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. Author manuscript; available in PMC: 2020 Feb 12.
Published in final edited form as: Chem Mater. 2019 Jan 11;31(3):1006–1015. doi: 10.1021/acs.chemmater.8b04486

Assembly of Bifunctional Aptamer–Fibrinogen Macromer for VEGF Delivery and Skin Wound Healing

Nan Zhao 1, James Coyne 1, Ming Xu 2, Xiaolong Zhang 1, Akiho Suzuki 1, Peng Shi 1, Jinping Lai 1, Guo-Hua Fong 3, Na Xiong 2, Yong Wang 1,*
PMCID: PMC6761992  NIHMSID: NIHMS1007750  PMID: 31558852

Abstract

Macromolecular assembly has been studied for various applications. However, while macromolecules can recognize one another for assembly, their assembled structures usually lack the function of specific molecular recognition. We hypothesized that bifunctional aptamer-protein macromers would possess dual functions of molecular assembly and recognition. The data show that hybrid aptamer-fibrinogen macromers can assemble to form hydrogels. Moreover, the assembled hydrogels can recognize vascular endothelial growth factor (VEGF) for sustained release. When the VEGF-loaded hydrogels are implanted in vivo, they can promote angiogenesis and skin wound healing. Thus, this work has successfully demonstrated a promising macromolecular system for broad applications such as drug delivery and regenerative medicine.

Keywords: molecular assembly, hydrogel, growth factor delivery, aptamer, regenerative medicine

1. Introduction

Macromolecular assembly is an important mechanism for building various natural structures such as ribosomes, microtubules, viruses and extracellular matrix 1–4. This mechanism has inspired the development of synthetic materials 3, 5–12 for biomedical applications such as immune regulation 13–15, drug delivery 16–18, regenerative medicine 19, and biosensing 20. However, synthetic materials developed via molecular assembly usually lack the ability to specifically recognize other biomolecules whereas the assembled structures in nature can recognize other biomolecules in a diverse array of biological processes. As molecular recognition is at the heart of various biological processes, lack of specific molecular recognition may represent a significant hurdle to the broad application of assembled synthetic materials. It is therefore important to explore macromolecular systems with dual functions of molecular assembly and recognition.

The purpose of this work was to develop hybrid aptamer-protein macromers with dual functions of both protein assembly and molecular recognition under physiological conditions. Specifically, we used fibrinogen (Fg) and an anti-vascular endothelial growth factor (VEGF) aptamer as a model to synthesize a hybrid aptamer-fibrinogen macromer. Fg is a natural protein that can assemble to form fibrin (Fn) hydrogels with thrombin-catalyzed hydrolysis 21, 22. Fn hydrogels have been widely used for drug delivery and regenerative medicine owing to their high biocompatibility 23. However, as Fn hydrogels have shortcomings such as low mechanical strength and high permeability, great effort has been made to functionalize Fg or Fn hydrogel derivatives. For instance, polyethylene glycol (PEG) was conjugated with Fg to synthesize PEG-Fg conjugates and the conjugates were polymerized via photo-initiated polymerization to form PEG-Fg composite hydrogels with high mechanical strength24–26. However, previous studies were not focused on the application of the principle of self-assembly for the development of affinity-based Fn hydrogels with improved molecular recognition and sequestration.

Aptamers are single-stranded oligonucleotides selected from RNA/DNA libraries. They are prominent synthetic ligands with high affinities and specificities against target molecules, holding great potential for various applications27–29. As aptamers are selected from large libraries, they have no theoretical limits in diversity. Moreover, aptamers form their functional structures through high-fidelity base pairing. This intramolecular hybridization is different from the interactions of amino acids and would not affect the functional conformation of the protein moiety in an aptamer-protein macromer and vice versa. Thus, Fg-aptamer macromers would assemble and form Fn hydrogels with the capability of molecular recognition.

In this work, we synthesized the aptamer-Fg macromer using the thiol-ene reaction. The assembly of this macromer was examined using numerous assays including circular dichroism, electron microscopy, fluorescence imaging and turbidity measurement. Its capability of molecular recognition was evaluated using surface plasmon resonance and protein retention. The functions of aptamer-Fn hydrogels in controlling VEGF release and promoting cell growth were examined both in vitro and in vivo. As VEGF is a highly potent angiogenic growth factor,30 the in vivo study was performed by examining the growth of new blood vessels and the procedure of skin wound healing.

2. Materials and Methods

2.1. Materials

Chemical reagents were obtained from Sigma-Aldrich (St. Louis, MO) and Thermo-Fisher Scientific (Waltham, MA). Biological reagents were obtained from Thermo-Fisher Scientific, Peprotech (Rocky Hill, NJ), Santa Cruz (Dallas, TX), Cell Signaling Technology (Beverly, MA), and Jackson ImmunoResearch Laboratories Inc (West Grove, PA). Nucleic acid sequences were obtained from Integrated DNA Technologies (Coralville, IA). Human umbilical vein endothelial cells (HUVECs) were obtained from Thermo-Fisher Scientific (Waltham, MA). Detailed information of materials is included in the Supporting Information.

2.2. Synthesis of aptamer-fibrinogen (Ap-Fg) macromers

50 mg of fibrinogen (Fg) was reacted with acrylic acid N-hydroxysuccinimide ester (AA-NHS) at a molar ratio of 1:100 in a NaHCO3 solution (0.1 M, pH=8). The reaction was carried out at 25 °C in a shaker (100 rpm) for totally 4 hours. Free monomers and byproducts were removed by washing the reaction mixture with a 100 kDa centrifugal filter. Thiol-modified anti-VEGF aptamers (Ap) and thiol-modified scrambled aptamers (Sc) were reduced in 50 mM tris (2-carboxy ethyl) phosphine hydrochloride (TCEP) at room temperature for 1 hour. The treated Ap and Sc were filtered in a 10 kDa filter. 30 nmol of Ap or Sc was reacted with 3 mg acrylate modified Fg at 37 °C in Tris-HCl buffer to synthesize aptamer-fibrinogen (Ap-Fg) macromer or scrambled aptamer-fibrinogen (Sc-Fg) macromer, respectively. After the reaction, the Ap-Fg macromer or Sc-Fg macromer was purified with a 100 kDa centrifugal filter, filtered through a 0.2 µm filter, and stored at −20 °C.

2.3. Characterization of Ap-Fg macromers

Dynamic Light Scattering (DLS).

Native Fg or Ap-Fg macromer was diluted with PBS to 0.1 mg/mL. 0.6 mL of the diluted solutions were transferred to a 12 mm square polystyrene cuvette and analyzed with a Zetasizer Nano ZS (Malvern, UK).

Circular dichroism.

Native Fg or Ap-Fg macromer was diluted to a final concentration of 500 µg/mL. 200 µL of the solution was added to a 1-mm path length quartz cell. Spectra were recorded over the range of 190–260 nm at room temperature with a J-1500 Circular Dichroism Spectrometer (JASCO, Easton, MD).

Transmission Electron Microscopy (TEM).

5 µL of 10 µg/mL native Fg or Ap-Fg macromer was absorbed onto the ultra-thin carbon grids blocked with 1% bovine serum albumin (BSA) for 60 min at room temperature. The grids were then incubated with biotin-labeled complementary sequence of the aptamers for 30 min at room temperature. Streptavidin-labeled gold nanoparticles (3 nm) were blocked with 20 mM glycine for 2 hours at room temperature. Then the grids were incubated with the solution of gold nanoparticles and stained with 2% phosphotungstic acid for 2 min. The grids were imaged under a TEM (Tecnai LaB6, FEI, Japan).

Surface Plasmon Resonance (SPR).

SPR Spectroscopy (SR7500DC, Reichert Analytical Instrument, Depew, NY) was used to examine the binding affinity of Ap-Fg macromer to vascular endothelial growth factor (VEGF) based on a previously published protocol 31. Briefly, a carboxyl-functionalized sensor chip was activated with N-hydroxysuccinimide (NHS) and carbodiimide hydrochloride (EDC) for 10 min. VEGF (10 µg/mL) was immobilized on the chip by injecting for 10 minutes in sodium acetate (10 mM, pH = 5.2). After equilibration, free aptamers, native Fg, Sc-Fg macromer, or Ap-Fg macromer was injected over the chip. The data were analyzed with Scrubber 2.0 software (BioLogic Software, Australia).

2.4. Assembly of Ap-Fg macromer

The assembly of Ap-Fg macromer was studied by measuring the turbidity at 350 nm 32. 50 μL of 8 mg/mL native Fg or 25 μL of 12 mg/mL native Fg mixed with 25 μL of 4 mg/mL Ap-Fg macromer was added to a 96-well cell culture plate (Ap, 10 µM). After the addition of 50 μL mixture of 0.4 U/mL thrombin and 20 mM CaCl2, the turbidity of the solution was monitored with an Infinite M200 Pro microplate reader (Tecan, Grödig, Austria).

2.5. Scanning Electron Microscopy (SEM) of assembled fibrin fibers

20 µL of 12 mg/mL native Fg mixed with equal volume of 4 mg/mL Ap-Fg macromer was allowed to assemble in the presence of 0.2 U/mL thrombin and 10 mM CaCl2 to form aptamer-fibrin (Ap-Fn) hydrogel (Ap, 5 µM). Ap-Fn hydrogel and native fibrin (Fn) hydrogel were frozen at −80 °C and lyophilized in a Freeze-dry System (Labconco, Kansas City, MO). The samples were sputter-coated with iridium and imaged with a Field Emission scanning electron microscope (Zeiss Sigma, US). The diameter of the fibrin fibers was evaluated with ImageJ (NIH, Bethesda, ML).

2.6. Preparation of aptamer-fibrin (Ap-Fn) hydrogel

For all the hydrogels used in the growth factor release, cell culture and in vivo studies, the final total concentration of fibrinogen was 10 mg/mL and the concentration of thrombin was 1 U/mL. To make the native fibrin hydrogel (Fn), Fg was diluted into 20 mg/mL. Thrombin and CaCl2 was combined to make 2 U/mL thrombin and 20 mM CaCl2 solution. Then equal volume of the two parts were mixed and transferred into a polydimethylsiloxane mold (diameter of 8 mm and thickness of 1 mm). To make the Ap-Fn hydrogel, Fg was mixed with Ap-Fg macromers to prepare the final solution with 20 mg/mL of total Fg. Then the mixture was mixed with an equal volume of thrombin and CaCl2 solution. To prepare VEGF-loaded hydrogels, the pregel solutions were mixed with VEGF. A scrambled aptamer-fibrinogen (Sc-Fg) macromer was also used to prepare hydrogel as control (i.e., Sc-Fn hydrogel).

2.7. Fluorescence imaging of Ap-Fn hydrogel

100 µL of 10 mg/mL Fn and 10 mg/mL Ap-Fn (Ap, 13 µM) were soaked into fluorescein amidite-labeled complementary sequence (FAM-cDNA) staining solution, washed with PBS, and imaged with a Maestro Imaging System (CRI, Woburn, MA). The optical images of the hydrogel were taken with a digital camera. For confocal imaging, the Ap-Fn hydrogels were fixed in 4% paraformaldehyde, stained with FAM-cDNA, washed with PBS, and imaged with a confocal microscope (Olympus FV1000, Center Valley, PA).

2.8. Examination of VEGF retention and release

VEGF retention.

Ap-Fn hydrogel (50 µL) with different concentrations of aptamer (0, 0.1, 0.2, 0.4, and 0.8 µM) were synthesized to test the effect of molar ratio of Ap to VEGF on VEGF retention. VEGF loaded hydrogels were incubated with 1 mL of release media (0.1% BSA in DPBS or medium 200). After 24 hours, the release media were collected and used for ELISA according to the protocol provided by the manufacturer.

VEGF release.

For the experiments of sustained VEGF release, Fn hydrogels were synthesized at the mole ratio of 20:1 (Ap:VEGF). The loading amount of VEGF was 200 ng. Hydrogels were incubated in 1 mL of release medium. At pre-determined time points, 1 mL of the release medium was collected, stored at −20 °C and replenished with fresh release medium. VEGF was analyzed using ELISA after all samples were collected. The data were presented as cumulative release.

2.9. Cell culture

Human umbilical vein endothelial cells (HUVECs) were expanded using medium 200 (M200) supplemented with 2% Low Serum Growth Supplement (LSGS) in 0.1% gelatin coated cell culture flasks. HUVECs of passage 4 to 8 were used in all cell experiments.

2.10. Cell migration assay

HUVECs were seeded in the plate with the density of 1×105 cells per well. Cells were starved in M200 with 1% FBS overnight. A simulated wound was created by scratching the cell monolayer with a p200 tip. Then cells were treated with different release media. After cultured for 24 hours, cells were stained with Calcein AM and imaged with an Olympus IX73 microscope (Center Valley, PA). The widths of the simulated wound were measured with ImageJ. The cell closure was calculated using this equation: closure=100× (1-final gap width/ initial gap width).

2.11. Degradation of Ap-Fn hydrogels

HUVECs were seeded with the density of 1×103 per well on the Fn hydrogel or the Ap-Fn hydrogel. At predetermined time points, the media were collected, and the amount of degraded fibrin was quantified by bicinchoninic acid (BCA) assay. At day 7, cells were removed by treating the materials with ice-cold DPBS. The materials were stained with FAM-cDNA and washed with PBS. Then the hydrogels were imaged with the Maestro imaging system.

2.12. Cell proliferation on VEGF-loaded Ap-Fn hydrogels

HUVECs were seeded with the density of 1×103 per well on native Fn hydrogels, Sc-Fn hydrogels loaded with 100 ng VEGF (Sc-Fn+VEGF), or Ap-Fn hydrogels loaded with 100 ng VEGF (Ap-Fn+VEGF). At predetermined time points, cells on the hydrogels were stained with the LIVE/DEAD viability kit and imaged with the Olympus IX73 microscope. The number of cells on the hydrogels was quantified with ImageJ. The cell viability was also examined using the MTS assay and the absorbance of the solution at 490 nm was measured with the Infinite M200 Pro microplate reader.

2.13. Measurement of VEGF retention during the cell culture

At day 7 after HUVEC culture on the hydrogels, cells were removed by washing the samples with ice-cold DPBS. Hydrogels were treated with 100 U/mL DNase. The amount of VEGF in the solution was quantified with ELISA.

2.14. Examination of p-VEGFR2 and endothelial gaps

p-VEGFR2 and VE-Cadherin were immunostained. 2×103 HUVECs were seeded on the hydrogels. The cell culture medium was changed every other day. At day 7, hydrogels with cells were fixed in 4% paraformaldehyde solution. Cells were blocked with 3% BSA solution, incubated with primary antibody overnight at 4 ºC, stained with fluorophore-labeled secondary antibody, mounted in ProLong diamond antifade mountant with DAPI, and imaged with the Olympus IX73 microscope. The fluorescence intensity of p-VEGFR2 was quantified in ImageJ. For VE-cadherin staining, fluorescein-labeled primary antibody was used to treat the cells. Endothelial gaps were calculated by dividing the gap areas by the total area of each image.

2.15. Mouse skin wound model

The animal experiments were performed according to the protocol approved by the Pennsylvania State University Institutional Animal Care and Use Committee. C57BL/6 mice (age of 8 to 10 weeks) with weight of 25 to 30 g were used. 1.2% avertin was injected intraperitoneally to mice (30 µL/g). The dorsal hair of the mice was removed by an electronic razor followed by Veet hair depilatory cream treatment. The hair removal cream was cleaned by washing with sterile DPBS and the skin was sterilized with 70% ethanol and povidone-iodine. An 8 mm circular full skin wound was created on the dorsal skin. After the hydrogels were applied, the wounds were covered with a transparent Tegaderm film. Fn hydrogels without VEGF, Sc-Fn hydrogels loaded with 200 ng VEGF (Sc-Fn+VEGF), and Ap-Fn hydrogels loaded with 200 ng VEGF (Ap-Fn+VEGF) were used to treat the wounds. At different days, the optical images of the wounds were taken by using a digital camera.

2.16. Histology analysis

Mice were euthanized at day 13 and the skin tissues were collected with an 8 mm punch. The tissue samples were cut from the center into two halves. One half was paraffin-blocked for histology staining and the other half was frozen in O.C.T. Compound for immunostaining. Paraffin blocked tissues were sectioned into 5 µm and stained with Hematoxylin and Eosin (H&E) in a Leica autostainer (Buffalo Grove, IL). The length of the epithelium layer and the distance between the wound margins were quantified in ImageJ. The epithelialization ratio was calculated by dividing the length of the epithelium layer by the initial length of the wound. For Trichrome staining, deparaffinized tissues were soaked into Bouin’s solution overnight, incubated with Weigert’s iron hematoxylin solution, and stained with Masson’s Trichrome staining kit. The stained samples were mounted into xylene substitute mountant and imaged with a BZ-X700 microscope (Keyence, Itasca, IL). Collagen positive areas were quantified using Fiji (Image J) with a built-in Masson Trichrome color deconvolution. The total blue area was normalized to the total tissue area.

2.17. Immunostaining

CD31 staining.

Paraffin blocked tissues were sectioned into slices of 5 µm, deparaffinized, and boiled in sodium citrate buffer (pH=6) for 20 min. The sections were equilibrated to room temperature and blocked with serum blocking solution (3% BSA and 3% goat serum in PBS) for 1 hour at room temperature. The blocked samples were incubated with rabbit anti-mouse CD31 antibody (1:200 dilution) overnight at 4 °C. After the antibody treatment, the samples were further treated with 3% H2O2 solution, biotinylated goat anti-rabbit secondary antibody, Avidin-Horseradish Peroxidase, and Diaminobenzidine substrate. Then the tissues were soaked into hematoxylin solution, dehydrated in gradient ethanol, cleared in xylene substitute, and mounted in xylene substitute mountant. The total number of blood vessels and the total area of blood vessels were quantified with ImageJ.

Keratin staining.

The frozen tissues were cryo-sectioned into slices of 10 µm. The tissue sections were equilibrated to room temperature and then soaked into PBS for 10 min. The tissue sections were further boiled in sodium citrate buffer for 20 min, blocked with serum blocking solution and stained with fluorophore-labeled keratin 5 and keratin 10 antibodies. Afterwards the tissues were mounted in ProLong diamond antifade mountant with DAPI and imaged with the Olympus IX73 Microscope.

2.18. Statistics

All the data were presented as mean± standard deviation. Statistical analysis was performed by using Prism 5.0 (GraphPad Software Inc., La Jolla). Mann-Whitney test was performed for the comparison of two groups. One-way analysis of variance (ANOVA) followed by the Bonferroni post-test or two-way ANOVA followed by the Bonferroni post-test was performed for the comparison of multiple groups. The result was considered statistically different if p < 0.05.

3. Results

3.1. Synthesis and characterization of aptamer-fibrinogen (Ap-Fg) macromers

We modified native fibrinogen (Fg) with acrylate and then conjugated Fg with aptamer through thiol-ene reaction (Fig.1a & Fig.S1). Transmission electron microscopy (TEM) and gel electrophoresis were used to confirm the formation of Ap-Fg macromers (Fig.1b & Fig.S2). In the TEM image, approximately two gold nanoparticles (black dot) were attached to one Ap-Fg macromer (rod-like structure). It suggests that each macromer had approximately 2 aptamers. The dynamic light scattering (DLS) analysis shows that the Ap-Fg macromer and Fg had a similar hydrodynamic diameter of ~30 nm (Fig.1c), indicating that aptamer conjugation did not affect the overall size of Fg. Consistent with the DLS measurement, the circular dichroism (CD) spectra show that Fg maintained the same secondary structure after the conjugation with aptamers (Fig.1d).

Fig.1|. Molecular assembly.

Fig.1|

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a, Schematic illustration of Ap-Fg macromer synthesis and assembly. Fg, fibrinogen; Ap, aptamer; Ap-Fg, aptamer-fibrinogen macromer. b, TEM images of native Fg and Ap-Fg macromer treated with nanoparticles. The rod-like structures are the Fg. The black dots are gold nanoparticles. c, Measurement of Ap-Fg macromer using dynamic light scattering. d, Circular dichroism of Fg and Ap-Fg macromer. e, Turbidity measurement of the solution of Ap-Fg macromer for showing dynamic assembly. f, SEM images. The right graph shows the size distribution of the fibers. g, Optical (left upper panel) and fluorescent (left lower panel) images of bulk hydrogels. The two fluorescence images on the right show the hydrated structure of fibers. Hydrogels were treated with FAM-labeled complementary sequences for fluorescent staining.

3.2. Evaluation of Ap-Fg macromer assembly

Fg is converted into fibrin (Fn) after the cleavage of fibrinopeptides 21, 22. During this conversion, Fn spontaneously polymerizes via self-assembly and forms an insoluble fibrin hydrogel. This procedure of Fn assembly can be characterized using the turbidity test 32, 33. The turbidity profiles show that the solutions of native Fn and aptamer-fibrin (Ap-Fn) hydrogels reached plateau in approximately 30 min (Fig.1e). More importantly, the two profiles nearly overlap, demonstrating that the presence of aptamer did not affect Fn assembly. To confirm the turbidity results, we examined the structures of Fn and Ap-Fn hydrogels using scanning electron microscopy (Fig.1f). Consistent with the literature 34–36, native Fn hydrogels exhibited the structure of a crosslinked mesh with threads. Ap-Fn hydrogels showed the same morphology as Fn hydrogels. A more detailed analysis was performed to examine the diameters of the threads, further confirming that Fn and Ap-Fn hydrogels virtually had the same structures (Fig.1f), which suggests that the incorporation of nucleic acid aptamers into Fg does not affect assembly behavior of Fg or the overall structure of assembled Fn hydrogels.

To further demonstrate the assembly of Ap-Fg macromers in forming hydrogels, we stained Ap-Fn hydrogels using the fluorescein amidite-labeled complementary sequence (FAM-cDNA) of the aptamer (Fig.1g). Both Fn and Ap-Fn hydrogels were opaque in the optical images. However, the Fn hydrogel did not exhibit fluorescence whereas the Ap-Fn hydrogel emitted strong fluorescence. Confocal microscopy images confirm that each strand of the Ap-Fn hydrogel was labeled with FAM-cDNA (Fig.1g). We also treated the hydrogels sequentially with biotin-labeled cDNA and streptavidin-labeled nanoparticles and imaged the hydrogels with TEM (Fig.S3). The Ap-Fn hydrogel retained more nanoparticles than the Fn hydrogel confirming the participation of Ap-Fg macromers during the formation of hydrogels.

3.3. Evaluation of molecular recognition and sequestration

After the demonstration of macromer assembly, we studied the function of the macromer in molecular recognition (Fig.2a). The ability of the Ap-Fg macromer in recognizing VEGF was examined using Surface Plasmon Resonance (SPR). A scrambled aptamer-fibrinogen (Sc-Fg) macromer was used as a control. The maximal signal of the Ap-Fg macromer was ten times as high as that of the Sc-Fg macromer. Moreover, the signal of the Sc-Fg macromer decreased more sharply than that of the Ap-Fg macromer during molecular dissociation. With curve fitting, we calculated the dissociation constant (Kd). The Kd values of free Ap and Ap-Fg are ~160 and 240 nM, respectively. The increase of Kd may result from the steric hindrance of Fg. Despite this increase, the SPR analysis shows that the Ap moiety of the macromer is still capable of recognizing VEGF.

Fig.2|. Molecular recognition and sequestration.

Fig.2|

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a, Molecular recognition examined by surface plasmon resonance spectroscopy. Fg, fibrinogen; Pr, protein; Ap, aptamers; Sc-Fg, scrambled aptamer-fibrinogen macromer; Ap-Fg, aptamer-fibrinogen macromer. Vascular endothelial growth factor (VEGF) was immobilized on the biochip. The solution of Ap, Fg, Sc-Fg macromer or Ap-Fg macromer was injected over the biochip. b, VEGF sequestration and release. Left (lower panel): effect of the aptamer-to-VEGF ratio on VEGF sequestration in the Ap-Fn hydrogel; right: sustained VEGF release; n=3. c, Wound closure in the in vitro wound healing assay. Fn, release media from the Fn hydrogel (no VEGF) at day 14; Ap-Fn, VEGF of 10 ng/mL in the release media from the Ap-Fn hydrogel collected at day 14; stock VEGF of 10 ng/mL. Cells were stained with Calcein AM. The quantitative analysis is shown in the right graph (n=6). ***, p<0.001.

We further examined whether the aptamer could recognize its cognate protein after molecular assembly (Fig.2b). A pre-gel solution was prepared to contain the Ap-Fg macromer and VEGF. When this solution was triggered to polymerize, VEGF was automatically incorporated into the Ap-Fn hydrogel. VEGF retention increased from 31.6% to 64.9% with the increase of the aptamer-to-VEGF ratio from 0 (i.e., no aptamer) to 40. Notably, simply increasing the Fn concentration without aptamer did not improve VEGF retention (Fig.S4). Thus, our data show that aptamer functioned as a specific VEGF-binding site in the Ap-Fn hydrogel after molecular assembly.

As aptamer can recognize and retain its cognate protein after molecular assembly, we then examined the ability of aptamer in controlling protein release. Before the protein release test, we examined the permeability of hydrogels using dextran release (Fig.S5a). Fn and Ap-Fn hydrogels were both highly permeable. Virtually 100% of dextran was released from these hydrogels within 24 hours. We also characterized the rheological properties of the hydrogels. The data suggest that aptamer functionalization did not change the mechanical properties of Fn hydrogels (Fig.S5b).

We then compared the release kinetics of dextran and VEGF from the Fn hydrogels. The release of VEGF from the Fn hydrogels was slower than the release of dextran. As VEGF and dextran have a similar molecular weight, it suggests that VEGF may bind to fibrin. However, the native Fn hydrogel released over 97% of VEGF within the first 3 days (Fig.2b), suggesting that VEGF-Fn interactions are too weak to achieve the sustained release of VEGF, which is consistent with the literature 37.

By contrast, the Ap-Fn hydrogel released VEGF in a biphasic manner (Fig.2b & Fig.S6). Within the first 6 hours, 35.7% of VEGF was rapidly released from the Ap-Fn hydrogel. We also examined whether the presence of VEGF affected Fn assembly and the structures of the Fn hydrogels (Fig.S7). Our results suggest that VEGF did not affect molecular assembly or hydrogel structures. After the first phase, VEGF was slowly released from the Ap-Fn hydrogel and the daily release rate of VEGF was ~3% to 4% between day 2 and 14.

While VEGF did not affect the macromer assembly, it is equally important to examine whether the assembly would affect VEGF. Thus, the released VEGF was used to stimulate human umbilical vein endothelial cells (HUVECs). Both the released VEGF and the stock VEGF could promote endothelial cell migration (Fig.2c). There was no statistically significant difference between the released VEGF and the stock VEGF, which suggests that VEGF could maintain high bioactivity during the Ap-Fg assembly. Based on those results, we concluded that aptamers can be used as specific growth factor-binding sites for protein assembly.

3.4. In vitro examination of the functions of the Ap-Fn hydrogel

Degradation is an important characteristic of biomaterials. Degradation of protein hydrogels can be mediated by enzymes in biological fluids. We first examined the degradation behavior of the hydrogels in the plasmin solution and the cell culture media (Fig.S8a & b). Ap-Fn and native Fn hydrogels exhibited the same degradation profiles in the two solutions. It suggests that aptamer did not affect the degradation behavior of Fn hydrogels. The degradation of the hydrogels was also independent of the presence of VEGF (Fig.S8c). We then cultured the cells on the hydrogels, finding that the degradation was not affected by endothelial cells (Fig.3a). However, Fn hydrogels can be quickly degraded during the procedure of natural tissue repair. This difference may result from the in vitro 2-dimensional cell culture condition and the limited amount of plasmin secreted by endothelia cells. Moreover, the fluorescence intensity of the stained aptamer did not decrease, showing that the degradation of aptamer in the hydrogel was negligible during the procedure of in vitro cell culture (Fig.3b). Resultantly, HUVCEs seeding on the hydrogel materials did not significantly change the release profile of VEGF (Fig.S9) and over 20% of loaded VEGF was retained in the Ap-Fn hydrogel after 7 days cell culture (Fig.3c).

Fig.3|. Cell-hydrogel interactions.

Fig.3|

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a, Degradation of hydrogels. +/− cell: with/without HUVECs. b, Imaging of aptamers in the Ap-Fn hydrogels for showing aptamer stability. c, Examination of VEGF retained in hydrogels at day 7 post HUVECs seeding. d, Cell quantification at day 7. Left, cell counting. Right, MTS analysis. e, Immunostaining of p-VEGFR2 of HUVECs cultured on the hydrogels at day 7. f, Examination of endothelial gap at day 7. Endothelial gaps are enclosed with yellow lines for clear legibility. a-d, n=3; e-f, n=4; ns, no significant difference; ***, p<0.001.

We also examined the effect of the hydrogels on cell growth. Keratinocytes grew only slightly faster on the VEGF-loaded Ap-Fn hydrogel compared to the two control hydrogels whereas fibroblasts grew similarly on all three hydrogels (Fig.S10). While HUVECs could grow on all three hydrogels, endothelial cells on the VEGF-loaded Ap-Fn hydrogel (Ap-Fn+VEGF) grew faster (Fig.S11). The viable number of endothelial cells on the VEGF-loaded Ap-Fn hydrogel was significantly higher than that of the two control hydrogels on day 7 (Fig.3d). To further prove that the increased endothelial cell growth on Ap-Fn hydrogel was due to the released VEGF, we measured the expression of phosphorylated vascular endothelial growth factor receptor-2( p-VEGFR2) of HUVCEs as VEGFR2 plays important role in the VEGF-induced cell proliferation 38. The HUVECs on the VEGF-loaded Ap-Fn hydrogel expressed a higher amount of p-VEGFR2 than those on the control hydrogels (Fig.3e), which is consistent with the western blot analysis showing that the released VEGF could stimulate the expression of p-VEGFR2 (Fig.S12). Correlated to the expression of p-VEGFR2, the endothelial gap increased on the VEGF-loaded Ap-Fn hydrogel (Fig.3f).

3.5. Examination of the function of Ap-Fn hydrogels in promoting skin wound healing

Based on the in vitro experiments, we hypothesized that sustained release of VEGF from Ap-Fn hydrogels might promote angiogenesis and tissue regeneration in vivo. We tested this hypothesis in the mouse model for full thickness skin wound healing. Different from the human skin, skin wounds in mice can quickly heal through contraction 39. To avoid this artifact, full thickness skin wounds were covered with transparent Tegaderm films that could effectively minimize contraction (Fig.S13).

We first tested the biocompatibility of Ap-Fn hydrogels. Ap-Fn hydrogels did not display significant cytotoxicity or elicit immune responses (Fig.S14). To test the angiogenic and wound-healing functions of VEGF delivered by Ap-Fn hydrogels, we loaded Ap-Fn hydrogels with VEGF (Ap-Fn+VEGF) and then implanted them into the dorsal skin wounds. Native Fn hydrogels without VEGF (Fn) and Sc-Fn hydrogels loaded with VEGF (Sc-Fn+VEGF) were used as controls. Hydrogels in all three groups were well attached to the wound bed during the observed period (Fig.4a). By day 7, wounds in Ap-Fn+VEGF -treated mice began to show significantly reduced sizes compared with the Fn group (Fig.4a). By day 11, wounds in the Ap-Fn+VEGF group were significantly smaller than both Fn and Sc-Fn+VEGF controls (Fig.4a). At day 13, the average wound size of the Ap-Fn+VEGF treated group was 7.2% ±3.2% of the starting size at day 0, whereas the corresponding values in the Fn and Sc-Fn+VEGF groups were 33.2% ± 10.3% (Fn) and 27.8% ± 12.5% (Fn-sc+VEGF), respectively (Fig.4a).

Fig.4|. VEGF delivery for skin wound healing.

Fig.4|

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a, Kinetics of wound closure. The wound areas measured at different time points were normalized to their original areas measured at day 0. Red stars: comparison between Ap-Fn+VEGF and Sc-Fn+VEGF. Green stars: comparison between Ap-Fn+VEGF and Fn. b, H&E staining of the skin tissue collected at day 13. E, epidermis; F, fibrin; G, granular tissue. Red arrows, the edges of epithelial tongues; blue arrows, wound margins. c, Distance of the wound margins. d, Quantification of wound re-epithelialization. n=8; ns, no significant difference; *, p<0.05; **, p<0.01; ***, p<0.001.

We harvested the wound tissues at day 13 and analyzed the histology by Hematoxylin & Eosin staining (Fig.4b). Faster wound closure was observed in the Ap-Fn+VEGF group, as manifested by the shorter distance between wound margins (Fig 4b, blue arrows) than that in Fn and Sc-Fn+VEGF groups (Fig.4c). The re-epithelialization ratio reached near 100% in the Ap-Fn+VEGF group (Fig.4d). Notably, six out of eight mice in this group showed complete re-epithelialization. By contrast, re-epithelialization ratios in Fn and Sc-Fn+VEGF treated wounds were 79.7% ± 9.3% and 84.7% ± 14.0%, respectively, leaving hydrogel material visible in epithelium-free regions (Fig 4b, where red arrows indicate the edge of epithelial tongues). Significantly, Ap-Fn+VEGF treated wounds formed more hair follicles than other two groups (Fig.S15), consistent with the effect of VEGF in stimulating hair follicle growth 40, 41.

To confirm that sustained VEGF release from Ap-Fn hydrogels promoted blood vessel formation, we stained the wound tissue sections with anti-CD31 antibodies (Fig.5a), and quantified the number of blood vessel and tissue area occupied by vascular lumens (Fig.5b). The number of blood vessels per unit area was 25% higher in the Ap-Fn+VEGF group compared with the two controls. Furthermore, the blood vessel area in Ap-Fn+VEGF was almost twice as that in Fn and Sc-Fn+VEGF. These data unequivocally demonstrate the proangiogenic effects of sustained VEGF release from Ap-Fn hydrogels.

Fig.5|. Examination of angiogenesis and wound remodeling.

Fig.5|

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a, Images of blood vessels stained with anti-CD31 antibody. The red arrows indicate blood vessels. b, Quantification of blood vessels in number and area (n=8) c, Trichrome staining of the skin tissue. The blue color indicates the collagen staining. Collagen-positive areas were normalized to the total areas of the skin tissue (n=6). d, Keratin staining. k5: keratin 5; k10: keratin 10. Nuclei were stained with DAPI. ns, no significant difference; **, p<0.01; ***, p<0.001.

Finally, we examined the remodeling of the skin tissue by Trichrome staining (Fig.5c). In the skin wound healing model, the replacement of the provisional extracellular matrix with collagenous matrix is a key part of wound remodeling 42. Around 60% of the wound area in the Ap-Fn+VEGF group was collagen-positive (blue color), which was much closer to the collagen content in the native skin compared with other two groups (Fig.5c). Because the in vitro results show that sustained VEGF delivery did not affect fibroblast proliferation (Fig.S10), the faster extracellular matrix remodeling of the Ap-Fn+VEGF treated group could be attributed to the indirect effect from sustained VEGF delivery. Sustained VEGF release promoted keratinocytes proliferation (Fig.S10). It has been reported keratinocytes can secrete growth factors such as basic fibroblast growth factor (bFGF) and transforming growth factor (TGF) 43, 44. Some of these growth factors such as bFGF can promote fibroblast proliferation and the secretion of extracellular matrix 45, which in turn could result in expedited extracellular matrix remodeling in vivo. The expression of keratin 5 and keratin 10, important cell markers for epithelialization, were also examined to further evaluate the wound healing procedure. Images of the epidermal layer were captured at 1.5 mm from the center of the wounds (Fig.5d & Fig.S16). While the expression of keratin 5 and keratin 10 was found in the newly formed epidermis in all three groups, keratin 5 and keratin 10 formed well-organized mature structures only in the Ap-Fn+VEGF group. In the neo-epidermis of the Ap-Fn+VEGF group, keratin 10 was restricted to the super-basal layer while keratin 5 in the basal layer. In sharp contrast, the epidermis in Fn and Sc-Fn+VEGF groups contained keratin 5 and keratin 10 in the entire epidermis. Taken together, our results demonstrate that VEGF released from the Ap-Fn hydrogel promotes angiogenesis and accelerates tissue regeneration in mouse skin wounds.

4. Discussion

Macromolecular assembly is an important principle for the development of synthetic materials. It has particularly received great attention in the field of biomedical engineering. A typical example is the assembly of proteins to form hydrogels or scaffolds for drug delivery and regenerative medicine. However, while these biomolecules can recognize one another for the success of assembly, their assembled structures usually do not have the ability to further recognize other proteins specifically with high affinities. Indeed our results show that while native Fn-mediated VEGF sequestration was slightly higher than dextran sequestration, this sequestration was extremely weak (Fig.2). Thus, the efficacy of native Fn in the clinical or bioengineering applications would be limited since most of these applications require to provide cells or tissues with soluble signaling molecules for a long period of time.

To solve this problem, we applied aptamers to functionalize Fg to develop Ap-Fn hydrogels for controlled VEGF delivery. Indeed, the anti-VEGF aptamer could significantly improve VEGF retention within Ap-Fn hydrogels for sustained VEGF release (Fig.2). However, while Ap-Fn hydrogels exhibited an improved release profile in comparison to the native Fn hydrogels, we still observed a rapid VEGF release in the first phase. It is different from our previous results showing the slow release of growth factors from aptamer-functionalized superporous hydrogels 31, 46. This rapid release of VEGF from Ap-Fn hydrogels may be attributed to two possibilities. When Fg is hydrolyzed for molecular assembly during the formation of Fn hydrogels, fibrinopeptides are produced 21, 22. If aptamers were conjugated to fibrinopeptides, aptamers would not be incorporated into the final Ap-Fn hydrogel. It would cause the rapid release of these aptamer-bound VEGF molecules. Another possibility may come from the initially unbound free VEGF during the formation of Fn hydrogels since this anti-VEGF DNA aptamer has a moderate affinity as indicated by ~100 nM Kd. In our previous studies, the aptamers bind to their cognate targets with Kd values at the level of 10 nM or even lower 31, 46. Thus, it is possible to reduce this initial burst release of growth factors from the Ap-Fn hydrogels by using aptamers of higher affinity, which will be studied in our future work.

A variety of wound dressings have been studied to treat skin wounds that affect millions of people each year 47. Of them, hydrogels have emerged as primary candidates for the development of advanced dressings. However, despite the diversity, synthetic hydrogels often lack the bioactivity and functions of the natural extracellular matrix that allows for the attachment of cells and the release of signaling molecules to further stimulate cell receptors. By using both Fg and the aptamer, we have demonstrated that the Ap-Fn hydrogel can not only release VEGF for the stimulation of cell growth but also allowe the cells to attach (Fig.3). Moreover, the VEGF-loaded Ap-Fn hydrogel could deliver VEGF to promote skin wound healing more effectively than the VEGF-loaded control hydrogel (Fig.4). However, this Ap-Fn hydrogel system may be further improved by using other methods such as the incorporation of PEG into Fn hydrogels.24–26 It is also important to note that the skin wound healing is a complex procedure involving multiple growth factors 48. Thus, while our data have shown that VEGF release from the Ap-Fn hydrogels can promote skin wound healing, it is also desirable to explore the co-delivery of multiple growth factors in the future. As aptamers have high binding affinities and specificities, it is possible to incorporate multiple aptamers into Fn hydrogels to control the release of their cognate growth factors with desired release kinetics.

5. Conclusion

This work has successfully demonstrated a promising assembly system based on the aptamer-protein macromer. The assembly procedure is driven by the protein and the ability of molecular recognition is provided by the aptamer. The assembled aptamer-protein hydrogel allows for not only the attachment of cells but also the sustained release of growth factors. Thus, this hybrid bifunctional assembly system holds great potential for drug delivery and regenerative medicine applications. Notably, while we used Fg as a model, it is possible to extend the same concept to other proteins and peptides or other types of macromolecules with the ability of self-assembly.

Supplementary Material

supplementary

Acknowledgements:

The authors thank Huck Institute of the Life Science and Material Characterization Lab at Pennsylvania State University (University Park, PA) for technical training and supports. We would also like to thank Dr. Ralph Colby and his group for assistance with the rheology characterization. This study was supported by the National Institutes of Health (HL122311; AR073364).

Footnotes

Competing interest: The authors declare no competing financial interests.

Supporting Information: Additional experimental methods and characterization figures

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