × 40 mm), making sure that the alginate solution remained sandwiched between the silicone layers in the wells.

FIG. 2.

One hundred millimeter glass dishes (a.ii) were used with a 90 mm bottom layer of silicone sheeting (a.iii). Three silicone strips (a.i) with 3 × 10 mm circular wells each were layered on primary sheet (a.iii). Resulting wells were filled with modified alginate solution (56 μL) before freezing at −20°C. Following freeze-gelation, scaffold seeding and culture are shown in (b–d). Black arrowheads indicate scaffolds in wells. Scanning electron microscopy images show sponge-like scaffold following freeze-gelation in (e) (magnification 1 × ) and scaffold surface (f) (magnification 1000 × ). Note the highly porous structure of freeze-gelled scaffold without need for lyophilization. hMPC-seeded light microscopic scaffolds are shown in (g) (40 × ) and (h) (200 × ). hMPC viability data (i) inside scaffolds following 1 week culture in vitro are shown. Cyclic RGDfK modification of alginate significantly enhanced hMPC viability compared to control (cyclic RADfK-modified or RADfK-unmodified scaffolds) at 20 mg/g. Student's t-test, two-way, equal variance. Bars represent mean ± SEM. *p < 0.05. SEM, standard error of the mean.

Alginate-containing plates were subsequently frozen at −20°C overnight. Then, 25 mL 1.1% CaCl₂ in 70% ethanol/molecular biology grade ddH₂O solution precooled at −20°C was added to the petri dishes, and top strips were removed. This exposed frozen alginate scaffolds to ethanol/molecular biology grade ddH₂O and the plates were placed back at −20°C for at least 4 h. By using this freeze-gelation approach as previously described,²⁵ we were able to generate highly porous scaffolds, without the use of lyophilization, by simultaneous liquefaction of the frozen cyclic RGDfK alginate solution, while solidifying it by binding calcium ions.

Following freeze-gelation, plates were warmed to room temperature and sterilely washed thrice in molecular biology grade ddH₂O; 10 mm diameter circular scaffolds were removed from the plate, air dried, and stored dry at −20°C until further use.

Scanning electron microscopy

Scaffolds were dehydrated through an ethanol gradient, then dried using a Bal-Tec Critical Point Dryer (Bal-Tec, Balzers, Liechtenstein), sputter-coated with 15 nm of Gold-Palladium with a Cressington Manual Sputter Coater (Cressington, Watford, United Kingdom), and imaged with a Hitachi 4700 scanning electron microscope (Hitachi Medical Systems, Twinsburg, OH) (Fig. 2b–d).

Cell culture

hMPCs were isolated from whole human bone marrow by magnetic bead separation using a novel mAb against STRO-3 as previously described.²⁶ This antibody isolates an hMPC population expressing Stro-1, CD44, CD90, and CD106 by flow cytometric analysis. First, we confirmed the ability of these cells to differentiate into osteoblasts, adipocytes, and mature chondrocytes according to the manufacturer's instructions,²⁷ followed by staining with Alizarin Red S, Oil Red O, and Alcian Blue, respectively.

Next, hMPCs were allowed to proliferate in T75 flasks with the full medium. For passage or cell usage, cells were lifted from flasks after incubation with 0.1% collagenase for 30 min followed by TrypLe Select treatment for 5 min, and resuspended in phosphate-buffered saline (PBS) with 10% FCS and DNase I (1 mg/mL) to prevent cell clumping. Passage 2–5 hMPCs were used for all studies. Neonatal rat cardiomyocyte and neonatal rat cardiac fibroblast viability in vitro were assessed in separate experiments (Supplementary Data; Supplementary Data are available online at www.liebertpub.com/tea).

Scaffold seeding

Dry scaffolds were placed in 35 mm tissue culture-treated dishes (Fig. 2b). Cells were suspended at 10⁷/mL followed by application of 1 × 10⁶ cells (100 μL) in the full medium directly onto the dry scaffold (Fig. 2c). Due to highly porous and hydroscopic nature of scaffolds, cells were quickly absorbed and retained inside the scaffold pores. 2.9 mL of full medium was subsequently added to each scaffold containing dish and they were placed on a shaker for 30 min at 20 rpm to complete seeding. Dishes with scaffolds were subsequently transferred to a humidified incubator at 37°C and 5% CO₂ until further use (Fig. 2d).

Viability studies

Scaffolds seeded with hMPCs were cultured in full α-MEM and maintained for 7 days at 37°C and 5% CO₂. The medium was replaced daily. After culture, scaffolds were removed from the medium and placed in citrate/EDTA buffer to liquefy alginate by Ca²⁺ chelation and permit collection of cells for viability studies. Removal of calcium by chelation through the citrate/EDTA buffer resulted in rapid degradation of the calcium alginate 3D structure.

Subsequently, cells were washed in 1 mL PBS/10% FCS solution. After washing, cells were resuspended in collagenase 0.1% solution for 30 min at 37°C followed by treatment in TrypLe Selekt (Invitrogen) to facilitate cell disaggregation. Cells were then washed in 1 mL DNase I (1 mg/mL) supplemented with 10% FCS. Cell viability was assessed by counting live and dead cells using trypan blue exclusion.²⁸ Studies were done thrice in duplicate. Additional viability studies using rodent cells were conducted (Supplementary Data).

Study 2: scaffolds implanted into the abdominal muscles

Intramuscular in vivo application of cyclic RGDfK-modified alginate scaffolds

Proof-of-concept studies were performed by intramuscular scaffold implantation as previously described.⁶ All animal studies were reviewed and approved by the Columbia University Institutional Animal Care and Use Committee (IACUC).

Male Lewis rats weighing between 200 and 250 g were obtained from Harlan Sprague Dawley, Inc. (Indianapolis, IN). Rats were anesthetized with isoflurane and after incising the abdominal skin, the rectus muscle was dissected to the peritoneum. A newly created pocket accommodated a 10 mm scaffold. Based on our previous studies,⁶ scaffolds were pretreated with recombinant VEGF-A 100 ng/mL and recombinant PDGF-BB 100 ng/mL to induce a vascular response and then implanted in the newly created space before reapproximating the muscle over the space and closing the incision in separate layers.

Sixty days after implantation, muscle plus scaffolds were harvested in toto and fixed in 2% paraformaldehyde for further analyses. In some animals, muscle was sectioned through the scaffold to optically inspect scaffold material at the time of harvest. Paraffin tissue sections were stained with Gill's hematoxylin/eosin (H&E) and factor VIII-related antigen mAb, diluted 1 in 200, according to the manufacturer's instructions, to examine vascularity of the tissue. Vessels within five separate high-power fields at 400× magnification were counted.

Study 3: scaffolds to repair MI

Epicardial in vivo application of cyclic RGDfK-modified alginate scaffolds

Epicardial scaffold implantation was performed as previously described.²⁹ Rowett (rnu/rnu) athymic nude rats (225–250 g; Harlan Sprague Dawley, Inc.) were used according to an IACUC-approved protocol. Rats were anesthetized with inhaled isoflurane (2–3%), endotracheally intubated, and mechanically ventilated. The heart was exposed through a left thoracotomy. Following opening of the pericardium, the left anterior descending artery (LAD) was ligated with 7–0 Prolene (Ethicon, Somerville, NJ) suture 2–3 mm below the edge of the left atrium, and 48 h later, an engineered scaffold was placed onto the epicardial surface of the infarcted area. The pericardium was not closed thereafter.

Three types of engineered scaffolds were implanted: (1) cyclic RGDfK-modified scaffolds without hMPCs (n = 11); (2) cyclic RGDfK-modified scaffolds with 1 × 10⁶ hMPCs (n = 14); and (3) cyclic RGDfK-modified scaffolds with 3 × 10⁶ hMPCs (n = 6). We did not use unmodified scaffolds in in vivo heart failure studies, since hMPC viability in such scaffolds was shown to be limited. We further opted to not pretreat epicardial hMPC-seeded scaffolds with VEGF-A/PDGF-BB, as we did in the intramuscular studies, since our data showed that hMPCs secrete several proangiogenic factors such as VEGF and MCP-1 that induce vasculogenesis in infarcted myocardium.²⁶ Seven days after scaffold implantation, hearts were harvested, sectioned at the mid-LAD level, imaged, and fixed in 2% paraformaldehyde for further histological analyses.

Effects of scaffold implantation on LV function

Rats were scanned in a parasternal short-axis view under inhaled isoflurane (2%) anesthesia, with 2D and M-mode echocardiographic imaging using a Sonos 5500 System (Philips Medical Systems, Framingham, MA) and a broadband high-frequency (12 MHz) transducer (S12). Left ventricular fractional shortening (LVFS) was used to assess cardiac function based on its reported reproducibility and lower variability compared to other cardiac parameters following MI using LAD ligation in rats.^30,31

LVFS was calculated using the formula LVFS (%) = (LV diastolic diameter [LVDd] − LV systolic diameter [LVDs])/LVDd × 100. The maximum and minimum values of LVDd and LVDs in one cardiac cycle were obtained after tracing the endocardial LV border. To address echocardiographic data variability, interquartile range (IQR) analysis was performed. Outliers were defined as observations that fall below Q1 − 1.5 × IQR or above Q3 + 1.5 × IQR.

Tissue analyses

Histological analyses

Sections were stained with mAbs directed against rat factor VIII-like protein, rat αSMA, and CD163/ED-2, a rat macrophage marker. Arterioles were differentiated from large capillaries by the presence of a smooth muscle layer, identified by staining with a mAb against αSMA. Staining was performed using immunoperoxidase, an avidin–biotin blocking kit, a rat-absorbed biotinylated anti-mouse IgG, and a peroxidase conjugate. Histological sections from three or four animals per group were analyzed by a blinded investigator at 40× magnification by image processing (using 24–48 fields of view) to determine (1) factor VIII—positive vessels (n = 4 animals per group) and (2) αSMA-positive vessels (n = 4 animals per group).

Statistical analysis

Groups were compared by Student's t-test or Welch's analysis of variance with Bonferroni correction, as indicated in figure legends. A p-value <0.05 was considered statistically significant. Bar graphs represent mean value ± standard error of the mean (SEM) or, in case of box plots for echocardiographic analysis, medians with IQRs ± minimum and maximum values. Calculations were performed using Microsoft Excel with the Real Statistics Add-In (www.real-statistics.com).

Results

Alginate purification

We used a novel purification protocol based on a modified protocol by Prokop and Wang with additional steps to improve removal of contaminants.³² The approach was based on our in-house developed protocol using hydrophobic PVDF membrane filtration. This method increased the final purity of poly-mannuronic acid threefold to fivefold compared to commercially available pharmaceutical grade poly-mannuronic/guluronic acid.¹⁷

Characterization of STRO-3 selected, culture-expanded hMPCs

hMPCs were immunoselected from fresh bone marrow aspirates as previously described.²⁶ Flow cytometric analysis of passage 5 hMPCs indicated that ∼95% of the population was positive for the Stro-1 antigen, which has been shown to be an important marker of clonogenic bone marrow stroma. Stro-1-positive hMPCs coexpressed a number of common MSC surface markers, namely CD13, CD29, CD44, CD90, and CD146, but were negative for the endothelial precursor marker, CD34, were plastic adherent, and displayed a fibroblastic morphology, which was retained following multiple passages (up to 5) (data not shown).

These cells were confirmed to be clonogenic in fibroblastic colony-forming unit assays. Moreover, culture-expanded hMPCs were multipotent, as evidenced by their ability to differentiate along osteogenic, adipogenic, and chondrogenic lineages in the presence of previously defined inductive stimuli (Fig. 1b–d). The isolated hMPCs secreted several chemokines and growth factors, notably IL-6, MCP-1, and VEGF (data not shown).

Cyclic RGDfK conjugation and effect on hMPC adhesion

Following covalent cyclic RGDfK modification, 2D cyclic RGDfK-modified scaffolds were produced and then evaluated for hMPC adhesion as determined by cell spreading. The spreading of hMPCs seeded on cyclic RGDfk-modified alginate scaffolds (Fig. 1f) was comparable to spreading on 2D tissue culture plates (Fig. 1b–d). Unmodified scaffolds did not show any hMPC adhesion, but instead, hMPCs rounded up indicating no adhesion (Fig. 1e).

Freeze-gelation and scaffold structure

Scanning electron microscopy photographs of the highly porous structure of scaffolds produced using freeze-gelation are shown in Figure 2e and f. Figure 2e shows 1× magnification, whereas Figure 2f shows 1000 × magnification. As opposed to lyophilization, freeze-gelation did not result in the formation of an impenetrable surface, thus allowing retention of the highly porous structure throughout the scaffold (images not shown). Pore sizes ranged between 40 and 100 μm, as estimated using scanning electron microscopy imaging.

Ex vivo studies

hMPC retention and viability using 3D freeze-gelled scaffolds

Interestingly, retention of cells inside scaffolds was significantly higher in cyclic RGDfK-modified scaffolds than in unmodified scaffolds (data not shown). Figure 2g and h show light microscopic images of hMPCs seeded inside cyclic RGDfK-modified scaffolds at 40 × and 200× modification, respectively. After seeding, there was a significant difference in cell viability between those being placed on the cyclic RGDfK-modified scaffolds and those being placed on unmodified scaffolds (Fig. 2i and Supplementary Fig. S1).

Furthermore, the higher the cyclic RGDfK concentration, the higher the cell viability or cell number (Supplementary Fig. S1 and Fig. 2i). For hMPCs, 20 mg/g cyclic RGDfK was associated with 38.3% ± 2.7% viable cells versus 17.9% ± 2.1% and 14.1% ± 1.1% at 20 mg/g negative control cRADfK peptide and 0 mg/g cyclic RGDfK after 7 days of culture, respectively (p < 0.05) (Fig. 2i). Additional data can be found in the Supplementary Data.

In vivo studies

Scaffolds implanted into the abdominal muscles

Preliminary safety studies showed that all animals tolerated intramuscular scaffold implantation. There were no clinical signs of distress or infection/inflammation in general or at the implantation site. Sixty days following implantation, abdominal tissues were harvested. Scaffolds could be identified optically in situ. H&E sections showed no scaffold cellularization of unmodified scaffolds (Fig. 3a). Scaffolds modified with cyclic RGDfK or cyclic RGDfK +100 ng VEGF-A + 100 ng PDGF-BB showed robust cellularization and vascularization (Fig. 3b, c), which was confirmed by factor VIII-like protein staining (Fig. 3d).

FIG. 3.

Alginate scaffolds with and without cyclic RGDfK peptide modification. Note virtual absence of cellularization of unmodified scaffold (a) (H&E, 40 × ). Note scaffold cellularization throughout scaffold material of cyclic RGDfK-modified scaffold (b) (H&E, 40 × ). Note vascularization and functional blood vessels filled with erythrocytes inside cyclic RGDfK-modified scaffolds (arrows, c) (H&E, 200 × ). Cyclic RGDfK-modified scaffold plus PDGF-BB and VEGF-A showed abundant vessel formation inside scaffold by von Willebrand factor-like protein staining (arrows, d) (40 × ). H&E, hematoxylin/eosin; PDGF-BB, platelet-derived growth factor-bb; VEGF, vascular endothelial growth factor.

Epicardial scaffold implantation in rats with MI

Epicardial scaffolds were also well tolerated. Scaffolds remained in situ for 7 days (Fig. 4a–d). Following explantation, tissue analyses showed minimal scaffold fibrosis (Fig. 4e) and no CD163⁺/ED-2 macrophage or fibroblast infiltration (Fig. 4f), indicating absence of foreign body responses. Scaffolds seeded with 1 × 10⁶ hMPCs showed significantly more vascularization at the infarct border zone (Fig. 5) than scaffolds without cells.

FIG. 4.

Scaffold application to epicardial surface. Scaffolds±cells were applied 48 h following MI through thoracotomy (a). Scaffolds sufficiently adhered to myocardial surface without sutures (b, d, arrows) and remained in place for the duration of the study (c, arrows). Hearts were harvested for histology 1 week after transplantation (c, d). Masson's trichrome staining for fibrosis (e) showed Scaf cellularization with minimal fibrosis in the scaffold, but significant fibrosis in the myocardium (blue). (f) Shows CD163/ED-2 staining for rat macrophages. There was no evidence of foreign body reaction against the scaffold material at 1 week following implantation. LVw, left ventricular wall; MI, myocardial infarction; Scaf, scaffold.

FIG. 5.

Numbers of erythrocyte-filled blood vessels in infarct zone, border zone, and scaffold shown in (a). Blood vessels in the border zone of the MI showed significantly increased numbers in animals treated with scaffolds +1 × 10⁶ hMPCs compared to animals that were treated with scaffolds only or with scaffolds +3 × 10⁶ hMPCs. (b, c) Show scaffold on epicardial surface using Masson's trichrome (b) and αSMA staining (c) at 40 × . (d) Shows Masson's trichrome stain with erythrocyte-filled arteriole (arrow; blue is interstitial connective tissue) inside scaffold at 400 × ; (e) shows αSMA-positive vasculature (brown) inside scaffold at 400 × (arrows). Scaf designates scaffold. LVw/I is left ventricular wall with infarction. Welch's ANOVA. Bars represent means ± SEM. *p < 0.05. αSMA, α-smooth muscle actin; ANOVA, analysis of variance.

Surprisingly, scaffolds seeded with a higher dose of hMPCs (3 × 10⁶) did not show increased vascularization, but rather a trend to decreased vascularization at either border or infarct zone, perhaps on the basis of cell “crowding,” resulting in lesser viability due to lack of nutrients when using a higher concentration of cells per scaffold (Fig. 5a).

Epicardial scaffolds and effect on cardiac function

Following epicardial scaffold application in the 1 × 10⁶ hMPC group, we found a slight, but significant positive effect of 4.7% on LVFS when day 0 (scaffold implantation) means were compared to means on day 7 (Fig. 6a, white diamonds). No effect on LVFS was observed in the group with unseeded scaffolds, nor in the group with scaffolds seeded with 3 × 10⁶ hMPCs, compared to saline injection. Data for each individual animal are shown (Fig. 6b–e) and summarized in Figure 6a.

FIG. 6.

LVFS by echocardiography showed significant increase in cardiac function 1 week following epicardial application of cyclic RGDfK scaffolds seeded with 1 × 10⁶ hMPCs (d) compared to saline injection (b). This effect was not observed using scaffolds without cells (c) or scaffolds seeded with 3 × 10⁶ hMPCs (e). Mean (white diamonds) and median change in LVFS (white horizontal lines) are summarized in (a). For means, white error bars show SEM. Black bars represent IQR, black whiskers represent minimum and maximum values. Horizontal white lines inside bars show medians. Welch's ANOVA with Bonferroni correction. *p = 0.047. cRGDfK, cyclic RGDfK; IQR, interquartile range; LVFS, left ventricular fractional shortening.

When comparing medians on day 0 versus 7, decrease in fractional shortening was −22.58% in saline control group, −24.54% in scaffold without cells group, −11.79% in 1 × 10⁶ hMPC group, and −17.14% in 3 × 10⁶ hMPC group (Fig. 6a, white lines).

For medians, IQR assessment was performed to address outlier and variability in the LVFS change data. For all groups, data fell between Q1 − 1.5 × IQR and Q3 + 1.5 × IQR (ranges −71.8% to 19.2% for saline, −80.2% to 57.2% for control scaffold, −101.0% to 106.4% for 1 × 10⁶ hMPC group, and −64.6% to 25.0% for 3 × 10⁶ hMPC group).

Discussion

We used a patented method for sodium alginate purification, which resulted in the production of nonimmunogenic material.¹⁷ The purified alginate could be covalently modified with cyclic RGDfK peptides and we were able to produce 3D scaffolds of this material. Scaffolds produced from cyclic RGDfK peptide-modified alginate significantly improved cell viability of a variety of cells, when compared with unmodified or negative control cyclic RADfK-modified alginate. Moreover, hMPC-seeded cyclic RGDfK peptide-modified scaffolds showed robust cellularization and neovascularization upon implantation in the abdominal wall or pericardially. No adverse immune response was observed.

In the heart, our results show that implantation of cyclic RGDfK-modified alginate scaffolds seeded with 1 × 10⁶ hMPCs resulted in a mean 4.7% increase in LVFS compared to saline controls (Fig. 6a, white diamonds). Regarding median changes, fractional shortening decreased ∼50% less in the 1 × 10⁶ hMPC scaffold-treated group compared to saline control or scaffold without cells groups: −22.58% in saline control group, −24.54% for scaffold without cells group, −11.79% for 1 × 10⁶ hMPC scaffold group, and −17.14% in 3 × 10⁶ hMPC scaffold group (Fig. 6a, white lines). These results suggest a protective effect of epicardial application of hMPC-seeded cyclic RGDfK-modified alginate scaffolds on deterioration of cardiac function following acute MI.

Of note, we did not test unmodified scaffolds in MI studies, since our preliminary data showed that unmodified scaffolds did not become vascularized and exerted a deleterious effect on hMPC viability. The enhanced in vitro viability of cells in cyclic RGDfK peptide-modified alginate is likely due to enhanced stability of cyclic RGDfK peptide, which is less susceptible to proteolytic degradation than linear Gly-Arg-Gly-Asp-Ser-Pro (GRGDSP) peptide.¹⁸ This would make cyclic RGDfK peptide more readily available to bind to cellular integrins on the cell surface.

In addition, cyclic RGDfK peptides bind to cellular integrin receptors with high affinity,²¹ which may further enhance signaling to maintain cell survival. Further optimization of cell survival with these peptide modifications, for instance by spacing the cyclic RGDfK moieties³³ or using multivalent cyclic RGDfK derivatives,³⁴ may be required.

Cell death inside scaffolds may contribute to deleterious effects on tissue regeneration and therefore be counterproductive.³⁵ Since scaffolds are generally not vascularized at the time of implantation, oxygen and nutrients to cells seeded inside scaffolds are initially delivered by diffusion. Cyclic RGDfK modification may enhance vessel growth by recruiting endothelial cells into scaffolds and promoting endothelial cell adhesion and proliferation.

A preexisting vascular network, either by implanting a vascular growth factor containing scaffold before introducing cells or grown ex vivo using bioreactors, would be more desirable, may decrease cell death, and enhance regenerative effects.^36,37 This is suggested by our studies with islet cell transplantation where the modified scaffolds were implanted between the abdominal rectus muscle and subsequently injected with pancreatic β-cells after a prevascularization period of 2 weeks.⁶

Scaffolds seeded with 1 × 10⁶ hMPCs increased blood vessel formation in the border zone of the myocardial infarct and augmented cardiac function. The functional improvement of cardiac function upon treatment with a variety of bone marrow-derived precursor cells has generally been attributed to paracrine stimulation and stimulated neovascularization.³⁸ Surprisingly, this effect was not observed using scaffolds with the higher dose of 3 × 10⁶ hMPCs. This finding may be explained by “overcrowding” of cells, which is related to levels of paracrine factors (i.e., cytokines) that hMPCs secrete, such as IL-6 and MCP-1.

Clinical dose escalation trials using direct intramyocardial mesenchymal stem cell injection have found similar dose-dependent effects.³⁹ Local excess cytokine concentrations may be toxic to cardiomyocytes and abolish the beneficial effects of transplanted cells. Alternatively, a high number of cells inside scaffolds may lead to accelerated cell death in vivo due to an initial lack of oxygen and nutrients, abrogating their beneficial effects. The efficacy of cell delivery in scaffolds can be maximized since cell dose can be controlled to a greater extent and cell loss by a lower extent compared to direct cell injection.⁴⁰

Limitations to echocardiographic assessment of rat hearts following MI are well known.³¹ Variability in measurements of ejection fraction in rats without MI in different studies have been reported to be 60%, 75%, and 91%, and the corresponding values for LVFS were 48%, 41%, and 56%. We used LVFS due to its reported lower variability compared to other parameters such as stroke volume or cardiac output; however, more advanced techniques such as magnetic resonance imaging may yield more accurate results.⁴¹

One animal's fractional shortening in the 1 × 10⁶ hMPC group normalized from 32.7% on day 0 to 57.4% on day 7, a 75.9% increase (Fig. 6d). We deemed this change to be plausible; another animal showed an increase in LVFS from 17.1% to 29.3%, a 71.0% increase; hence, we included the data point as part of the primary data of overall LVFS change, which was within the Q3 + 1.5 × IQR range of 106.4 (Fig. 6a). Of note, the animal's LVFS (i.e., infarct size) on day 0 did not differ significantly from the other animals' “LVFS,” which is relevant in this setting since initial degree of functional impairment of infarcted myocardium has been shown to affect functional outcome following intramyocardial cell transplantation.⁴²

Grubbs' outlier detection test⁴³ showed that both overall primary and normalized LVFS change data did not contain statistically significant outliers (not shown). It would need to be established whether animals in the 1 × 10⁶ hMPC group were either good responders, for instance due to genetic polymorphisms, or whether other causes (i.e., data variability) are responsible for the results. Given the relatively low data base numbers, additional animal studies would add to robustness of the data and increased statistical validity.

Many natural and synthetic materials are being evaluated in combination with a number of growth factors and/or cell types for clinical applications in cardiac tissue engineering and cell transplantation.⁴⁴ This study sought to evaluate an economical, practical, and clinically feasible scaffold material that promotes cell survival without provoking adverse immunological reactions. Cyclic RGDfK-modified alginate is such a biomaterial that meets these properties.

Estimated material retail costs to produce 1 cyclic RGDfK-modified alginate scaffold suitable for clinical use (size 100 × 0.75 mm using 2% cyclic RGDfK-modified alginate) would be approximately US$1500, excluding cells and cell culture materials. Financial modeling would be needed to calculate whether this is an acceptable price in addition to standard treatments. It would add to the costs of direct cell injection by percutaneous coronary intervention (PCI) or coronary artery bypass grafting. Depending on effectiveness and efficacy, and assuming scaffolds would not need replacement frequently, scaffold treatment could offer an alternative cost-effective approach.

Conclusion

Purified cyclic RGDfK peptide-modified 3D alginate scaffolds are biocompatible and nonimmunogenic, enhance cell viability, promote angiogenesis, and may be used as a means to deliver cells to myocardial infarct areas to improve neovascularization and cardiac function.

Disclosure Statement

H.P.S., P.W., and M.A.H. are provisional patent holders of the procedures described in this article (patent application No. 20100196441). S.I. is Executive Director of Mesoblast Limited and holds several patents on HMPL technology. All other authors have no competing financial interests.