Hierarchical Porous Protein Scaffold Templated from High Internal

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Interface Components: Nanoparticles, Colloids, Emulsions, Surfactants, Proteins, Polymers

Hierarchical Porous Protein Scaffolds Templated from High Internal Phase Emulsion co-Stabilized by Gelatin and Gelatin Nanoparticles Huan Tan, Zhao Tu, Hongqian Jia, Xiaojun Gou, and To Ngai Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.7b04047 • Publication Date (Web): 09 Apr 2018 Downloaded from http://pubs.acs.org on April 9, 2018

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Hierarchical Porous Protein Scaffold Templated from High Internal Phase Emulsion co-Stabilized by Gelatin and Gelatin Nanoparticles

Huan Tan†, Zhao Tu ‡, Hongqian Jia†, Xiaojun Gou†, *, and To Ngai§, *

†

Key Laboratory of Medicinal and Edible Plants Resources Development of Sichuan Education

Department, Sichuan Industrial Institute of Antibiotics, Chengdu University, Chengdu, 610052, PR China ‡

Bazhong Hospital of Traditional Chinese Medicine, Bazhong, 636000, PR China

§

Department of Chemistry, The Chinese University of Hong Kong, Shatin, Hong Kong

ACS Paragon Plus Environment

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ABSTRACT: Recently, three-dimensional (3D) scaffolds produced using poly Pickering high internal phase emulsions (polyHIPEs) technology are particularly attractive in biomedical application. However, until now the most investigated polyHIPEs are hydrophobic composites originating from synthetic polymers. Here we present an investigation of a hierarchical porous protein scaffold templated from oilin-water (O/W) HIPEs co-stabilized by fully natural materials, gelatin and gelatin nanoparticles. Fairly monodispersed gelatin nanoparticles were firstly synthesized through a two-step desolvation method, and then they were used as emulsifiers together with gelatin to fabricate stable HIPEs with adjustable droplet size distribution and rheology. Monolithic scaffolds were formed by cross-linking the HIPEs with polymers as low as 2.5 wt% in the continuous phase, which appropriately presented a general high porosity and had an interconnected porous morphology with smooth pore walls and textured structures. Furthermore, the scaffolds were degradable and showed reasonably good biocompatibility; L929 cells could adhere to the surface of the materials and exhibited intensive growth and well-spread morphology. This hierarchical porous protein scaffold could, therefore, have important application as a 3D scaffold that offers enhanced cell adhesion and functionality.

KEYWORDS: Pickering emulsion, high internal phase emulsion, gelatin, gelatin nanoparticle, 3D scaffold


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INTRODUCTION In recent years, porous three dimensional (3D) scaffolds with a fully interconnected geometry and high surface area have been attractive matrices for biotechnological and biomedical applications. To mimicking the native extracellular matrix (ECM) as much as possible, the physical structure and bulk chemical composition of a biomaterial must be the first consideration when designing a scaffold.1-2 The fabrication techniques of scaffolds with desired physical structure, such as electrospinning, solvent casting, freeze-drying, melt-based technologies, and high-pressure-based methods are reasonably commonplace.1, 3 Recently, porous polymers templated from high internal phase emulsions (polyHIPEs) are particularly attractive scaffolds for 3D cell growth.4 HIPEs are commonly known as highly concentrated emulsions with the volume fraction of the internal phase exceeding 74%.5 Fabrication of polyHIPEs involves the polymerization or cross-linking of the external (non-droplet) phase and the subsequent extraction of the dispersed phase.6-8 Approaches in this respect give well-defined hierarchical structures, high material porosities (typically > 85%) as well as high pore interconnections, which help to attain desired mechanical function to support the weight of large cell numbers and facilitate mass transport to cells growing in the materials.6,

9-10

Furthermore, the scaffolds can be

processed into desired 3D anatomical shapes by introducing the polymerization reaction of the asprepared HIPEs into predefined molds. The most investigated polyHIPEs are originated from the surfactant-stabilized water-in-oil (W/O) HIPEs with styrene (STY), divinylbenzene (DVB), and 2-ethylhexyl acrylate (EHA) as monomers in the organic phase. Those polymers are hydrophobic and scarcely or at all biodegradable. The obtained polyHIPEs consequently need surface modification to enhance the hydrophilicity aiming at biomedical applications.9-12 Therefore, there has been increasing studies addressing hydrophilicity and biocompatibility of the scaffolds by introducing O/W HIPE templates and natural starting polymers. However, O/W HIPEs always require more careful emulsion stabilization and polymerization,13 which by far leads to few relative approaches, just including several methodologies for the production of


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monolithic polyHIPEs from 2-hydroxyethyl methacrylate,13 2-hydroxyethyl acrylate,14 acrylamide,15-16 as well as co-monomers.17 Natural polymer like gelatin has been introduced in preparing polyHIPEs, also following the thoughts of free radical polymerization from vinylated or methacrylated gelatin in the continuous phase.2, 18 In addition, the above methods usually involve tedious polymerization process and massive surfactants (like Triton X-405, net 5 ~ 50 vol%). Previous studies have reported on the potential cytotoxicity of several surfactants over a certain concentration, and revealed that they may cause chronic toxicity to many kinds of aquatic organisms.19-22 Thus, an alternative strategy to stabilise HIPEs is based on replacing the surfactant altogether by using small particles. Particle-stabilized emulsions, also known as Pickering emulisons, exhibit excellent droplet stabilization due to the nearly irreversible adsorption of colloid particles with their high energy of attachment at the oil-water interface.23 Therefore, Pickering HIPE templates can be extremely stable against coalescence during the polymerization process and hence commendably maintain the defined porous structure of the resulting polyHIPEs. To mimic the biological (extracellular matrix) environment, it is desirable to make use of biopolymers as the constituents of the polyHIPEs. Gelatin, which is the denatured-type collagen derived from the controlled hydrolysis of the native collagen, is a commonly used natural macromolecule in medical fields and tissue engineering.24-25 Gelatin has relatively low antigenicity whilst maintaining many integrin-binding sites for cell adhesion and differentiation.26 More importantly, gelatin has high emulsifying capacity, and it can form a monomolecular layer of protein around the oil droplets, thus stabilizing the emulsions. Our work therefore describes the employ of gelatin as stabilizers for HIPEs preparation, simultaneously being the major constituents for the generation of porous scaffolds. Recently, we also synthesized a new protein nanoparticle by a two-step desolvation method using gelatin as starting material (Scheme 1). Considering the fairly monodispersed gelatin nanoparticles were ideal candidates for Pickering stabilizers, herein we introduced them as co-stabilizers with gelatin to prepare HIPE templates. An interesting finding in our preliminary experiments was that very small


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amount of gelatin nanoparticles could improve the emulsion stability. The nanoparticles irreversibly adsorbed at the oil-water interface to form viscoelastic stabilizing layers, which could strongly hinder droplet coalescence and enhance the emulsion stability during cross-linking reaction.9 Chemically crosslinking the continuous phase and the subsequent extraction of the dispersed phase led to the formation of stable polyHIPE scaffolds (Scheme 1). In addition, the in vitro biodegradation and cell culture assay of the scaffolds were also investigated. The aim of this study is to propose a facile route to prepare biocompatible natural polyHIPEs with hierarchical and highly interconnected porous structures, rendering them suitable as scaffolds for promising biomedical applications.

Scheme 1. Schematic representation of the preparation of a hierarchical porous protein scaffold.

EXPERIMENTAL SECTION Materials. Gelatin type B (~250 g bloom) and Cell Counting Kit-8 (CCK-8) were obtained from Aladdin Chemistry Co., Ltd. Genipin was kindly supplied by Lin chuan zhi xin biotechnology company (Jiangxi, China). Dulbecco’s modified Eagle’s medium (DMEM), fetal bovine serum (FBS), trypsin, glutamine, and penicillin-streptomycin were obtained from Gibco (Grand Island, USA). Thermolysin was purchased from Sigma-Aldrich. L929 cells were purchased from American Type Culture Collection


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(ATCC), and the cells were grown at 37 °C, 5% CO2 in DMEM complete medium, supplemented with 10% FBS, penicillin, and streptomycin (both at 100 units/mL). All other chemicals were of analytical grade and were obtained commercially from Kelong Chemical Reagent (Chengdu, China). Distilled water was freshly prepared and used for all the experiments. Preparation of Gelatin Nanoparticles. The synthesis of gelatin nanoparticles has been described in our previous work in detail.27 Briefly, 1.25 g of gelatin type B was dissolved in 25 mL of distilled water to prepare a gelatin solution under constant heating (50 °C) and stirring. Then, 25 mL of desolvating agent, acetone, was added to precipitate the high molecular weight (HMW) gelatin. After that, the HMW gelatin was collected and re-dissolved in 25 mL of distilled water with the pH of the solution adjusted to 12.0. A total of 75 mL of acetone was added drop-wise into the gelatin solution, followed by the addition of 125 µL of glutaraldehyde solution (25% aqueous solution) as the cross-linking agent. The resulting mixture was stirred at 50 °C for 3 h to form gelatin nanoparticles. Finally, the dispersion was centrifuged at 10 000 g (Thermo ST16R, USA) for 40 min, and the nanoparticles were purified by threefold centrifugation and redispersion in an aqueous acetone mixture (30 vol%, acetone). After the last redispersion, the remaining acetone was evaporated using a rotary evaporator (EYELA, Japan) and the resultant nanoparticles were stored at 4 °C for further experiments. Preparation of the HIPE Templates. To understand the influence of each emulsifier on the resultant emulsion templated polymers, we varied either gelatin nanoparticle content or gelatin molecules content but kept the volume fraction of the internal phase constant (80 vol%). For preparing a typical HIPE template, a total of 2 mL of aqueous phase containing certain volume of gelatin nanoparticles (18 mg/mL) or gelatin solution (10 wt%) was transferred into glass vials followed by adding 8 mL of hexane (oil phase). And then, the mixture was mechanically sheared with an Ultra Turrax T18 homogenizer (IKA, 10 mm head) operating at 12000 rpm for 30 s to form HIPEs. A series of HIPE were stabilized by gelatin nanoparticles and gelatin with mass ratio (g/g) of 0.25:1 (M1), 0.4:1 (M2), 0.68:1 (M3), and 1.5:1 (M4), and a HIPE solely stabilized by 18 mg/mL gelatin nanoparticles was prepared as


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control (M0). Preparation of Porous Scaffolds Templated from the HIPEs. To cross-link the continuous phase of the HIPE templates, 50 µL of genipin (cross-linker) solution (1 wt%) was mixed with 1.95 mL of mixture with certain volume of gelatin nanoparticles (18 mg/mL) and gelatin solution (10 wt%). Immediately, the mixture was mechanically homogenized with 8 mL of hexane to obtain HIPE templates. Set the HIPEs at room temperature for 72 h to let the genipin fully react with the gelatin molecules and gelatin nanoparticles. After that, the resulting monoliths were alternately washed with ethanol and distilled water to remove the internal phase, followed by the freeze-drying to yield polyHIPEs. Scanning Electron Microscopy (SEM). To observe the morphology and structure of the gelatin nanoparticles, SEM images were carried out using a Quanta 400 F (FEI Company, USA) operated at 15 kV. A drop of dilute gelatin nanoparticle dispersion was placed on a silica wafer and dried in air for 12 hours, and the samples were sputtered with gold for 3 minutes in an argon atmosphere before observation. The cross-sectional morphologies of the macroporous scaffolds were observed using a FEI Quanta 400 F microscope operating at 10 kV. The monoliths were cut with a knife and coated with Au nanoparticles before imaging, and all the samples were placed onto carbon-coated lacy substrates. Contact Angle Measurement. The water-in-oil contact angles θow were measured at 25 °C using a DataPhysics OCAH200 (Germany) setup by the captive drop method. Microscopic slides have been treated with gelatin nanoparticles and gelatin with mass ratio of 1:0, 1.5:1, and 0.25:1. A drop of water was deposited on the pre-treated slides immersed in hexane with a needle, and the drop shape was recorded with a camera and analyzed for the contact angle. Confocal Laser Scanning Microscopy (CLSM). CLSM pictures of the HIPEs were taken on a Nikon Eclipse Ti inverted microscope (Nikon, Japan) using a 543 nm laser to excite the emulsions. Note that the gelatin nanoparticles prepared in our study were autofluorescent, and they exhibited red fluorescence


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when excited by the commonly used 543 nm laser. The average droplet size was determined by manually measuring over 150 droplets from the CLSM images using image analysis software (Nikon EZ-C1 FreeViewer). Rheology Measurement. The rheology measurement of the resulting emulsions was carried out at room temperature with a rheometer (Anton Paar Physica MCR 301) by a plate-plate mode PP 25. The strain sweep was performed with frequency set at ƒ =1 Hz, and the temperature was carefully controlled with a water bath. In Vitro Biodegradation. The in vitro biodegradation of the scaffolds was conducted with minor modifications to previously described methods.2 The freeze-dried scaffolds were incubated in a thermolysin solution (0.02 mg/mL in 0.05 M Tris-HCl, 2.5 mM CaCl2, pH = 7.4) maintained at 37 °C under mild stirring. At certain time intervals, solution aliquots were measured at 280 nm via a UV−visible spectrometer (Shimadzu, UV-2000) until the solid monoliths were completely dissolved. The percentage of scaffold degradation was determined by the equation as follows: Scaffold degraded (%) =

A280(t ) A280(tfin )

(1)

Where A280(t) represented solution absorbance at time t, A280(tfin) was the plateau absorbance corresponding to the completely dissolved scaffolds. Indirect Cytotoxicity Assay. The viability of L929 fibroblast cells exposed to the extraction liquid of the scaffolds was determined by the colorimetric CCK-8 assay. Briefly, scaffold pieces (10 mm diameter, 0.5 ± 0.1 mm thickness) were sterilized with UV radiation from both sides for 30 min each. Then, they were incubated at 37 °C in DEME supplemented with 10% fetal bovine serum and 1% antibiotic/antimycotic for 24 h to obtain the extraction medium. 100 µL of L929 fibroblasts suspension was seeded in a 96-well plate (2 × 103 per well) and cultured in a 5% CO2 humidified incubator at 37 °C for 24 h. After that, the culture medium was removed, replaced with the corresponding extraction medium, and the cells were further incubated. The cells cultured with fresh medium was used as control.


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At time intervals of 24, 48, and 72 h, 100 µL of CCK-8 (10%, v/v) solution in fresh medium was added to each well and incubated at 37 °C for 2.5 h. Finally, the absorbance values were recorded with a microplate reader (BioTek Instruments Inc. ELX 800, USA) at 450 nm. The cell viability was expressed as a percentage of the absorbance of control, repeated for five times of each sample. Furthermore, the cell morphology was observed by using an optical microscope (OLYMPUS CKX53, Japan) at time intervals of 24, 48, and 72 h. Cell Morphology Study on Scaffold. L929 fibroblast cells (1 × 105 per well) were seeded in 24-well culture plates containing sterilized swollen scaffold pieces (12 mm diameter, 0.5 ± 0.1mm thickness), and then cultured at 37 °C in a wet atmosphere in the presence of 5% CO2 for 72 h. For analyzing the morphology and proliferation of cells in the scaffolds, cells were fixed with 2.5% glutaraldehyde in pH 7.5 phosphate buffer, dehydrated in increasing concentrations of ethanol from 30 to 100%. Samples were then freeze-dried and sputter-coated with gold prior to observation on a JEOL (JSM-7500F, Japan) SEM operating at 10 kV.

RESULTS AND DISCUSSION PolyHIPEs Templated from Gelatin-Stabilized HIPEs. The emulsifying power of gelatin is owing to the side chains which have charged groups and that certain parts of the collagen sequence contain either hydrophilic or hydrophobic amino acids.28 Both hydrophilic and hydrophobic parts tend to migrate to oil-water surface and form a layer of protein around the oil droplets, thus stabilizing the emulsions. The HIPEs can be successfully prepared by using 2.5 wt %, 5 wt %, and 10 wt% gelatin as emulsifiers (Figure S1 in the Supporting Information). The typical HIPE shown in Figure 1A reveals that it is a kind of gel-like emulsion and shows very little phase separation even after one month. In addition to decrease the surface tension of aqueous systems, gelatin can form required films around the droplets, strengthening the dispersed phase by gel formation. Moreover, it can also increase the viscosity of the aqueous phase, trapping the oil droplets in the gel matrix and finally resulting very stable HIPEs.


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It seems that the gelatin-stabilized HIPEs are perfect templates for scaffold preparation based on the characteristic of natural derivative, extreme stability, and biocompatibility. We therefore use a naturally occurring cross-linking agent, genipin, to cross-link the dispersed phase to form a permanent pure protein scaffold. The microstructure of the resultant polymer was determined by SEM, as shown in Figure 1B. Nevertheless, the polymeric monolith presents random morphology with poor interconnectivity, which is not accordance with the usually reported interconnected polyHIPEs with a hierarchical pore distribution. In fact, the unique pore structure of polyHIPEs is determined by the thermodynamics of polymer gel phase separation from the continuous phase and the delicate equilibrium of emulsion phase separation versus droplet stabilization.11 It is proposed here that the cross-linking reaction proceeding in the continuous phase as well as on the interface dramatically affect the stabilization of the HIPE templates, leading to a large-scale collapse of the droplets and the subsequent failure of structural integrity. As a consequence, HIPE solely stabilized by gelatin is failed to be used as template to fabricate highly interconnected porous scaffolds.

Figure 1. (A) Photograph of HIPE stabilized with 10 wt% gelatin taken one month after preparation and (B) the SEM image of the corresponding cross-linked polymers.


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Figure 2. (A) SEM image of the synthesized gelatin nanoparticles. Water-in-oil contact angles θow (see insets) of mixed gelatin nanoparticles and gelatin with mass ratio of (B) 1:0, (C)1.5:1, and (D) 0.25:1 (scale bar is 250 µm).

Characterization of HIPEs co-stabilized by gelatin and gelatin nanoparticles. It has been extensively demonstrated that Pickering emulsion is rather stable for several months or even years. Based on the facts, we now consider introducing solid colloidal nanoparticles, namely gelatin nanoparticles, to serve as co-stabilizers to obtain stable HIPE templates resisting the droplet rapture caused by cross-linking reaction. As shown in Figure 2A and Figure 2B, gelatin nanoparticles are ideal candidates for Pickering stabilizers with a fairly monodispersed average size of 235.9 ± 2.2 nm (PDI = 0.119) and suitable intermediate wettability according to our previous work27,

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. The interfacial

entrapment of gelatin nanoparticles is mainly determined by the contact angle (θ) of the particles at the water-oil interface — surface wettability. As for co-stabilizers, the absorption of molecular surfactants may change the wetting properties of colloidal particles.30 We therefore carried out a wetting experiment on a model macroscopic system to investigate the effect and to explore the ability of gelatin ACS Paragon Plus Environment

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and gelatin nanoparticles to stabilize emulsions. Figure 2B and Figure 2D reveal that adding 40 wt% gelatin of the total stabilizers to the aqueous phase increases the contact angle measured through bare gelatin nanoparticles from θ = 25 ± 3° to θ = 31 ± 2°, and then increases to θ = 37 ± 2°in the presence of 80 wt% gelatin. The results indicate a significant change of wettability of gelatin nanoparticles with a slight preference for wetting by the oil phase. It is proven that gelatin nanoparticles are autofluorescent and they exhibit red fluorescence when excited by the commonly used 543 nm laser.27 The formation of CH = N bonds from Schiff base in conjunction with the C = C double bonds from the glutaraldehyde was responsible for the autofluorescence of the particles.31 This feature provides great convenience for observing the behavior of gelatin nanoparticles on the oil-water interface as well as in the continuous phase. Figure 3 shows the CLSM images of the HIPEs stabilized by different content of gelatin and gelatin nanoparticles. The results reveal as expected that gelatin nanoparticles migrate to the oil-water interface to form thick packing layers (Figure 3A). As for co-stabilizers (Figure 3B-E), gelatin and gelatin nanoparticles tend to form large aggregates and agglomerates either at oil-water interface or in the aqueous phase, and the number of which decreases with the increase of gelatin nanoparticles. In the so crowded continuous phase of the HIPEs, gelatin molecules are easy to be adsorbed at the surface of gelatin nanoparticles probably via hydrogen bonds, thus forming aggregates. Furthermore, it seems that upon increasing the content of gelatin nanoparticles leads to smaller and more uniform aggregates, because the welldispersed gelatin nanoparticles dramatically disaggregate the gelatin molecules and impart steric repulsion, thereby leaving them well dispersed in the continuous phase.


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Figure 3. CLSM images of HIPEs co-stabilized by gelatin nanoparticles and gelatin with mass ratio (g/g) of 1:0 (A), 0.25:1 (B), 0.4:1 (C), 0.68:1 (D), 1.5:1 (E), and the magnification image of E (F). The arrows represent the gaps formed between the adjacent droplets.

Stabilization of the HIPEs is achieved when gelatin, gelatin nanoparticles, and the aggregates diffuse to the interfacial region and remain there in a stable mechanical equilibrium. Thus, the packing layers residing at the interface provide a physical barrier to emulsion droplet rupture and coalescence. It ACS Paragon Plus Environment

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has also found that aggregates of particles can be transported to the interface of droplets more easily than single nanoparticle during high-shear mixing because of their larger relative inertia for adsorption. At the same time, large aggregates have small radii of curvature that approaches the oil-water interface and thus a lower energy barrier for adsorption.32-33 The nanoparticles to be in a state of weak aggregation in the aqueous phase could create the most stable emulsions mainly owing to the enhanced viscosity of the continuous aqueous phase,34 which provides sufficient stability and mechanical support for withstanding experimental procedures in the generation of scaffolds. Particularly, as shown in Figure 3F, the random small aggregates transported to the interface lead to the formation of some gaps between the droplets, which is completely different from the dense packing particle layers around the droplets in the case of HIPE solely stabilized by gelatin nanoparticles (Figure 3A).

Figure 4. Droplet size distributions of HIPEs co-stabilized by gelatin nanoparticles and gelatin with mass ratio (g/g) of 0.25:1 (M1), 0.4:1 (M2), 0.68:1 (M3), 1.5:1 (M4).


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The stabilizer content set the amount of available stabilized interface at a fixed volume fraction of the oil phase — meaning out of little stabilizers have less opportunity to transport to the interface to stabilize the emulsions, thereby increasing the droplet size.29, 35 As for M1-M4, the total content of gelatin and gelatin nanoparticles, namely stabilizers, is gradually decreased as follows: 0.1 g, 0.834 g, 0.067 g, and 0.051 g. The CLSM images have shown a clear tendency that a decrease in stabilizers concentration results in an increase in droplet size. We further analyze the droplet size distribution of the HIPEs using the incorporated image software in the CLSM. As shown in Figure 4, the main approximately calculated droplet size for all the HIPEs is in the range of 5 ~ 10 µm, and this ratio decreases with the increase of gelatin nanoparticles, which is also observed for the presence of oil droplets < 5 µm. Conversely, the number of large droplets in the range of 15 ~ 25 µm increases as the content of gelatin nanoparticles increases in the emulsions. This is significantly important for improving the pore size of the polyHIPEs since the emulsion structure at the gel point of polymerization is the template for the pore structure of the resultant polymers.36 More importantly, compared with M1, the amount of stabilizers used in M4 has been reduced by half, highlighting the ability of gelatin nanoparticles to stabilize the emulsions. Rheological Measurement. The rheology of M1-M4 has been carried at a strain sweep spectrum at 1 Hz. Figure 5A reveals that both storage (G′) and loss (G″) moduli are constant in a very wide strain range, and G′ is always higher than the corresponding G″ below 40% strain. This indicates that the HIPEs behave like elastic media in this domain owing to the packing of gelatin and gelatin nanoparticles into a jammed conformation. However, upon increasing applied strain, a solid-like to liquid-like transition takes place (G′ exhibits a distinct decrease with G′ < G″), showing shear thinning. It probably relates to a large scale of structure rearrangement or flow of the dispersed droplets at high applied strains.37 Moreover, G′ and G″ increase with the increase of stabilizers below the yield value (G′ = G″), revealing that higher content of stabilizers enhancing the viscoelastic layer formed at the


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oil-water interfaces, thus leading to more stable and solid-like HIPEs.

Figure 5. (A) Strain dependence of the storage (G′) and loss (G″) moduli of M1, M2, M3, and M4 measured at a frequency of 1 Hz. (B) The strain dependence of the storage G′ and loss G″ moduli of M4 without and with genipin crosslinking, measured at frequency f = 1 Hz.

We further investigate the rheology of the typical HIPE, M4, before and after the cross-linking


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reaction at frequency 1 Hz, as shown in Figure 5B. After cross-linking, both G′ and G″ increase, especially for G ″ ， which indicates that the cross-linking indeed strengthens the stability and mechanical properties of the resultant polyHIPE. To better understand the effect of cross-linking on the viscoelastic properties of the HIPEs, we also compare the rheology property of the cross-linked M4 with the most solid HIPE, M1. As can be seen in Figure S2 (see the Supporting Information), the crosslinked M4 has higher G′than the uncross-linked M1, which further confirms that the cross-linking give rise to a stronger gel network. Additionally, the results also indicate that before cross-linking, the highly elastic behavior of the HIPE is attributed to the packing viscoelastic layer formed by gelatin and gelatin nanoparticles around the droplets. Nevertheless, with the addition of genipin, the packing layers at the oil-water interfaces together with molecules and nanoparticles in the continuous phase are further cross-linked, leading to form a soft composite with enhanced mechanical properties.37 Characterization of the Scaffolds. The morphological structures of the freeze-dried scaffolds are shown in Figure 6. These composites present a general high porosity and have a typical polyHIPE morphology with an average pore size around 25 µm, and interconnecting throat size in the range of 5 ~ 10 µm, which agrees well with the droplet size distribution of the emulsions shown in Figure 3 and Figure 4. More importantly, the scaffolds possess open-cell structure which is not usually seen in polymerization of Pickering HIPEs.38-39 It has been proposed that interconnecting pores are formed during the polymerization of the continuous phase. Concretely, the density between polymer gel phase and liquid continuous phase is generally different, and hence partial shrinkage occurs causing ruptures in the polymer film at its thinnest point.40-41 Moreover, the gaps formed around the droplets (see Figure 2) also benefits triggering the collapse and rupture of the droplets during cross-linking, leading to form open-cell pores. A tiny increase in void diameter, whereas a distinct decrease in thickness of the pore walls (see the Supporting Information, Figure S3) are observed with decreasing stabilizer content. Moreover, smooth pore walls and textured structures are also observed in the magnification images, which are quite different from the polymers templated from inorganic particle-stabilized HIPEs or ACS Paragon Plus Environment

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conventional polyHIPEs. Presumably, such smooth pore walls likely derive from the interfacial adsorbed soft nanoparticle layers and gelatin molecules separating the oil and aqueous phase. As we know, gelatin is derived from the hydrolysis of collagen and can imitate the ECM for biocompatibility.42 As a consequence, these highly gelatin-decorated surface may introduce extra benefits to the prepared scaffolds including, for example, cytocompatibility,43 cell differentiation, and mineralization.44

Figure 6. SEM images of freeze-dried porous scaffolds template from HIPEs of M1 (A), M2 (B), M3 (C), and M4 (D).

We further performed a controlled experiment concerning the scaffold templated from HIPE solely stabilized by gelatin nanoparticles. As can be seen in Figure S4 (see the Supporting Information), the composite almost retains the original morphology of the droplets, but it does not have intact pore walls and pore throat structures that are commonly seen in polyHIPEs. The preparation of gelatin nanoparticles has consumed partial amine groups of gelatin molecules; the content of residual amine


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groups therefore is not enough to form stable polymer networks with genipin to strengthen the HIPE as well as retain the integrity of the resultant porous structures. Generally, only HIPEs co-stabilized with gelatin and gelatin nanoparticles are successful templates to prepare interconnected porous scaffolds. Concretely speaking, gelatin can provide sufficient mechanical support for withstanding experimental procedures in the generation of scaffolds, while gelatin nanoparticles give high emulsion stability to guarantee the integrity of the emulsion templates during cross-linking reaction.

Figure 7. Typical degradation kinetics of pure gelatin, M3, and M4 composite scaffolds. The solubilization was catalyzed by thermolysin at 37 °C.

As for tissue engineering application, the scaffolds should be degraded in a physiological environment to allow the proliferating cells to gradually replace the scaffold itself and allow optimal regeneration.2 Hence, the degradation behavior of the pure gelatin, M3, and M4 composite scaffolds were typically monitored with a spectrophotometric assay that measured the appearance of solubilized hydrolysis products during the reaction with proteinases under cell-free conditions. Note that the concentration of the proteinase used was much higher than that normally found in the human body. As


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shown in Figure 7, the degradation ratio of all the samples increases with the increase in the soaking time, following the order: pure gelatin composite < M4 < M3. The rate of degradation is strictly connected to the degree of porosity, and a high porosity can reduce the risk of accumulation of acidic degradation products, thus reducing any possible further degradation reaction.1 Compared with pure gelatin composite, M3 and M4 have highly interconnected open-cell structures, which benefit improving the interaction with lipases, thus producing faster reaction velocities. In addition, gelatin nanoparticles cross-linked by glutaraldehyde has stable network structure with high cross-linking density, suggesting that the content of gelatin nanoparticles also affects the degradation of the scaffolds. As for M3 and M4, the content of gelatin nanoparticles in M4 is 18% (w/w) higher than M3, thus making it more resistant to enzymatic degradation. Furthermore, this also explains the phenomenon that the time reached a maximum rate of degradation of 100% is in the following order: pure gelatin composite < M3 < M4. Cell Culture Assay. The indirect cytoxicity of the prepared scaffolds analyzed by CCK-8 assay is used to evaluate the release of cytotoxic compounds from the scaffolds to the extraction medium, including the possible soluble components like genipin or glutaraldehyde. Cell viability on the scaffolds was examined for 24, 48, and 72 h using L929 cell line (Figure 8A). All the samples show a rapid cell growth during the first 24 h of culture, followed by a stationary phase. M4 present a significant increase in cell number and finally exhibit higher levels of cell viability when compared to the other two scaffolds. As a supplement, the general external morphology of L929 cells exposed to the extraction liquid of M4 was investigated by optical microscopy, as shown in Figure 8B. The treatment group shows no indication of a change in morphology and cell number compared to the control group, which further indicates that the scaffold does not induce any cytotoxic effects on L929 mouse fibroblast cells.


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Figure 8. (A) Indirect cytotoxicity evaluation of M2-M4 scaffolds based on the viability of fibroblasts (L929) cultured in the extraction media. (B) The viability of L929 cells cultured with extraction media


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of scaffolds, as relative percentage of control, i.e., fresh medium (100%).

Figure 9. SEM image (A) and the corresponding magnification image (B) of L929 fibroblasts cultured on M4 scaffold for 72 h.

The viability of the cells is greatly influenced by the cell adhesion processes, and the ability of adherent cells to spread has important consequences. Cell-substrate contact area can determine whether or not a cell proliferates, becomes quiescent or dies.45-46 In present work, the morphology and proliferation of cells in the scaffolds were investigated by SEM. Figure 9 displays the SEM images illustrating the adhesion and spreading of L929 cells seeded on M4 at incubation time of 72 h, as an example. It shows that the cells exhibit intensive growth that partially covered the scaffold surfaces, and have a well-spread morphology with multiple cellular extentions adhering to the substrate. These results reveal that the composite scaffolds templated from HIPEs promotes the L929 mouse fibroblast cell adhesion and proliferation, which thus demonstrates their potential application in tissue engineering.

CONCLUSIONS Hierarchical porous protein scaffold can be fabricated by cross-linking of O/W HIPEs co-stabilized with gelatin and gelatin nanoparticles. Gelatin can provide sufficient mechanical support for withstanding experimental procedures in the generation of scaffolds, while gelatin nanoparticles give high emulsion stability to guarantee the integrity of the emulsion templates during cross-linking reaction. Behind this, ACS Paragon Plus Environment

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gelatin molecules tend to be adsorbed on the surface of gelatin nanoparticles to form large aggregates and agglomerates, changing the wetting properties of gelatin nanoparticles, thus forming emulsion droplets with different size distribution. The rheological properties of the HIPEs can be tuned within a certain range by adjusting the stabilizer amounts. Cross-linking the dispersed phase not only strengthens the viscoelastic properties of the resultant polyHIPEs but also retains the integrity of the hierarchical porous structures. The freeze-dried scaffolds present a general high porosity and have an interconnected porous morphology with smooth pore walls and textured structures. Moreover, the scaffolds are degradable and the rate of degradation is strictly connected to the degree of interconnectivity and the content of gelatin nanoparticles. The scaffolds show reasonably good biocompatibility; L929 cells can adhere to the surface of the materials and exhibit intensive growth and well-spread morphology. Therefore, our fabrication technique paved the way toward novel hierarchical porous scaffold production using a facile process, and resultant scaffolds are based on fully natural materials, showing the possibility to be used as culture matrices for soft tissue engineering.


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ASSOCIATED CONTENT Supporting Information. Digital pictures of HIPEs stabilized by different concentration of gelatin; rheological properties of the cross-linked scaffolds; SEM imges of the macroporous scaffolds.

AUTHOR INFORMATION Corresponding authors *(Xiaojun Gou) Tel: +86-028-84216578; Fax: +86-028-84333218; E-mail: [email protected]. *(To Ngai) Tel: (+852) 39431222; Fax: (+852) 2603 5057; E-mail: [email protected].

ORCID Xiaojun Gou: 0000-0003-3211-3985 To Ngai: 0000-0002-7207-6878

Notes The authors declare no competing financial interest.

ACKNOWLEDGMENTS

The financial support of Scientific Research Foundation of Sichuan Education Department (17ZB0102), and Youth Fund Program of Chengdu University (2017XJZ01) are gratefully acknowledged. We also acknowledge the fudding from the Collaboration and Innovation on New Antibiotic Development and Industrialization (2016-XT00-00023-GX) for financial support. T. N. acknowledges the supports from the Direct Grant for Research (3132681, 3132682 & 4053195) of the Chinese University of Hong Kong. Furthermore, we greatly appreciate the contact angle measurement done by Wanbo Xue from Prof. Wei Lin’s group in Sichuan University.


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Table of Contents (TOC) Graphic


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