Toward Structured Macroporous Hydrogel Composites: Electron Beam

Article pubs.acs.org/Biomac

Toward Structured Macroporous Hydrogel Composites: Electron Beam-Initiated Polymerization of Layered Cryogels Anna Golunova,† David Chvátil,‡ Pavel Krist,‡ Josef Jaroš,§ Veronika Jurtíková,§ Jakub Pospíšil,§ Ilya Kotelnikov,† Lucie Abelová,† Jiří Kotek,† Tomás ̌ Sedlačík,† Jan Kučka,†,‡ Jana Koubková,† Hana Studenovská,† Libor Streit,∥ Aleš Hampl,§ František Rypácě k,† and Vladimír Proks*,† †

Institute of Macromolecular Chemistry, Academy of Sciences of the Czech Republic, Heyrovsky Square 2, 162 06 Prague 6, Czech Republic ‡ Nuclear Physics Institute, Academy of Sciences of the Czech Republic, Ř ež 130, 250 68 Ř ež, Czech Republic § Department of Histology and Embryology, Faculty of Medicine, Masaryk University, Kamenice 5, 625 00 Brno, Czech Republic ∥ Department of Plastic and Aesthetic Surgery, St. Anne University Hospital Brno, Pekarska 53, 656 91 Brno, Czech Republic S Supporting Information *

ABSTRACT: The ability to tailor mechanical properties and architecture is crucial in creating macroporous hydrogel scaffolds for tissue engineering. In the present work, a technique for the modification of the pore size and stiffness of acrylamide-based cryogels is demonstrated via the regulation of an electron beam irradiation dose. The samples were characterized by equilibrium swelling measurements, light and scanning electron microscopy, mercury porosimetry, Brunauer−Emmett−Teller surface area analysis, and stiffness measurements. Their properties were compared to cryogels prepared by a standard redox-initiated radical polymerization. A 125I radiolabeled azidopentanoyl-GGGRGDSGGGY-NH2 peptide was bound to the surface to determine the concentration of the adhesive sites available for biomimetic modification. The functionality of the prepared substrates was evaluated by in vitro cultivation of adipose-derived stem cells. Moreover, the feasibility of preparing layered cryogels was demonstrated. This may be the key to the future preparation of complex hydrogelbased scaffolds to mimic the extracellular microenvironment in a wide range of applications.

■

INTRODUCTION Hydrogels are water-insoluble, chemically or physically crosslinked, three-dimensional polymer networks composed of hydrophilic or water-swellable polymers.1 These networks are able to swell and retain a significant amount of water when placed in an aqueous solution.1 Hydrogel materials are widely used in various biomedical and tissue engineering applications, including soft contact lenses,2,3 drug-releasing implants,4 drug delivery systems,5,6 wound-healing matrices,7−10 and cell cultures.11−14 The materials used in cell and tissue engineering should support cell adhesion, proliferation and differentiation. The ways to reach this aim typically include tuning the mechanical properties of the material according to the desired type of tissue.15 For example, studies of glial and neural cell,14 muscle16 and bone tissue15 growth have demonstrated the correlation between biomaterial stiffness and cell behavior. The incorporation of biomimetic motifs, which can support cell adhesion and direct differentiation, is another important feature for an artificial matrix. Peptides derived from various extracellular matrix proteins are widely used for this aim. For instance, a hydrogel matrix can be modified with the fibronectin-derived RGD peptide15,17 or with special functional groups to enable further biomimetic surface modification.18−21 © XXXX American Chemical Society

Moreover, the stem cells differentiation into the desired tissue phenotype can also be stimulated by the 3D microenvironment surrounding of the cells.22,23 In the case of crosslinked hydrogels, suitable matrix architecture can be created by conducting polymerization in the presence of a porogen. Any solid, low molecular weight compound, even a polymer that is not soluble in the reaction mixture but can be easily solubilized and leached out with other solvents, is utilized, e.g., inorganic salts24 or fibrous polymers that are decomposed to soluble degradation products.17,25 Obtained in a such way macroporous materials were utilized for various tissue engineering applications.17,18,25−27 Cryogels represent a group of macroporous hydrogels with interconnected pores.28−33 The preparation of these materials is carried out under freezing conditions and crystals of the frozen solvent act as porogen. The main advantage of this method is in producing gels with a high swelling capacity and spongy structure while preserving good mechanical stability.34−36 Received: December 15, 2014 Revised: February 24, 2015

A

DOI: 10.1021/bm501809t Biomacromolecules XXXX, XXX, XXX−XXX

Article

Biomacromolecules

Microscopy observations were performed using Olympus IX71 Cell∧R imaging station (Olympus C&S Ltd., Prague, Czech Republic) with the objective LUCplan FL N 20x/0.45 and images captured by a Hamamatsu ORCA-R2 digital camera. Formation of Hydrogels Using a Redox Initiation System. Hydrogel formation was carried out in 5 mL glass vials (ampules). Appropriate amounts of acrylamide (AAm), N,N′-methylenebis(acrylamide) as a cross-linking agent and N-propargylacrylamide (PrAAm) as a functional comonomer were dissolved in Q-water to obtain 7.5 w/v% solutions with different monomer compositions. A portion of the initial solution was added to each vial; the solution was then purged with nitrogen and brought to a temperature near zero. TEMED was then added. APS (0.02 g/mL) was subsequently added at 2 w/w% to all of the solids. The vials were placed into a thermostat, where a temperature of −15 °C was maintained. Then, all of the vials with frozen solutions were thermostated at −18 °C for 12 h. After freezing, the samples were defrosted and rinsed with distilled water. The washed samples were dried in a freeze-dryer until a constant weight was reached. Formation of Hydrogels Using Irradiation. The formation of hydrogels using irradiation was performed similarly to the procedure described above, except for the addition during the initiator stage. After the samples were incubated at −15 °C, the initiation process using irradiation took place. The irradiated samples were maintained under subzero conditions (−15 °C) using an MT 25 microtron. The absorbed dose was regulated according to the experimental conditions. To estimate the yields, the samples were weighed on an analytical balance (Sartorius GmbH, Germany). Formation of Layered Hydrogels. A layered structure was formed by sequentially adding solution to the previous layer while incubating the whole vial at a subzero temperature. After the added layer had frozen, the process was repeated until the reaction vial was filled. The layers were distinguished by the presence or absence of 5 mol % propargylacrylamide with respect to the acrylamide component (3.2 mol % with respect to all of the solids). The frozen samples were irradiated under the same conditions used for the nonlayered gels, as described above. After thawing and washing, compact bulk gels were obtained. Electron Beam Irradiation of the Cryogel. The MT 25 microtron is a cyclic relativistic accelerator of electrons located in Prague in the NPI of the ASCR, v.v.i. The main parameters of the microtron are summarized in Supporting Information Table S1, and a schematic is shown in Figures S2 and S3. Water Uptake Measurement. Dried hydrogels were weighed and then swollen at room temperature in Q-water until equilibrium was reached. At least five measurements were performed for each hydrogel sample, and the average weight of the swollen hydrogels was calculated. The value of equilibrium water content was determined using eq 1.

A wide and diverse range of polymer compositions has been used to fabricate hydrogels.37 Among others, there are materials based on acrylamide/N,N′-methylenebis(acrylamide) monomers that have shown extracellular matrix (ECM)-like mechanical properties38 that have already been used in tissue engineering applications.39,40 The polyacrylamide gels are primarily obtained by free radical polymerization. In general, the ammonium persulfate/ N,N,N′,N′-tetramethylethylenediamine (APS/TEMED) system is used for initiation.28,41,42 The alternative route for initiation of polymerization process is realized with gamma or beta beams or other types of irradiation.10,43−45 The present work is focused on the improvement of recently proposed electron-beam initiated polymerization of cryogels,46,47 tailoring their mechanical properties, and comparison with hydrogels obtained by the standard red-ox initiation mechanism. Moreover, alkyne groups were incorporated into the hydrogel composition. They can be used for the biomimetic surface modification by the click-chemistry.48 In the present work, the concentration of these groups was determined by the radioassay. The interaction of scaffold with cells was performed using adipose-derived stem cells (ADSC) to understand their response to different hydrogel chemical composition and mechanical properties. Another goal was to explore the potential of the irradiation technique for creation of complex 3D-architectures, in particular, for the preparation of bulk cryogels composed of layers with different consist. Because of the extracellular matrix complexity,49 which even in one organ could provide growth of different types of tissue,50 the potential of using scaffolds with various domains is higher than for the homogeneous ones. The electron beam irradiation way of carrying out the polymerization process gives the possibility to obtain layered cryogels, which, in comparison to layered nonporous hydrogels,51−53 can combine a composite structure that supports the growth of different types of cells and a controlled system of macropores with all the profit it has.

■

MATERIALS AND METHODS

Acrylamide (AA), N,N′-methylenebis(acrylamide) (MBAA), copper sulfate pentahydrate (CuSO4·5 H2O), and sodium ascorbate were obtained from Sigma-Aldrich (Czech Republic). N-propargylacrylamide (PrAAm) and 125I radiolabeled azidopentanoyl-GGGRGDSGGGY-NH2 peptide were prepared according to the method of Mackova et al.,54 and 3-azido-7-hydroxycoumarin was prepared according to the method of Sivakumar et al.55 Electron-beam initiation was carried out using an MT 25 microtron instrument (NPI, ASCR v.v.i., Prague). The samples were incubated in a Ministat 2400 cryostat (Huber, Offenburg/Germany). For UV-mediated visualization of the modified layers, a thin layer chromatography (TLC) lamp (Camag, Muttenz, Switzerland) was used. The sample morphology was evaluated by light microscopy on a SZ61 stereomicroscope (Olympus) and by scanning electron microscopy (SEM) on a Vega Plus TS 5135 (Tescan, Czech Republic). A radioassay was performed using a Bqmetr 4 ionization chamber (Empos Ltd., Czech Republic). The stiffness of the hydrogels was characterized by measuring their compressive modulus using an Instron 6025R5800 (Norwood, MA, USA) universal tester. Collagenase Type IV, Dulbecco's modified Eagle's medium (DMEM; Gibco, Life Technologies, Paisley, UK), fetal bovine serum (FBS), penicillin/streptomycin (PAA Laboratories GmBH, Pasching, Austria), and gelatin were used for the cell cultivation. Acridine orange (A6014, Sigma-Aldrich) and ethidium bromide (46065, Sigma-Aldrich) were used for the biological live/dead analysis.

S=

ms − md 1 ρ md

(1)

where ms is the weight of the gel with equilibrium water content or squeezed gel (g), md is the weight of dry gel (g), and ρ is the solvent density (g/mL). The water uptake of the macroporous hydrogel network was measured by wringing out the swollen samples between several layers of filter paper until constant weight was reached. When all free water was removed from the hydrogel pores, the swelling was estimated using the equation given above. Water content adsorbed by the cryogel polymer network was estimated through the water vapor adsorption experiments according to Plieva et al.56 Briefly, dried cryogel samples were placed in a water vapor-saturated chamber with no direct contact of the sample with water. The increase in sample weight with time due to absorbed water vapor was checked after 14 days. This value gave the weight of the gel matrix with polymer bound water. Cyclohexane Uptake Measurement. Dried hydrogels were weighed and then swollen with room temperature Q-water. Swollen samples were successively washed with ethanol, acetone, and finally B


Article

Biomacromolecules

with λ = 366 nm, emitted by a TLC lamp (Camag, Muttenz, Switzerland). Mechanical Properties. The studied hydrogels were subjected to compressive testing. The stiffness of the hydrogels was characterized by the compressive modulus of elasticity. Cylindrical specimens having a h/D (height-to-diameter) ratio of ∼1/3 were subjected to compressive loading using an Instron 6025R5800 universal testing machine. The specimens had a diameter of approximately 16.5 mm and were deformed at a constant test speed of 1.0 mm/min at ambient temperature. During the test, the specimens were immersed in distilled water. The modulus of elasticity was calculated from the linear portion of the stress−strain diagram by dividing the change in stress, σ2 − σ1, by the corresponding change in strain, ε2 = 0.10 minus ε1 = 0.05. Cell Seeding. Adipose-derived stem cells were isolated from adipose tissue by centrifugation and collagenase extraction based on Zuk’s method.58 Briefly, adipose tissue was digested with 0.1% collagenase type IV for 30 min at 37 °C. Following enzyme activity neutralization by DMEM-F12 with 10% FBS, cells were separated by centrifugation at 600g. The pellet was resuspended in cultivation medium (10% FBS, 0.5% penicillin/streptomycin in DMEM Glutamax) and propagated for five passages on a culture dish coated by 0.01% gelatin. Thin layers of cryogels were cut to working pieces 3 × 3 mm and washed three times with 0.1 M phosphate buffer. Adipose-derived stem cells were released from cultivation plastic dish by trypsin digestion and seeded on cryogels in 500 uL suspension with concentration of 3 × 105 cells/ mL and incubated in 5% CO2 atm at 37 °C for 24 h. Fluorescent Staining of Viable Cells and Actin Cytoskeleton. Samples of hydrogels seeded with cells were stained with live/dead fluorescent solution for 3 min at 22 °C and observed in fluorescence microscope to examine cell viability. Fluorescent stock solution was prepared by diluting 0.03% w/v acridine orange and 0.1% w/v of ethidium bromide to 2% EtOH in distilled H2O. Final dilution was 1/ 1000 in 0.1 M phosphate buffer. Microscopy observations were performed with Olympus IX71 Cell∧R imaging station equipped with the objective LUCplan FL N 20x/0,45 and images captured by a Hamamatsu ORCA-R2 digital camera. To visualize cell adhesion and cell spreading in hydrogels, actin cytoskeleton of cells was stained. Cells grown on hydrogels 24 h were fixed with 4% paraformaldehyde dissolved in 0.1 M phosphate buffered saline (PBS), permeabilized by 0.1% Triton TX-100, and fibrillar actin was labeled with 60 nM Phalloidin Rhodamine (R415, Life technologies, Czech Republic) dissolved in 0.1 M PBS. Samples were washed twice by PBS, and cell nuclei were counterstained with 1 μg/mL 4′,6-diamidino-2-phenylindole (DAPI, 32670, Sigma-Aldrich, Czech Republic). Images of actin were obtained using confocal laser scanning microscope Fluoview 500, (Olympus C&S Ltd., Prague, Czech Republic). Scanning Electron Microscopy. Hydrogel samples were fixed in 3% glutaraldehyde (Polysciences, Inc., Warrington, USA) dissolved in 0.1 M phosphate buffer for 1 h at room temperature. Subsequently, the samples were washed three times for 10 min in 0.1 M phosphate buffer, dehydrated in ascending ethanol grade (30, 50, 70, 80, 96, 100%) and dried in a critical point dryer (CPD 030, Balzers Union Limited, Balzers, Liechtenstein) using liquid carbon dioxide. Samples were sputtered with gold in a sputter coater (SCD 040, Balzers Union Limited, Balzers, Liechtenstein) and examined in a scanning electron microscope (Vega, Tescan Orsay Holding, Brno, Czech Republic). SEM micrographs were obtained using secondary electron imaging at 15 kV.

cyclohexane. Using the solvent-exchange method, a thermodynamically good solvent (“good” solvent) in the swollen gel was replaced by a thermodynamically poor solvent (“bad” solvent). The experiment was carried out using samples prepared with redox initiation as well as different irradiation doses. SEM and Light Microscopy. An SZ61 stereomicroscope (Olympus) and a Vega TS 5135 scanning electron microscope (Tescan, Czech Republic) were employed to investigate the morphology of the hydrogels. Before observation with the electron microscope, the hydrogel samples were sputtered with a Pt layer (SCD 050 vacuum sputter coater; Leica). SEM micrographs were obtained using secondary electron imaging at 30 kV. Mercury Porosimetry and Specific Surface Area Measurement. The pore structure was characterized by mercury porosimetry on Pascal 140 and 440 instruments (Thermo Finigan, Rodano, Italy) over two pressure intervals, 0−400 kPa and 1−400 MPa, respectively. This method allowed for a determination of the meso- (4−50 nm) and macroporous structure (0.05−116 μm). The pore volume and the most frequent pore size were calculated from cumulative pore volume curves in the Pascal software using the Washburn equation and a cylindrical pore57 model. The porosity was calculated according to eq 2, where V is the cumulative pore volume (mL/g) and ρ is the density of the polymer used: P=

Vc·100 V+

1 ρ

(2)

The specific surface area of the samples was determined on a Gemini VII 2390 (Micromiretics, Instruments Corp, Norcross, USA) with nitrogen as the sorbate. The surface area was calculated from the Brunauer−Emmett−Teller (BET) adsorption/desorption isotherm using the Gemini software. Prior to the porosity and surface area measurements, the samples were vacuum-dried at 130 °C for 16 h. For all calculations, an apparent density of ρ = 0.561 g cm−3 was applied for the polymer. Modification of the Sample Surface with a Radiolabeled RGD Peptide. In this step, 0.001 g of radiolabeled RGD peptide (specific activity 30 MBq/mg) was synthesized according to the method of Mackova et al.54 A stock solution was prepared by dissolving the peptide in 1.5 mL of Q-water. Subsequently, 70 μL of sodium ascorbate (100 mg/mL) was added. Dried hydrogels (six samples) were weighed (average weight of 0.0026 g) and then swollen at room temperature in 200 μL of the stock solution in centrifuge tube filters with a cellulose acetate membrane with a pore size of 0.45 μm (Corning, Inc., USA). All samples were purged with nitrogen for 15 min to remove the oxygen, and then, 70 mL of 0.05 M CuSO4 solution was added as the second component of the click-reaction initiation system. The ratio of nonspecifically bound peptide was determined using the same reaction conditions but without the copper catalyst. The click-reaction was carried out for 30 min with nitrogen purging. Centrifugation was the next step of the procedure. The process was carried out in a Personal Microcentrifuge MiniSpin (Eppendorf AG, Germany) at 14 500 rpm. Then, all samples were washed with 200 μL of Q-water four times. The wastewater was separated by centrifugation. The radioactivity was measured with a Bqmetr 4 ionization chamber (Empos Ltd., Czech Republic). Modification of the Sample Surface with the Azidocoumarine Derivative. First, 3-azido-7-hydroxycoumarin was synthesized according to the method of Sivakumar et al.,55 weighed and dissolved in a water−ethyl alcohol mixture (1:1). The concentration of the solution was 0.3 μg/mL. Weighed hydrogels (layered and nonlayered; mass of ∼0.1 g) were placed in a filtration funnel. Subsequently, the 3azido-7-hydroxycoumarin solution and 150 μL of a sodium ascorbate water solution (20 mg/mL) were added. The samples were purged with nitrogen for 25 min to remove the oxygen, and then, 60 μL of a CuSO4 solution (0.05 M) was added to the reaction mixture. The hydrogels were left to react for 30 min with nitrogen purging. Afterward, the hydrogels were washed with distilled water. Final visualization of the layers was performed by placing the entire volume of the gel onto the quartz cell for UV spectroscopy under a UV light

■

RESULTS AND DISCUSSION In the present work, the properties of materials obtained by a standard initiation method and by electron beam irradiation were compared. For this aim, a series of hydrogels based on acrylamide/N,N′-methylenebis(acrylamide) was prepared. Optionally, N-propargylacrylamide was used as a comonomer to introduce the alkyne group affording subsequent modifications C


Article

Biomacromolecules

uptake. It was demonstrated that the contribution of the polymer itself was small in comparison to the amount of water held in the pores of the samples. Moreover, the bounded water content was estimated by the water vapor adsorption method. It was shown that it was almost equal for cryogels obtained by Red-Ox and e-beam initiation (all the doses) and was around 0.8 mL/g. Hence, we can say that the type of the initiation as well as the irradiation doze influence the pores volume, but not the polymer network. Additionally, the cyclohexane uptake was measured. As cyclohexane does not swell the polymer network of the hydrogel, the inner volume of the permanent pores in the samples can be estimated with this solvent. The results obtained in cyclohexane demonstrated differences in the inner volumes of the samples: the inner volume was larger for the samples obtained by electron beam irradiation (Figure 2).

with various biomimetic ligands using azide−alkyne clickchemistry.48 All polymerization experiments were carried out at −15 °C. For the standard initiation method, the ammonium persulfate/ N,N,N′,N′-tetramethylethylenediamine system was used. The electron beam initiation was performed using a microtron MT 25 electron accelerator (see Supporting Information Figures S2 and S3 and Table S1 for a detailed description). Solvent uptake by the samples was measured in water and cyclohexane. The inner structure (morphology) of the samples was analyzed by SEM and light microscopy (LM), and the stiffness of the hydrogels was measured by compression testing. It was found that the yields of the macroporous polymer hydrogels had slightly depended on the irradiation doze, but not on the composition of samples (see Supporting Information Figure S1). Equilibrium Water Content and Solvent Uptake. The equilibrium water content analysis is crucial in the characterization of hydrogels. This parameter was expected to be relatively high for the cryogels due to the presence of an interconnected network of permanent pores. Although the obtained water uptake values for the two hydrogel types (redox and electron beam with 5 kGy) were different, it was found that the results were similar for both gel types and that the dependences on the cross-linker concentration followed the same trend. The increased crosslinker concentration contributed to the decrease in the water uptake. Figure 1 shows the dependence of water uptake on the crosslinker concentration for both methods of initiation. The

Figure 2. Dependence of solvent uptake in water and cyclohexane on the initiation method. The total monomer concentration was 7.5 wt %, the MBAA concentration was 2.4 mol %, the temperature was −15 °C, the concentration of the redox initiation system was 2 wt %, and the irradiation doses were 5 kGy, 10 kGy, and 15 kGy.

The hydrogels obtained by the two initiation methods exhibited different behavior in “good” and “bad” solvents (Figure 2). In the “good” solvent (water), the hydrogels formed with redox initiation showed higher water capacity than those obtained by irradiation. In the “bad” solvent (cyclohexane), the opposite trend was observed. This method allowed for the inner volume of the samples to be estimated, and the results indicated that the different initiation methods produced samples with different internal structures. The Light Microscopy Results. The main difference between the electron beam and standard redox polymerization is due to the conditions at which the polymerization reaction starts. In the redox initiating system, the initiator has to be added to the reaction mixture, while this is in a homogeneous solution and, hence, the polymerization starts before the sample is completely frozen. On the other hand, the electron beam can initiate the polymerization after sample freezing was complete. Thus, one of the important advantages of this approach is the ability to avoid the polymerization of any samples before they are completely frozen. This eliminates the problems with heterogeneity of samples that may result when the rapid progression of the polymerization (cross-linking) reaction occurs concurrently with the sample freezing process and

Figure 1. Dependence of the equilibrium water content on the crosslinker concentration for the hydrogels obtained by different initiation methods. The total monomer concentration was 7.5 wt %, the temperature was −15 °C, the concentration of the initiation system was 2 wt %, and the irradiation dose was 5 kGy.

equilibrium water content values decreased with increasing cross-linker concentration due to the reduced network flexibility. For both cases, the curves have a similar profile, but the hydrogels obtained with redox initiation had higher swelling values. It could be connected to the additional crosslinks appearance in the polymer network under the e-beam initiation conditions. The described method was not very precise but allowed for an estimation of the contributions of the solvent held in the pores and in the polymer itself to the total amount of water D


Article

Biomacromolecules

Figure 3. Light microscopy images of hydrogels obtained using redox initiation (A) and irradiation (B). The total monomer concentration was 7.5 wt %, the MBAA cross-linker concentration was 2.4 mol %, and the temperature was −15 °C. The concentration of the initiation system was 2 wt % compared to the monomers, and the irradiation dose was 5 kGy. The scale bar is 10 mm.

Figure 4. Scanning electron micrographs of hydrogels obtained using different irradiation doses. The total monomer concentration was 7.5 wt %, the cross-linker (MBAA) concentration was 2.4 mol %, and the temperature was −15 °C. The irradiation dose was A-5 kGy, B-10 kGy, and C-15 kGy. The scale bar is 500 μm.

which provide information regarding the pore volume, size distribution and surface area (see Table 1).

may produce different morphology of the gel as going from the sample edges to the center. Figure 3 presents light microscopy images showing differences in the morphology of cryogels prepared by the two different initiation methods. The samples obtained by electron beam irradiation (Figure 3B) display a more homogeneous inner structure. In contrast, the samples formed by the redox initiation (Figure 3A) exhibited different morphologies near the edge and in the middle, which were likely caused by a gradient in the sample freezing accompanied by progressive increase of viscosity caused by concurrent polymerization of the reaction mixture. Additionally, during the redox polymerization, nonporous areas would most likely appear as a result of the sharp increase in the rate of reaction in the nonfrozen phase. Hence, the ability to produce hydrogels with a more regular porous structure by the realization of distinct stages of freezing and initiation has been demonstrated. The Electron Scanning Microscopy Results. In addition, cryogels prepared using different irradiation doses were studied by SEM to examine the effects of this variable on the pore structure. Figure 4 demonstrates a clear dependence of the average pore size on the irradiation dose applied during synthesis. An increase in the irradiation dose from 5 to 15 kGy resulted in a decrease in the average pore size from 250 to 150 μm. This trend could be related to slight melting of the ice crystals during the high-dose irradiation and subsequent filling of this volume with the reaction mixture. Mercury Porosimetry and the BET Adsorption. In addition to characterizing the solvent uptake and investigating the microstructure by SEM, the porous structure was characterized by mercury porosimetry and BET adsorption,

Table 1. Characteristics of Hydrogels Obtained Using Different Irradiation Doses mercury porosimetry

dose 5 kGy 10 kGy 15 kGy

pore diameter (μm)

pore volume (mL/g)

porosity (%)

BET surface area (m2/g)

cyclohexane uptake

87 86

2.6 2.3

59.5 55.3

5.6 11.9

12.1 ± 0.3 9.1 ± 0.5

83

6.5

77.6

27.6

8.6 ± 0.9

Mercury porosimetry is usually used for systems with pores up to 116 μm in diameter,59 which represents a drawback for characterizing the macroporous systems obtained in this work. Nevertheless, the combination of this method with the cyclohexane uptake data can characterize the pores to some extent. Here, the solvent uptake study provided information regarding the large macropores, while the mercury porosimetry was employed for characterizing the small macro- and mesoporous structures.59 According to the mercury porosimetry data, as the irradiation dose increased, the average pore diameter decreased from 87 to 83 μm. A small volume of pores in the range of 0.05−116 μm (2.63 and 2.27 mL/g, respectively) was found for samples obtained using 5 and 10 kGy irradiation. However, the values obtained for the solvent uptake were approximately 12 and 9 mL/g, respectively. For the samples obtained under the highest E


Article

Biomacromolecules

Figure 5. Cumulative pore volume (-■-) and the derived pore size distribution (bars) of samples prepared with 5 kGy (A), 10 kGy (B), and 15 kGy (C), obtained by mercury porosimetry.

the alkyne groups may become unavailable for modification due to steric hindrance or other difficulties related to the structure of the sample. These obstacles were particularly expected for samples obtained with a higher irradiation dose. Hence, an experiment involving surface modification with a radiolabeled RGD peptide was performed to estimate the quantity of alkyne groups available for the reaction. Despite the strong possibility of damage because of the high irradiation dose applied, the results revealed that most of the alkyne groups were available for modification. The difference between the results for the samples with and without the copper catalyst indicates that the radiolabeled peptides were specifically coupled with alkyne groups on the surface of the hydrogel. The average radioactivity value was 0.710 ± 0.040 MBq for samples with the copper catalyst and 0.047 ± 0.003 MBq for samples without the catalyst. This result corresponds to 31.0 ± 1.7 pmole of peptide/cm2 for the specific modification, while nonspecific adsorption only accounted for approximately 6.6% of the incorporated peptide (eq 3). This amount of peptide exceeds the requirement for cell adhesion and proliferation.

irradiation dose, the values of both experiments were closer than for the other samples. This result indicates the presence of structured pores that are larger than the mercury porosimetry detection limit. Thus, we cannot determine the exact size of the pores in the samples, but we can conclude that a correlation between the irradiation dose and pore size exists. Furthermore, by comparing these two methods, it is possible to conclude that the samples obtained with 5 and 10 kGy primarily had macropores with diameters greater than 116 μm and that the pores became smaller as the irradiation dose increased (Table 1 and Figure 5). Data for the different irradiation doses are shown in Table 1. As the pores become smaller, their volume decreases, which can be seen from the values for cyclohexane uptake (Table 1). At the same time, according to the BET adsorption test, an increased irradiation dose resulted in an increased inner surface area, which is directly related to the increased numbers of pores coupled with their shrinking. These results are in good agreement with the SEM findings (Figure 4). Biomimetic Modification. The data for the inner surface area are crucial for the biomimetic modification of the hydrogels. To promote cell adhesion and proliferation, the peptide concentration on the matrix surface should be in the range of 1 to 1000 fmole/cm2.60 The initial concentration of alkyne groups was sufficiently high to meet this modification requirement, but some of the groups may be affected by the irradiation during the gel formation process and may lose their functionality. In addition, after the formation process, some of

C=

A SA ·M ·S

(3)

where C is the concentration of the peptide on the hydrogel surface (mol/cm2), A is the radioactivity of the sample after the reaction (MBq), SA is the specific activity of the peptide (MBq/ mg), M is the molecular weight of the peptide (g/mol; M = F


Article

Biomacromolecules 1064 g/mol), and S is the surface area (m2/g), estimated by BET. Elasticity. As the irradiation dose was increased to 15 kGy, the mechanical characteristics of the polymer network, such as the modulus of elasticity, changed. Figure 6A illustrates the

This hypothesis was also supported by the water uptake test (Figure 6B). Because the internal structure changed as the irradiation dose was increased, the values of equilibrium water content were also modified. This dependence was more obvious for samples with a low cross-linker concentration. The transition from the system with a lower cross-linker content to systems with a higher content led to a moderate change in the swelling rate. Apparently, a limit exists at which the cross-linking density is sufficient to achieve a different combination of cross-links formed by the cross-linker and additional cross-links caused by the higher irradiation dose, thus producing different outcomes in the water uptake. The irradiation dose had an influence on the internal structure of the hydrogels, as well as their mechanical properties and water uptake amount. An increased dose led to the formation of smaller pores, higher stiffness, and lower water uptake by the hydrogels. Thus, it is possible to tailor the average pore size, the mechanical properties, and the swelling properties of the hydrogels by changing the irradiation dose, which could be very useful for tissue and cell growth. According to Reilly et al.,61 hydrogels with various stiffness values can be used as growth supports for different tissue types. For example, the modulus of elasticity of a hydrogel used for the cultivation of skeletal muscle and smooth muscle should be ∼8−17 kPa and ∼5−9 kPa, respectively. Furthermore, by tuning the matrix stiffness, specific stem cell behavior can be achieved.62 Thus, the different types of obtained hydrogels can be utilized to facilitate the growth of specific cell types. Cell Viability, Adhesion, and Spreading in Hydrogels. The technique of preparation the acrylamide hydrogels is interesting mainly because of possibility to adjust hydrogel elasticity and porosity along with modification of material surface. These factors are important for precise manipulation and control of cell behavior, which is crucial for engineering scaffolds and its future utilization. The perspectives of these hydrogels can be evaluated by using human cells encapsulated in materials and determining the interaction of cells with material surface. To do so, we have

Figure 6. (A) Dependence of the compressive modulus on the irradiation dose. The total monomer concentration was 7.5 wt %, the cross-linker (MBAA) concentration was 2.4 mol %, and the temperature was −15 °C. The modulus values were 5 kGy = 2.76 ± 0.06 kPa, 10 kGy = 5.63 ± 0.12 kPa, and 15 kGy = 11.21 ± 0.05 kPa. (B) Dependence of the water uptake values on the irradiation dose for hydrogels with different compositions. The total monomer concentration was 7.5 wt %, the temperature was −15 °C, and the samples were incubated for 7 days to reach equilibrium.

correlation between the modulus of elasticity and the irradiation dose. Irradiation with a higher dose during hydrogel formation could contribute to a higher stiffness of the samples by increasing the cross-linking density. The increased stiffness of the samples could then decrease their ability to stretch after water uptake. Hence, the water uptake would also be reduced.

Figure 7. Live/dead assay of ADSCs seeded on irradiated hydrogels with or without the surface modification by RGD peptide. Elastic modulus is related to irradiation dose. Viable cells were labeled with acridine orange (green), dead cells were stained by ethidium bromide (red) and observed rarely. Hydrogels were made with the total monomer concentration 7.5 wt %, the cross-linker (MBAA) concentration 2.4 mol %, and the temperature −15 °C. Scale bar is 500 μm. G


Article

Biomacromolecules

Figure 8. Actin cytoskeleton (red) and nuclei (blue) staining of adipose-derived stem cells observed with confocal microscopy. Cells adhered only to RGD modified hydrogels, and they were most spread on the material with the lowest elastic modulus (3 kPa), whereas stiffer materials allowed cell adhesion but limited cell spreading. ADSCs encapsulated to unmodified hydrogels formed cell clusters without attachment to material. Scale bar 100 μm.

Figure 9. Scanning electron microscopy of adipose-derived stem cells adhered and spread on hydrogels w/o surface RGD modification and different stiffness. Scale bar 20 μm.

These values correspond to the distance of RGD ligands equal to 1.1, 1.6, and 2.5 nm, which is less than one-third the diameter of the integrin head of 7 nm.63 Several peptides become then accessible for one integrin unit under this condition, therefore the capacity of binding sites was considered to be sufficient for cell adhesion and spreading.60 From fluorescence microscopy of actin cytoskeleton, we observed that ADSCs formed strong stress fibers mainly on soft hydrogels (3 kPa) and cells were spread inside as well as across material pores. Although cells adhered also to surface of stiffer gels, the actin fibers were thinner, and cells were much less spread (Figure 8). These morphological observations were also confirmed by SEM (Figure 9). Cells seeded to unmodified hydrogels were round-shaped and formed cell clusters without notable interaction between cells and material. The stiffer materials (6, 11 kPa) with surface modification supported cell attach-

chosen cells isolated from human adipose tissue, because they are representing noncancerous human stem cells, and moreover they are accessible for applications in clinical practice. Adipose-derived stem cells were homogeneously seeded within irradiation polymerized hydrogels with and without surface modification. Cells were examined by fluorescent live/ dead assay to determine whether cells survive in hydrogels during 24 h cultivation. Viable ADSCs (green) were observed throughout all hydrogel scaffolds, and dead cells (red) were observed only occasionally. No differences in cell viability were noted among different hydrogel scaffolds (Figure 7). The presence of integrin-binding motif RGD was necessary for adipose-derived stem cell attachment. Concentrations of incorporated peptide were determined from measurements of peptide radioactivity and BET surface area of hydrogels, and they were calculated as 156, 74, and 31 pmoles of peptide/cm2, related to irradiation doses 5, 10, and 15 kGy, respectively. H


Article

Biomacromolecules

cell for UV spectroscopy (λ = 366 nm) (Figures 4 and S13). For comparison, the same experiment was performed with a nonlayered cryogel before and after the click-reaction. Figure 10 shows the contrast in the emission among the layers of varying composition. This result confirms the layered structure of the studied gels (see Figure S4 for the color image).

ment, but cells were usually round-shaped, or they stretched through pores or formed cell clusters. RGD modified hydrogels with lowest elasticity allowed cells to fully spread on its surface and connect with other cells. It is important to note that the concentration of RGD peptides on the surface material was excessive for cell attachment and comparable among hydrogels regardless of irradiation dose. Consequently, the presumptive factor for influencing cell spreading is stiffness and porosity of hydrogels. Curiously, described cellular response is in contrast with published analysis of cell behavior on planar substrates, where cells presented round-shaped morphology on soft surfaces and extend spreading with increasing stiffness of material (for example, see refs 64−67). It is necessary to put such observations into the proper context: native extracellular matrix is fibrillar and has a specific spatial organization and distribution of RGD-containing binding sites. Most studies to date have employed flat surfaces, which could dramatically alter how a cell generates traction forces and senses its environment.61 Thus, we assume that in our work these opposite effects on cell spreading could be related to 3D structure and porosity of examined hydrogels. These results, though, confirm the hydrogel biocompatibility and potential for precise stimulation of cellular behavior and design scaffolds for tissue engineering. Multilayered Cryogels. Another aim of this work was to provide a proof-of-principle for producing layered macroporous hydrogels by electron beam irradiation. The model samples contained layers of reaction mixture with and without propargylacrylamide. The presence of alkyne groups located in separate layers of a layered structure in these materials was confirmed by a click-reaction with an azidocoumarin derivative. When considering multilayered hydrogels, it is important to remember that significant interlayer mixing must be avoided; however, limited interpenetration of the layers is necessary to obtain a bulk material with interconnected layers. If interlayer cross-linking cannot be achieved, delamination of the layers would be observed after polymerization. The possibility of preparing samples with layers of different composition by redox-initiated polymerization is very limited, because the initiation starts while the system is still in a liquid phase. Using the electron beam initiation approach, the layered structure was produced by sequentially adding the solution up to the previous layer while incubating the entire vial at a subzero temperature until the reaction vial was filled. Thus, the samples contained layers with and without propargylacrylamide, which was added at 5 mol % with respect to the acrylamide component (3.2 mol % with respect to all of the solids). Importantly, the given procedure allows for partial thawing of the frozen phase at the edge of the layer, and mixing of the layers occurs as a result of the addition to the already frozen reaction mixture solution at a higher temperature. The frozen layered samples were finally irradiated under the conditions described above for nonlayered gels. After thawing and washing, compact bulk gels were obtained. The next step was to visualize the obtained layered structure. For this purpose, 3-azido-7-hydroxycoumarin was reacted with the alkyne groups contained in some of the sample layers. The advantage of using an azidocoumarin derivative was its ability to fluoresce only after the click-reaction. The initial coumarin derivative does not fluoresce under UV light.55 Finally, the layers were visualized by placing the entire volume of the gel under a lamp for TLC and by using a quartz

Figure 10. Hydrogels visualized under UV light. (A) Thin slice of a nonlayered hydrogel before reacting with 3-azido-7-hydroxycoumarin. (B) Thin slice of a nonlayered hydrogel after reacting with 3-azido-7hydroxycoumarin. (C) A bulk sample of layered hydrogel after reacting with 3-azido-7-hydroxycoumarin. The monomer concentration was 7.5 wt %, the temperature was −15 °C, and the irradiation dose was 5 kGy.

■

CONCLUSIONS The electron beam and redox initiation methods are suitable for obtaining porous polymer hydrogels under subzero conditions, such as cryogels. The differences in the characteristics of hydrogels obtained by the two distinct methods were caused by variations in their morphology. However, the electron beam irradiation proved to be advantageous for producing hydrogels with a regular internal structure and tunable physical properties, where decreased irradiation dose contributed to changes in the pore size, modulus of elasticity, and water uptake. The adhesion and viability of seeded cells on the surfaces with different RGD content prove nontoxicity of cryogels and the preference of ADSC cells to the matrix with the lowest elasticity among the revised samples. Further investigation of cryogels with adjustable substrate elasticity and biomimetic ligand composition/concentration opens the way to design scaffolds for various tissue engineering applications. Moreover, a method for preparing multilayered cryogels by electron-beam-initiated polymerization was demonstrated. This finding opens possibilities for the future preparation of macroporous cryogels with domains exhibiting different mechanical properties and biomimetic groups for more complex tissue engineering scaffold design and fabrication.

■

ASSOCIATED CONTENT

S Supporting Information *

Supporting Information including the formation of hydrogels using irradiation, electron beam irradiation of the cryogel, and hydrogels visualized under UV light is available free of charge via the Internet at http://pubs.acs.org.

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. I


Article

Biomacromolecules Notes

(25) Sun, J.; Wei, D.; Zhu, Y.; Zhong, M.; Zuo, Y.; Fan, H.; Zhang, X. Biomaterials 2014, 35, 4759−4768. (26) Studenovska, H.; Slouf, M.; Rypacek, F. J. Mater. Sci.: Mater. Med. 2008, 19, 615−621. (27) Kubinova, S.; Horak, D.; Hejcl, A.; Plichta, Z.; Kotek, J.; Proks, V.; Forostyak, S.; Sykova, E. J. Tissue Eng. Regen. Med. 2013, n/a. (28) He, X.; Yao, K.; Shen, S.; Yun, J. Chem. Eng. Sci. 2007, 62, 1334−1342. (29) Lozinsky, V. I.; Galaev, I. Y.; Plieva, F. M.; Savinal, I. N.; Jungvid, H.; Mattiasson, B. Trends Biotechnol. 2003, 21, 445−451. (30) Chen, G.; Gulbranson, D.; Hou, Z.; Bolin, J.; Ruotti, V.; Probasco, M.; Smuga-Otto, K.; Howden, S.; Diol, N.; Propson, N. Nat. Methods 2011, 8, 424−429. (31) Hsieh, C.-Y.; Tsai, S.-P.; Ho, M.-H.; Wang, D.-M.; Liu, C.-E.; Hsieh, C.-H.; Tseng, H.-C.; Hsieh, H.-J. Carbohydr. Polym. 2007, 67, 124−132. (32) Noppe, W.; Plieva, F. M.; Vanhoorelbeke, K.; Deckmyn, H.; Tuncel, M.; Tuncel, A.; Galaev, I. Y.; Mattiasson, B. J. Biotechnol. 2007, 131, 293−299. (33) Damshkaln, L. G.; Simenel, I. A.; Lozinsky, V. I. J. Appl. Polym. Sci. 1999, 74, 1978−1986. (34) Gun’ko, V. M.; Savina, I. N.; Mikhalovsky, S. V. Adv. Colloid Interface Sci. 2013, 187, 1−46. (35) Fromageau, J.; Gennisson, J. L.; Schmitt, C.; Maurice, R. L.; Mongrain, R.; Cloutier, G. IEEE Trans. Ultrason. Ferroelectr. Freq. Control 2007, 54, 498−509. (36) Topuz, F.; Okay, O. React. Funct. Polym. 2009, 69, 273−280. (37) Hoffman, A. S. Adv. Drug Delivery Rev. 2012, 64, 18−23. (38) Saha, K.; Keung, A. J.; Irwin, E. F.; Li, Y.; Little, L.; Schaffer, D. V.; Healy, K. E. Biophys. J. 2008, 95, 4426−4438. (39) Plieva, F. M.; Galaev, I. Y.; Mattiasson, B. J. Sep. Sci. 2007, 30, 1657−1671. (40) Kuyukina, M. S.; Rubtsova, E. V.; Ivshina, I. B.; Ivanov, R. V.; Lozinsky, V. I. J. Microbiol. Methods 2009, 79, 76−81. (41) Dragan, E. S.; Loghin, D. F. A. Chem. Eng. J. 2013, 234, 211− 222. (42) Reichelt, S.; Becher, J.; Weisser, J.; Prager, A.; Decker, U.; Moeller, S.; Berg, A.; Schnabelrauch, M. Mater. Sci. Eng. C: Mater. Biol. Appl. 2014, 35, 164−170. (43) Reichelt, S.; Prager, A.; Abe, C.; Knolle, W. Radiat. Phys. Chem. 2014, 94, 40−44. (44) Hennink, W. E.; van Nostrum, C. F. Adv. Drug Delivery Rev. 2002, 54, 13−36. (45) Zhao, L.; Gwon, H.-J.; Lim, Y.-M.; Nho, Y.-C.; Kim, S. Y. Carbohydr. Polym. 2014, 102, 598−605. (46) Reichelt, S.; Abe, C.; Hainich, S.; Knolle, W.; Decker, U.; Prager, A.; Konieczny, R. Soft Matter 2013, 9, 2484−2492. (47) Reichelt, S.; Prager, A.; Abe, C.; Knolle, W. Radiat. Phys. Chem. 2014, 9, 40−44. (48) Rostovtsev, V. V.; Green, L. G.; Fokin, V. V.; Sharpless, K. B. Angew. Chem., Int. Ed. 2002, 41, 2596−2599. (49) Frantz, C.; Stewart, K. M.; Weaver, V. M. J. Cell Sci. 2010, 123, 4195−4200. (50) Lu, T.-Y.; Lin, B.; Kim, J.; Sullivan, M.; Tobita, K.; Salama, G.; Yang, L. Nat. Commun. 2013, 4, 2307. (51) Nguyen, L. H.; Kudva, A. K.; Saxena, N. S.; Roy, K. Biomaterials 2011, 32, 6946−6952. (52) Lee, Y.-H.; Chang, J.-J.; Lai, W.-F.; Yang, M.-C.; Chien, C.-T. Colloids Surf., B 2011, 86, 409−413. (53) Lin, T. Y.; Ki, C. S.; Lin, C. C. Biomaterials 2014, 35, 6898− 6906. (54) Macková, H.; Proks, V.; Horák, D.; Kučka, J.; Trchová, M. J. Polym. Sci., Polym. Chem. 2011, 49, 4820−4829. (55) Sivakumar, K.; Xie, F.; Cash, B. M.; Long, S.; Barnhill, H. N.; Wang, Q. Org. Lett. 2004, 6, 4603−4606. (56) Plieva, F. M.; Karlsson, M.; Aguilar, M.-R.; Gomez, D.; Mikhalovsky, S.; Galaev, I. Yu. Soft Matter 2005, 1, 303−309. (57) Rigby, S. P.; Barwick, D.; Fletcher, R. S.; Riley, S. N. Appl. Catal., A 2003, 238, 303−318.

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS The financial support of the Czech Science Foundation (No. 14-14961S, P108/11/1857 and 13-29009S), projects “BIOCEV− Biotechnology and Biomedicine Centre of the Academy of Sciences and Charles University” (CZ.1.05/1.1.00/02.0109) from the European Regional Development Fund, and “HistoPARK − Centre for analysis and modeling of tissues and organs” (CZ.1.07/2.3.00/20.0185) from the European Social Fund in the Czech Republic is gratefully acknowledged. The authors also acknowledge the Charles University in Prague, Department of Physical and Macromolecular Chemistry, and University of Pardubice, Institute of Chemistry and Technology of Macromolecular Materials, Faculty of Chemical Technology, for the opportunity for doctoral studies.

■

REFERENCES

(1) Peppas, N. A., Ed. Fundamentals; CRC Press, Inc.: Boca Raton, FL, 1986; Vol. 1. (2) Wichterle, O.; Lim, D. Nature 1960, 185, 117−118. (3) Xinming, L.; Yingde, C.; Lloyd, A. W.; Mikhalovsky, S. V.; Sandeman, S. R.; Howel, C. A.; Liewen, L. Contact Lens Anterior Eye 2008, 31, 57−64. (4) Campbell, S. B.; Hoare, T. Curr. Opin. Chem. Eng. 2014, 4, 1−10. (5) Bertz, A.; Woehl-Bruhn, S.; Miethe, S.; Tiersch, B.; Koetz, J.; Hust, M.; Bunjes, H.; Menzel, H. J. Biotechnol. 2013, 163, 243−249. (6) Mazzitelli, S.; Pagano, C.; Giusepponi, D.; Nastruzzi, C.; Perioli, L. Int. J. Pharm. 2013, 454, 47−57. (7) Li, H.; Yang, J.; Hu, X.; Liang, J.; Fan, Y.; Zhang, X. J. Biomed. Mater. Res., Part A 2011, 98A, 31−39. (8) Heilmann, S.; Kuechler, S.; Wischke, C.; Lendlein, A.; Stein, C.; Schaefer-Korting, M. Int. J. Pharm. 2013, 444, 96−102. (9) Gong, C.; Wu, Q.; Wang, Y.; Zhang, D.; Luo, F.; Zhao, X.; Wei, Y.; Qian, Z. Biomaterials 2013, 34, 6377−6387. (10) Huang, X.; Zhang, Y.; Zhang, X.; Xu, L.; Chen, X.; Wei, S. Mater. Sci. Eng. C:Mater. Biol. Appl. 2013, 33, 4816−4824. (11) Cavalcanti, B. N.; Zeitlin, B. D.; Noer, J. E. Dent. Mater. 2013, 29, 97−102. (12) Matricardi, P.; Di Meo, C.; Coviello, T.; Hennink, W. E.; Alhaique, F. Adv. Drug Delivery Rev. 2013, 65, 1172−1187. (13) Radhakrishnan, J.; Krishnan, U. M.; Sethuraman, S. Biotechnol. Adv. 2014, 32, 449−461. (14) Saha, K.; Keung, A. J.; Irwin, E. F.; Li, Y.; Little, L.; Schaffer, D. V.; Healy, K. E. Biophys. J. 2008, 95, 4426−4438. (15) Adeloew, C.; Segura, T.; Hubbell, J. A.; Frey, P. Biomaterials 2008, 29, 314−326. (16) Discher, D. E.; Janmey, P.; Wang, Y. L. Science 2005, 310, 1139−1143. (17) Spencer, H.; Keramari, M.; Ward, C. Methods Mol. Biol. 2011, 690, 81−94. (18) Kubinová, S.; Horák, D.; Kozubenko, N.; Vanecek, V.; Proks, V.; Price, J.; Cocks, G.; Syková, E. Biomaterials 2010, 31, 5966−5975. (19) Yu, T. T.; Shoichet, M. S. Biomaterials 2005, 26, 1507−1514. (20) Hacker, M.; Tessmar, J.; Neubauer, M.; Blaimer, A.; Blunk, T.; Gopferich, A.; Schulz, M. B. Biomaterials 2003, 24, 4459−4473. (21) Rahmany, M. B.; Van Dyke, M. Acta Biomater. 2013, 9, 5431− 5437. (22) Guvendiren, M.; Burdick, J. A. Curr. Opin. Biotechnol. 2013, 24, 841−846. (23) Mason, B.; Califano, J.; Reinhart-King, C. In Engineering Biomaterials for Regenerative Medicine; Bhatia, S. K., Ed.; Springer: New York, 2012; pp 19−37. (24) Michalek, J.; Pradny, M.; Artyukhov, A.; Slouf, M.; Smetana, K. J. Mater. Sci.: Mater. Med. 2005, 16, 783−786. J


Article

Biomacromolecules (58) Zuk, P. A.; Zhu, M.; Ashjian, P.; et al. Mol. Biol. Cell 2002, 13, 4279−4295. (59) Horák, D.; Hlídková, H.; Hradil, J.; Lapčíková, M.; Šlouf, M. Polymer 2008, 49, 2046−2054. (60) Proks, V.; Jaros, J.; Pop-Georgievski, O.; Kucka, J.; Popelka, S.; Dvorak, P.; Hampl, A.; Rypacek, F. Macromol. Biosci. 2012, 12, 1232. (61) Reilly, G. C.; Engler, A. J. J. Biomech. 2010, 43, 44−62. (62) Dingal, P. C. D. P.; Discher, D. E. Curr. Opin. Biotechnol. 2014, 28, 46−50. (63) Missirlis, D.; Spatz, J. P. Biomacromolecules 2013, 15, 195. (64) Young, D. A.; Choi, Y. S.; Engler, A. J.; Christman, K. L. Biomaterials 2013, 34, 8581−8588. (65) Trappmann, B.; Gautrot, J. E.; Connelly, J. T.; Strange, D. G. T.; Li, Y.; Oyen, M. L.; Cohen Stuart, M. A.; Boehm, H.; Li, B.; Vogel, V.; Spatz, J. P.; Watt, F. M.; Huck, W. T. S. Nat. Mater. 2012, 11, 642− 649. (66) Ananthanarayanan, B.; Kim, Y.; Kumar, S. Biomaterials 2011, 32, 7913−7923. (67) Engler, A. J.; Sen, S.; Sweeney, H. L.; Discher, D. E. Cell 2006, 126, 677−689.

K


Toward Structured Macroporous Hydrogel Composites: Electron Beam

Recommend Documents