In Situ Fabrication of Fiber Reinforced Three ... - ACS Publications

Jun 7, 2017 - PEO hydrogels with a stiffness of 8.71 ± 0.21 MPa. Fibroblast cells .... and S2.51 Three classes of coextruded composites were fabricat...
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Article pubs.acs.org/journal/abseba

In Situ Fabrication of Fiber Reinforced Three-Dimensional Hydrogel Tissue Engineering Scaffolds Alex M. Jordan, Si-Eun Kim, Kristen Van de Voorde, Jonathan K. Pokorski, and LaShanda T. J. Korley* Center for Layered Polymeric Systems, Department of Macromolecular Science and Engineering, Case Western Reserve University, Cleveland, Ohio 44106-7202, United States S Supporting Information *

ABSTRACT: Hydrogels are an important class of biomaterials, but are inherently weak; to overcome this challenge, we report an in situ manufacturing technique to fabricate mechanically robust, fiber-reinforced poly(ethylene oxide) (PEO) hydrogels. Here, a covalent PEO cross-linking scheme was implemented to derive poly(ε-caprolactone) (PCL) fiber reinforced PEO hydrogels from multilayer coextruded PEO/PCL matrix/fiber composites. By varying PCL fiber loading between ∼0.1 vol % and ∼7.8 vol %, hydrogel stiffness was tailored from 0.69 ± 0.04 MPa to 1.94 ± 0.21 MPa. The influence of PCL chain orientation and enhanced mechanics via uniaxial drawing of PCL/PEO composites revealed a further 225% increase in hydrogel stiffness. To further highlight the robust nature of this manufacturing process, we also derived rigid poly(L-lactic acid) (PLLA) fiber-reinforced PEO hydrogels with a stiffness of 8.71 ± 0.21 MPa. Fibroblast cells were injected into the hydrogel volume, which displayed excellent ingrowth, adhesion, and proliferation throughout the fiber reinforced hydrogels. Finally, the range of mechanical properties obtained with fiber-reinforced hydrogels directed differentiation pathways of MC3T3-E1 cells into osteoblasts. This innovative manufacturing approach to achieve randomly aligned, well-distributed, micrometer-scale fibers within a hydrogel matrix with tunable mechanical properties represents a significant avenue of pursuit not only for load-bearing hydrogel applications, but also targeted cellular differentiation. KEYWORDS: fibers, hydrogels, tissue engineering, cell scaffolds

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INTRODUCTION Hydrogels, both natural and synthetic, have received significant interest in recent years as tissue engineering scaffolds. The function of these scaffolds is to mimic the extracellular matrix (ECM) and provide a platform for cell adhesion, proliferation, and differentiation by controlling the structure and function of the hydrogel scaffold.1 Natural materials, including proteins and polysaccharides, have been used extensively to form hydrogels for tissue engineering applications because of their inherent biocompatibility, lack of immunogenicity, and flexibility in network formation via the diversity of their chemical structures. However, these natural materials are challenged by their weak mechanics and limited long-term stability due to dissolution behavior.2−7 To overcome the rapid dissolution of natural materials in vivo, researchers have turned to synthetic polymers, such as poly(vinyl alcohol) (PVA), poly(acrylic acid) (PAA), poly(hydroxyethyl methacrylate) (PHEMA), poly(N-isopropylacrylamide) (PNIPAAm), poly(ethylene glycol) (PEG), and its higher-molecular-weight analogue poly(ethylene oxide) (PEO) as hydrogel building blocks.8,9 Specific cross-linking chemistries, including thiol−ene photopolymerization10 and multiarm PEG cross-linked by Michael-type addition,11 are more commonly implemented because of their selective, highyield pathways to develop 3D PEG hydrogels for drug delivery © XXXX American Chemical Society

and tissue engineering. However, PEG/PEO hydrogels have also been fabricated via chain scission and recombination using high energy electron-beam or Co60 γ-radiation,12,13 physical associations in supramolecular networks,14 peroxide mediated coupling,15 and ultraviolet (UV) radiation induced radical coupling,16 and have found many uses as synthetic tissue scaffolds. While attractive for biomedical applications ex vivo, such as cell culture in a laboratory setting, these PEO and other synthetic hydrogels, which consist primarily of water, are inherently weak, limiting their utility in vivo.17 Recently, Zhao reviewed potential strategies, including network architectures, incorporation of both strong covalent and weak ionic cross-links, and the inclusion of fiber elements, for improving the mechanical behavior of synthetic hydrogels under physiological conditions.17 The formation of dualnetwork architectures have been shown to enhance the extensibility and strength of synthetic hydrogels, at the expense of diffusivity through the hydrogels.18−21 Incorporation of strong and weak cross-links has also increased hydrogel toughness via an enhancement in extensibility.22−30 Sun et al. Received: April 12, 2017 Accepted: May 23, 2017

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rigid to be deformed by the gel, and as a result the gel matrix simply deformed around the embedded fibers. However, the size scale of fibers manufactured utilizing this strategy is limited to the resolution of 3D printing, currently ∼25 μm feature size−well above the nanoscale dimensions of collagen. It is proposed that there are three critical design considerations when fabricating fiber reinforced hydrogels: (1) fiber distribution throughout the hydrogel volume, (2) random fiber alignment, and (3) fiber diameter less than 40 μm. Recent innovations in multilayer coextrusion43−49 have utilized this scalable, melt-processing strategy to manufacture polymer composites containing distributed domains of microand nanoscale fibers.50,51 Extruded composites of rectangular PCL fibers in a PEO matrix have been investigated for relevance in biomedical applications. Uniaxial PCL/PEO composite drawing and subsequent removal of the PEO matrix yielded PCL fiber scaffolds with tunable mechanics, high surface area, and nanoscale dimensions.51,52 Functional coextruded PCL fibers were also achieved by Kim et al. via surface modification using “click” chemistry, imparting gradient density modification and multifunctionality to enhance cellular proliferation and direct differentiation along the fiber scaffolds.53−56 Although these extruded PCL fiber mats have potential in a range of biomedical applications, one can envision that this manufacturing platform may address key challenges in the development of fiber-reinforced hydrogels with synergistic mechanics due to the structural organization of these fiber/ matrix coextruded composites with a distributed microscale PCL fiber architecture throughout the “sacrificial” PEO matrix. Here, we implement a facile, UV-initiated covalent cross-linking scheme to fabricate polyester fiber-reinforced PEO hydrogels as cell scaffolds with tunable mechanics as a function of fiber loading and fiber type.

demonstrated a complex toughening mechanism in hybrid hydrogels of ionically cross-linked alginate and covalently crosslinked polyacrylamide; the dissociation of ionic cross-links absorbed internal damage, whereas the covalent cross-links formed a network for crack bridging, facilitating large scale (∼20× original size) deformation of hybrid hydrogels.30 Fiber reinforcement of synthetic hydrogels has received significant interest in recent years with toughness enhancement reported to occur via fiber fracture/pull-out mechanisms associated with crack propagation and energy dissipation.31−38 This recent attention has coincided with the development of new fiber manufacturing techniques, such as three-dimensional (3D) printing and the resurgence of electrospinning and its many adaptations, to fabricate nano- to microscale polymeric fibers with architectures similar to that of collagen fibrils and bundles found in natural systems. However, there are a number of architectural obstacles that must be overcome in the additive manufacturing process to impregnate these fibrous constructs with gel matrix to fabricate fiber reinforced hydrogels.36 To truly function as a synergistic reinforcing network, individual fibers must not only be on the nano- and microscale, but must also be distributed throughout the volume of the hydrogel matrix. The use of solution electrospun fibrous constructs as a reinforcing material for hydrogels has been recently reviewed by Bosworth and colleagues.39 Utilizing solvent-based electrospun fibers incorporated into hydrogels as a layered structure, the water uptake was significantly reduced in this configuration because of the restricted elasticity imposed by the fiber network. To date, a majority of the work involving embedded electrospun mats within hydrogel constructs has focused on the impact of fiber-directed cell proliferation.40,41 Even though solution electrospinning provides a platform for fabrication of nanoscale fibers, there exists a lack of architectural and spatial control over the fiber distribution within the hydrogel matrix. For example, the reinforcing effect of electrospun poly(ε-caprolactone) (PCL) fibers embedded in an alginate hydrogel was examined, highlighting an enhancement in tensile strength and strain-to-break compared to the neat alginate hydrogels; however, a synergistic effect was not achieved in these composite hydrogels with mechanical response similar to the neat electrospun PCL fibers.42 To achieve a randomly dispersed fiber architecture, Vlassak et al. formed an alginate and double network alginate-polyacrylamide hydrogel around a mass of steel wool.38 Although this approach has its obvious limitations in biomedical applications, it demonstrated a synergistic enhancement in tensile strength, modulus, and strain-to-break of the reinforced hydrogel. To overcome the architectural challenges presented via the use of electrospun fibers as the reinforcing component of bioscaffolds, recent studies have utilized the advent of 3D printing technology to fabricate reinforcing scaffolds.37 Recently, a controlled 0−90° architecture of PCL fibers down to ∼20 μm was obtained via direct write melt electrospinning as a reinforcing element for a cross-linked gel-methacrylate hydrogel.36 A synergistic effect was revealed, suggesting a cooperative mechanism between the small (20−40 μm) fibers and the surrounding hydrogel matrix, which improved the observed stiffness by almost 2 orders of magnitude compared to the use of larger fibers. This cooperative mechanism was explained by the fact that small fibers (40 μm diameter) were too

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EXPERIMENTAL SECTION

Coextruded Composite Fabrication. Coextruded polyester fiber/PEO matrix (Dow POLYOX WSR-N:80/WSR-N:10, 70/30 blend) composites were fabricated using a previously reported technique (Figure 1, Step 1); full details are provided in Figures S1 and S2.51 Three classes of coextruded composites were fabricated with a PEO matrix (Table 1). Coextruded Composite Dissolution. Coextruded composites were cut into 5 mm strips and dissolved in distilled water (H2O), separating the soluble PEO from the insoluble fibrous phase (PCL or poly(L-lactic acid) (PLLA)). Composites were first agitated overnight at 800 rpm before ultrasonication at 50 °C for 90 min to separate and disperse aggregated fiber bundles throughout the H2O/PEO mixture (Figure 1, Step 2). Once individual fibers were randomly aligned and distributed throughout the mixture, a PEO cross-linking agent, pentaerythritol triacrylate (PETA, Sigma-Aldrich Technical grade, CAS: 3524-68-3), was added in a 60:40 (PETA:PEO) weight ratio. The fiber-containing H2O/PEO/PETA mixture was agitated again to ensure adequate mixing between PEO and PETA (Figure 1, Step 3). PEO Cross-Linking. After mixing, the fiber-containing solution was poured into a Teflon mold and the H2O allowed to evaporate, leaving a solid film (Figure 1, Step 4). Each film was exposed to ultraviolet (UV, Honle Blue Print 4 Ecocure) radiation with radiation wavelength 320−390 nm for 30 min at a radiation intensity of 33 ± 2 mW/cm2 to facilitate cross-linking between the acrylate moieties of PETA and the PEO backbone (Figure 1, Step 5). A range of PETA:PEO concentrations and UV exposure times were investigated. The high PEO molecular weight required for melt coextrusion and the absence of reactive end groups necessitated a higher weight ratio of PETA:PEO to achieve an appropriate gel fraction within l hr. Following UV exposure, the xerogel (e.g., dry cross-linked composite) film was swollen in distilled H2O again, forming a fiber-reinforced PEO B

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ACS Biomaterials Science & Engineering A1 =

Table 1. Fabricated Coextruded Fiber-Reinforced PEO Composites

poly(ε-caprolactone), PCL (CAPA 6800)

PCL + 3 wt % Sunset Yellow FCF (Sigma-Aldrich, CAS: 2783-94-0) poly(L-lactic acid), PLLA (NatureWorks 2003D)

uniaxial drawing

role

2.6 6.0 10.9 15.3 22.4 30.5 30.5

DR = 1

examine influence of fiber loading on hydrogel mechanics

DR = 6

30.5

DR = 1

probe effect of fiber chain orientation visualize fiber distribution

30.5

DR = 1

)

−2

(1)

m3 − A1m1 m1(1 − A1)

(2)

SR =

m 2 − m3 m3 − A1m1

(3) A1 ρf

A1 ρf

+

0.6(1 − A1) ρPEO

+

0.4(1 − A1) ρPETA

+

SR XD(1 − A1) ρH O 2

(4)

Fiber Distribution. Hydrogels fabricated from PEO/PCL composites with embedded fluorescent dye were utilized to image the fiber distribution within fiber-reinforced PEO hydrogels. Four vertical sections of these dye-containing PEO/PCL composites were obtained, and imaged on a laser scanning confocal microscope (LSCM, Olympus FV1000 BX2 Upright Confocal Microscope) to observe fiber/dye distribution throughout the hydrogel. Four images were obtained per vertical section. These images were divided into 16 equal coordinate sections using ImageJ, and a threshold analysis was applied to determine the fiber coverage (by area). Compressive Hysteresis. Uniaxial compression was performed on a Zwick/Roell instrument with a 100 N load cell. Hydrogel samples were prepared by swelling the corresponding dried xerogel in distilled H2O for 72 h. Following swelling, each hydrogel sample was trimmed to a cubic 8 cm3 nominal volume (2 cm × 2 cm × 2 cm) using a razor blade. Individual hydrogel specimens (minimum 5 specimens per sample) were placed between circular plates and compressed to 75% of the initial thickness (25% engineering strain) at a rate of 50% min−1. The compressive load was then removed, and the force unloading curve of each sample was recorded. This process was repeated 19 more cycles to obtain 20 cyclic loading/unloading force curves, which were normalized by the initial dimensions of each specimen. The stiffness of each specimen was calculated as the slope of the linear derivative of the first loading curve at the point of increased load bearing between 10 and 15% compressive engineering strain. The stability of each specimen was assessed as the normalized integration ratio between the first and twentieth loading/unloading cycle. Porcine Cartilage Samples. Articular cartilage with full thickness of 1.3 ± 0.2 mm was harvested from the knee joint of a pig (approximately 2.5 years of age) with consent of a local farmer (Amherst, OH) and stored in phosphate buffered saline (PBS, pH 7.4) from the time of harvest until testing (∼12 h). Porcine cartilage was chosen as a model system due to its relative similarity to human cartilage and its availability. Care was taken to ensure no unnecessary suffering of the donor pig in accordance with established ethical research practices.57 Gel Strength Analysis. The strength of each sample was analyzed via small amplitude oscillatory shear (SAOS, TA Ares G2) using parallel 25 mm plates with a minimum of 3 specimens each. The amplitude of deformation was set to 1% in the linear viscoelastic region, and a range of frequencies (ω) were examined between 1 × 10−2 to 1 × 102 Hz at room temperature. The plateau modulus was determined as the stable storage modulus (G′) region of each frequency sweep. Loss modulus (G″) and G′ curves were plotted as the point-by-point average of 3 specimens. NIH 3T3 Cell Culture. NIH 3T3 cells were cultured in a T-75 cm2 flask using Dulbecco’s modified eagle medium (DMEM) with addition of 10% newborn calf serum (NCS), 1% penicillin (Invitrogen), and 1% GlutaMAX (Gmax) in a humidified atmosphere (37 °C, 5% CO2). The growth medium was changed every 48 h. Upon reaching 80−90% confluency, NIH 3T3 cells were detached using a cell detachment solution containing PBS and ethylenediaminetetracetic acid (EDTA) at 37 °C for 10 min. The detached cells were collected by centrifugation at 500 g for 5 min. After removing the supernatant above the cell pellet, cells were resuspended in new media. Cells were counted using a hemocytometer before seeding them in each sample.

Figure 1. Schematic representation for the formation of in situ fiberreinforced PEO hydrogels.

fiber material

(

XD =

vf =

fiber loading (wt %)

3 5 Wf

investigate glassy vs elastic fiber elements

hydrogel (Figure 1, Step 6). Control PEO hydrogels containing no fibers were also fabricated under similar processing conditions. Gel Fraction and Equilibrium Swelling Determination. After determining the fiber weight fraction (Wf) via 1H NMR (Varian Anova 600 MHz), the fiber mass fraction of each xerogel (A1) was calculated (eq 1). Five individual samples were evaluated to determine both gel fraction (Gf) and the equilibrium swelling ratio (SR) of the PEO hydrogel. The initial mass of each specimen (m1) was recorded before submerging in distilled H2O for 72 h to ensure complete removal of uncross-linked PETA and PEO (Figure S3). Distilled H2O was chosen for this study instead of organic solvents to prevent dissolution of the polyester fiber components. After swelling of the cross-linked PEO, the mass of each specimen (m2) was recorded. Swollen samples were dried under vacuum for 24 h, and the final dry mass of each specimen was recorded (m3). The mass of hydrophobic, polyester fiber filler was accounted for in each sample to ensure the determination of Gf and SR were based solely on the PEO hydrogel phase (eqs 2-3). The volume fraction fiber content (vf) in each hydrogel sample was calculated using Wf and the average SR and Gf of each sample (eq 4). Optical images of the xerogels and hydrogels were captured with a Canon (EOS Rebel T3i) camera. C

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ACS Biomaterials Science & Engineering 3D Cell Culture. Hydrogels derived from coextruded PEO (control), undrawn coextruded PCL/PEO, drawn coextruded PCL/ PEO, and undrawn PLLA/PEO composites (n = 3) were evaluated as 3D culture platforms using the above cultured NIH 3T3 cells (5th passage). Hydrogel samples (300 mg) were injected with 7 × 105 cells. Cells were grown for 72 h at 37 °C. After 72 h, growth medium was removed and NIH 3T3 cells were washed with PBS three (3) times. Cells were fixed using 4% (w/v) paraformaldehyde for 15 min at room temperature and again washed with PBS 3 times. The NIH 3T3 cells were permeabilized with 1 mL of 0.2% (w/v) Triton X-100 in PBS for 5 min before washing 3 times with PBS. The fixed cells were incubated with ActinGreen488 (excitation wavelength of 488 nm) for 30 min using 2 drops/ml of media before washing 3 times with PBS. The cell nuclei were stained using 4′,6-diamidino-2 phenylindole (DAPI). DAPI (100 μL) was diluted in 10 mL of DPBS (5 μL DAPI/ml DPBS) before staining cell nuclei for 10 min at room temperature with subsequent washing 3 times using DPBS. Each sample was mounted on a glass slide, and images were obtained using LSCM (Leica TCS SPE Confocal Microscope). NIH 3T3 Cell Viability. Cell viability (n = 3) for the control PEO hydrogel and the undrawn PCL fiber-reinforced PEO hydrogel was assessed using an 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay after 72 h of cell growth. An MTT solution (5 mg/mL PBS) was prepared and sterilized via microfiltration (0.2 μm filter size). In the absence of light, 500 μL of DMEM/MTT (85/15 by volume) solution was added to cells grown in each sample and incubated for 4 h. Following incubation, the solution was removed and replaced with 1 mL of dimethyl sulfoxide (DMSO, Sigma-Aldrich) to dissolve the formazan crystals by shaking at 100 rpm for 30 min. Relative viability between the two sample types was determined using relative absorbance at 570 nm. MC3T3-E1 Cell Culture. MC3T3-E1 Subclone 4 cells were cultured in Alpha minimum essential medium (MEM) with ribonucleosides, deoxyribonucleosides, 2 mM L-glutamine, and 1 mM sodium pyruvate, but without ascorbic acid. To prepare complete growth medium, we included 10% fetal bovine serum (FBS) and 1% penicillin in the MEM medium. Cells were incubated in 75 cm2 cell culture flasks at 37 °C in a 95% air and 5% CO2 environment. At 80− 90% confluency, the cells were detached with PBS/EDTA for 10 min at 37 °C. The detached cells were collected by centrifugation at 800 g for 5 min. The in situ fiber reinforced hydrogels (control, undrawn PCL, drawn PCL, and undrawn PLLA) were each injected with 5 × 105 cells in a 24-well cell culture plate (n = 6 for each type) and cultured at 37 °C, 5% CO2 in a humid environment for 7 days. The cell culture medium was refreshed every other day (5th passage). MC3T3-E1 Calcification. To evaluate localized calcium content within the hydrogel matrix, cells were washed with DPBS 3 times. Cells in each matrix were fixed with 4% paraformaldehyde for 10 min at room temperature and then incubated with 2% Alizarin red solution (titrated to pH = 4.2 by 0.5% ammonium hydroxide) at room temperature for 5 min. After incubation, the control and three in situ (undrawn PCL, drawn PCL, and undrawn PLLA) fabricated PEO hydrogel samples were washed to remove excess Alizarin red with PBS buffer five times. All samples were mounted on glass slides and covered using a glass coverslip sealed with nail polish. Hydrogels were imaged using an optical microscope (Olympus BX 51).

composites contained between 0 and 30.5% PCL by weight, resulting in hydrogel fiber loadings between 0 and 7.8% PCL fiber by volume (vf) (Figure S2). The lowest vf obtained for any of the fiber reinforced hydrogels was calculated as 0.1 ± 0.02 vol %, demonstrating the tunability of fiber loading over 2 orders of magnitude utilizing this in situ fabrication strategy. The largest lateral fiber dimensions were kept under 10 μm (9.3 ± 1.7 μm width by 6.1 ± 1.2 μm thickness), well below the 40 μm threshold for efficient fiber reinforcement, allowing fibers to deform under the contact forces of the hydrogel matrix acting upon individual fibers (Figure S4).36 The composite resulting from the multilayer coextrusion process contained thousands of individual polyester fibers of infinite length surrounded by a PEO matrix because of the continuous nature of the process. PCL fibers functioned as a soft elastomer at room temperature,51 whereas PLLA served as a rigid reinforcing element at room temperature with a Tg of 61 °C and a Tm of 152 °C (Figure S5). In contrast to conventional PEG hydrogel cross-linking, this in situ fabrication method enables direct incorporation of exfoliated micron-scale fibers; however, irregularity in the coextruded PEO hydrogel network is a potential disadvantage due to the nonspecific cross-linking pathway. During hydrogel formation via UV irradiation, a number of variables influence architecture and mechanics, including cross-linker functionality, radiation intensity, cross-linker:PEO concentration ratios, and exposure time. PETA, which contained three acrylate moieties activated with UV radiation (λ = 320−390 nm), was chosen as the cross-linker. PETA may either react with other PETA molecules or abstract a hydrogen atom from the PEO backbone before radical coupling between PEO and PETA to cross-link PEO.16 Both of these reaction pathways likely occur, but the high gel fractions observed suggest that the majority of the PEO chains undergo covalent cross-linking under sufficient PEO:PETA ratios (40−70 wt % PEO) and UV exposure times (20−120 min). By systematically varying the PEO:PETA concentration ratio and UV irradiation time, an appropriate balance of water uptake and gel fraction was found using a 60:40 (PEO:PETA by weight) ratio and 30 min UV irradiation (Figure S6). These conditions were used throughout this investigation to maintain similar gel fractions (∼70%) in PEO hydrogels imbibed with either PCL or PLLA fibers. Hydrogel Architecture. The hydrogel fabrication process yielded a dried film, or a “xerogel” intermediate, which was swollen in distilled water for 72 h to form the hydrogel (Figure 2a). A nominal volume composition was calculated based on feed rate and compared with experimentally determined composition of each sample based on 1H NMR integrations. The experimentally determined PCL composition was systematically lower (∼2−5%) than the calculated values, likely due to perturbations in feed rate monitoring (Figure S2). A constant ratio PEO:PETA was utilized for each fiber loading to obtain a similar gel fraction in each hydrogel (Figure S7a), maintaining gel matrix mechanics and allowing comparison of reinforcement behavior based solely on PCL fiber loading. Even at similar gel contents, a clear decrease in water uptake as the PCL fiber volume fraction increased was observed, despite controlling for the nonswellable hydrophobic fibrous phase (Figure S7b). This swelling behavior is likely due to PCL fibers within the hydrogel matrix restricting expansion with water uptake, which has been previously reported in fiber-reinforced hydrogels fabricated via additive manufacturing techniques.39

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RESULTS & DISCUSSION In Situ Hydrogel Fabrication Technique. Having established the applicability of this continuous, solvent-free, and scalable processing technique to fabricate functional PCL fiber scaffolds, this research highlights the in situ fabrication of micro- and nanoscale fiber reinforced hydrogels utilizing coextruded composites. For this work, a series of polyester fiber/PEO matrix composites were fabricated via multilayer coextrusion (Table 1). Implementing a one-step PEO crosslinking reaction postextrusion allowed high-volume hydrogel production utilizing commodity polymers. The coextruded D

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dimensions of individual fibers were indeed on the micronscale (Figure S8). To visualize fiber distribution throughout the hydrogel volume, we embedded a fluorescent dye within PCL fibers during multilayer coextrusion. The in situ hydrogel manufacturing technique was then used to transform these coextruded composites to fiber-reinforced hydrogels. Laser scanning confocal microscopy (LSCM) analysis of these hydrogels also revealed distinct regions of high dye intensity with similar size scale to the PCL fiber phases formed during coextrusion of PCL/PEO composites, suggesting a significant amount of dye was retained upon hydrogel formation. Depth imaging of these dye-load PCL/PEO hydrogels (∼7.8 vol %) was obtained under a wide field of view (210 × 210 μm) (Figure 2b). Representative LSCM images were recorded at four z-depths, highlighting individual fiber features (Figure 2b, left column). Quantitative analysis of these fiber-reinforced PCL/PEO hydrogels were conducted at each depth, sectioning into a matrix of four vertical columns (1−4) and four horizontal rows (A−D) and calculating the relative fiber coverage was calculated for each of the 16 sections at all four depths (Figure 2b, right column). The variability represented by the error bars is attributed to the relatively low fiber loading. However, there is good agreement between the relative fiber coverage obtained via LSCM analysis and the fiber composition calculated based on eq 4. Visual analysis via SEM and LSCM imaging support achievement of two key design characteristics for synergistic mechanical reinforcement in fiber-loaded hydrogels: (1) micrometer-scale fiber inclusion, and (2) fiber distribution throughout the hydrogel volume. Limited evidence of a third design consideration−random fiber alignment−was also revealed. Additional support for this final design aspect will be discussed later. Effect of Fiber Loading. Two key parameters were utilized to assess the reinforcing effect of the hydrogels: compressive mechanical stability and stiffness. Mechanical stability can be defined as the relative hysteresis ratio between the 20th loading−unloading and first loading−unloading cycle (Figures S9−S12), and was determined from compressive hysteresis experiments (Figure 3). Stiffness was determined by taking the linear derivative of the first loading curve and determining the slope of the derivative, which reflects the both the degree of concavity of the loading curve and the maximum compressive stress (Figure S13). Porcine articular cartilage was utilized as an ideal standard for hydrogel mechanics due to its similarity to adult human articular cartilage.58 The hysteresis loop obtained from porcine articular cartilage revealed near ideal mechanical stability (94.5 ± 1.7%) and minimal resistance to loading until approximately 15% compressive engineering strain before a sharp increase in resistive force, both key requirements of this supportive load bearing biological tissue, known as a “J-curve” loading profile. The characteristic force−displacement “J-curve” displayed by most biological tissues is characterized by a large initial displacement requiring minimal force followed by a drastic force increase required to achieve additional deformation.59 Examination of the control PEO hydrogel revealed a lack of the characteristic “J-curve” behavior, coupled with extremely poor mechanical stability (6.8 ± 2.1%). It should be noted that even a minimal fiber loading (∼0.1 vol %) shifts the characteristic loading curve from gradual to a two-step process, similar to that observed in the cartilage sample. The mechanical stability of each sample was quantified (Figure 4a). Quantitative analysis reveals a significant increase in mechanical stability even at low

Figure 2. (a) Optical image of a xerogel and corresponding swollen hydrogel sample containing undrawn PCL fiber elements at a volume fraction of 7.8 ± 1.5%; (b) LSCM images showing the fluorescent dyeloaded PCL fiber (yellow) distribution throughout the body of the hydrogel (black).

A random distribution of fibers has been identified as a critical requirement for efficient mechanical reinforcement in hydrogel constructs.38 Cross-sectional SEM images of PCL fiber/PEO hydrogels demonstrate that fiber integrity was maintained during hydrogel formation, and the lateral E

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Figure 3. Representative 1st and 20th loading−unloading compressive hysteresis loops for the control PEO hydrogel which contains no fibers, varying fiber loading composition, and porcine articular cartilage.

Figure 4. (a) Hydrogel stability increased with increased fiber loading; (b) hydrogel stiffness fit well with the predictive model based on the Halpin− Tsai model with a sharp increase in stiffness between 1.2 vol % and 2.4 vol %; (c) phenomenological deformation model of in situ fabricated fiber reinforced hydrogels.

fiber loadings with stability ratios of 18.5 ± 3.5% (vf ≈ 0.1 vol %), 22.7 ± 4.6% (vf ≈ 0.4 vol %), and 27.8 ± 4.1% (vf ≈ 1.2 vol %) before increasing dramatically to a final ratio of 72.7 ± 7.3% (vf ≈ 7.8 vol %). The ability to tune the characteristic compressive loading and unloading curves of PEO hydrogels with the addition of fibers using the in situ fabrication technique from one-stage gradual loading to a two-stage “Jcurve” represents an important mechanics innovation in loadbearing hydrogels, as the “J-curve” permits minor displacement without resistance before a steep increase in resistance to further, undesirable loading. Mimicking the characteristic “J-

curve” is critical for in vivo applications; if the mechanics do not match that of the biological soft tissue, the healing process, long-term use, and performance are negatively impacted.60 It should be highlighted that, although enhanced and tunable mechanical stability was achieved, other critical factors, including stiffness and lubrication, must be considered in the development of synthetic cartilage.61 Beyond hydrogel mechanical stability, PCL fiber-reinforced hydrogel stiffness was also investigated as a function of fiber loading (Figure 4b) for comparison to articular cartilage (1.61 ± 0.17 MPa). Previous studies have reported the stiffness of F

DOI: 10.1021/acsbiomaterials.7b00229 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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ACS Biomaterials Science & Engineering

fiber reinforced PEO hydrogels were fabricated from composites with similar PCL content (Figures S18−S20). Hydrogels derived from drawn PCL/PEO composites at a similar loading possessed a gel fraction of 71.5 ± 4.1% and an equilibrium swelling ratio of 2.9 ± 1.1, which was comparable to the undrawn PCL/PEO hydrogels (Figure S21). This similarity in both gel fraction and water uptake suggested that the matrix gel structure was similar for undrawn and drawn PCL/PEO hydrogels, allowing isolation of the effect of chain alignment in the reinforcing fiber elements. Interestingly, the 2D WAXS profile of the PEO hydrogel containing drawn PCL fibers displayed isotropic scattering patterns for both PEO and PCL (Figure S22). The isotropic scattering from both the PEO and PCL phases was expected. Chain dissolution during the hydrogel fabrication process induced random ordering of the PEO chains. The isotropic scattering observed from PCL can be attributed to random alignment of individual fibers within the hydrogel. This confirmed our previous observations via SEM and LSCM and satisfied the third design criteria: random fiber alignment. After isolating the effect of fiber modulus, compression analysis revealed similar mechanical stability between hydrogels containing undrawn (72.7 ± 7.3%) and drawn (73.1 ± 8.2%) PCL fibers (Figure S24). However, Kc increased significantly to 6.31 ± 0.18 MPa with the 820% increase in fiber modulus during PCL drawing. Although this represented an increase of 225% in Kc (Figure 5b), it was significantly lower than the predicted Kc of 11.4 MPa from the Halpin−Tsai model, which was attributed to discrepancies in the term η due to the extreme disparity between Ef and Em.65 Yet, a high degree of synergistic reinforcement between the drawn PCL fibers and the PEO hydrogel was achieved (Figure S14c). Not only was cooperative reinforcement behavior observed under static compressive loading, but also during dynamic small amplitude oscillatory shear (SAOS) deformation (Figure 5c). Over 4 orders of magnitude in frequency, the PEO hydrogels with no fibers, undrawn PCL fibers, and drawn PCL fibers all exhibit gel-like behavior. By incorporating undrawn PCL fibers, the plateau storage modulus (G′) increased ∼56% from 2.5 ± 0.2 MPa to 3.9 ± 0.3 MPa. Strikingly, uniaxial drawing before hydrogel fabrication increased G′ to 12.3 ± 1.2 MPa, an enhancement of 390% compared to the control PEO hydrogel and 215% compared to the undrawn PCL fiber reinforced PEO hydrogel. Material Selection. The modular nature of the multilayer coextrusion process allowed processing of a PLLA/PEO composite to obtain a PLLA fiber reinforced PEO hydrogel with vf = 7.5 ± 1.6 vol %, similar to that of undrawn (highest loading) and drawn PCL fiber reinforced PEO hydrogels. Rectangular PLLA fibers with dimensions 2.3 ± 0.5 μm (width) and 1.5 ± 0.4 μm (thickness) were isolated from coextruded composites with an elastic modulus of 167 ± 24 MPa (Figure S25). The lack of PLLA chain alignment (Figure S26) allowed a comparison between elastic and rigid fiber elements using undrawn PCL and undrawn PLLA fibers (Figure 5a). PEO hydrogels reinforced with undrawn PLLA fibers exhibited similar water uptake (3.0 ± 0.7 g H2O/g Gel) and gel fraction (68.6 ± 5.2%) compared to reinforcement with undrawn PCL fibers (Figure S21). Using the Halpin−Tsai model and the determined Ef of coextruded PLLA fibers, the overall Kc was predicted to be 13.3 MPa; however, the observed Kc was 8.76 ± 0.21 MPa (Figure 5b). Again, we attributed this discrepancy to the large modulus

traditional low-molecular weight end-functionalized PEG hydrogels to range between 13 and 233 kPa, varying with photoinitiator content.62 Because of the high viscosities required for coextrusion, it was not feasible to use conventional end-functionalized PEG in this process. Using high molecular weight PEO and UV cross-linking yielded hydrogel stiffness of 0.69 ± 0.04 MPa. Incorporating PCL fibers increased hydrogel stiffness to 1.94 ± 0.21 MPa at ∼7.8 vol % fiber loading. The enhancement in hydrogel stiffness was not explained by a simple volume additive model because of an underestimation of Kc, suggesting a cooperative and synergistic reinforcement phenomenon (Figure S14). To understand the synergistic impact with fiber loading, we fitted the stiffness data using a modified form of the Halpin−Tsai model (eqs 4−7).63 ⎛ 1 + ξηvf ⎞ Kc = K m ⎜ ⎟ ⎝ 1 + ηvf ⎠

(5)

ξ = 1 + (40vf )10

(6)

η=

Ef Em Ef Em

−1 +ξ

(7)

The predicted hydrogel stiffness (Kc) was determined to be a function of matrix stiffness (Km), which was taken as the stiffness of the hydrogel without fiber elements, and vf. The geometric shape factor (ξ) was determined using the HewittMalherbe approximation, which relates ξ to vf.64 By relating ξ to vf instead of fiber aspect ratio, it was possible to use the Halpin−Tsai model to predict Kc, as the theoretical aspect ratio of individual fibers varied by altering coextrusion feed rates (Table S5). The reinforcement factor (η) accounted for the tensile stiffness of both the fibrous PCL elements (Ef), determined previously to be 15.0 ± 1.8 MPa,51 and the matrix material (Em), determined to be 0.45 ± 0.09 MPa (Figure S15). The utilization of tensile elastic constants can be explained by the structural changes experienced during deformation of each hydrogel sample (Figure 4c). In the uncompressed resting state, the randomly distributed, contorted and crimped fibers are surrounded by gel matrix. During compressive loading, the hydrogel matrix is compressed in the direction of applied force, resulting in axial expansion perpendicular to the applied force. During the first compressive phase, the crimped fibers were straightened, requiring minimal deformation force. While under compressive loading (2nd stage), these straightened fibers require significantly more contact force for continued deformation under tension, resulting in a significant increase in deformation force. Together, these two phases produced the “J-curve” observed in Figure 3. It is clear from the crosssectional SEM images that some individual PCL fibers were contorted during the matrix solvation step, resulting in a significant decrease in fiber aspect ratio. Ideally, in a percolated network architecture, straight, rigid fibers are completely exfoliated. The aspect ratio is the ratio of length-to-diameter for circular fibers, or length-to-width/thickness for rectangular fibers (Figure S16, Table S5). The contortion and crimping of fibers was confirmed by determining an effective fiber aspect ratio of 95, which was significantly less than the predicted theoretical range (650−2380) and confirmed via SEM analysis (Figures S8 and S17). PCL Chain Orientation. To explore the impact of composite drawing on hydrogel mechanics, drawn and undrawn G

DOI: 10.1021/acsbiomaterials.7b00229 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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ACS Biomaterials Science & Engineering

Hydrogel Viability as Cell Scaffolds. Three-dimensional hydrogel scaffolds provide an inclusive cellular growth environment with pathways for nutrient influx and waste efflux.66 Fiber reinforced hydrogels were evaluated as a 3D tissue culture model using NIH 3T3 murine fibroblast cells, a common in vitro model for connective tissue (Figure 6).

Figure 6. Fibroblast cell adhesion and proliferation in PEO hydrogel containing (a) no fibers; (b) undrawn PCL fibers; (c) drawn PCL fibers; (d) undrawn PLLA fibers: the cytoskeleton is stained with ActinGreen488 and the cell nuclei are stained with DAPI (blue).

Murine fibroblast cells were injected into the interior volume of each hydrogel system and incubated for 72 h. Following 72 h incubation, cells were first stained with ActinGreen488 to visualize the cytoskeleton and DAPI (blue) to visualize the nuclei of adhered fibroblast cells. Confocal micrographs showed fibroblast cell growth in the control PEO hydrogel with cytoskeletal connections between cell nuclei (Figure 6a). As a proof of viability, hydrogels derived from undrawn PCL/PEO (Figure 6b), drawn PCL/PEO (Figure 6c), and undrawn PLLA/PEO (Figure 6d) composites at ∼8 vol % fiber loading were also injected with fibroblast cells. All three fiber reinforced hydrogel systems displayed similar cell density and morphology, which suggested that embedded fibers did not perturb cellular morphology. Additionally, there was no observed decrease in cell viability when comparing the control PEO hydrogel and PCL-fiber reinforced hydrogel (Figure S29). Having demonstrated the viability of in situ fiber reinforced PEO hydrogels as tissue engineering scaffolds, there is also the possibility to utilize the tunable mechanical nature of these hydrogels to direct cell differentiation.67−69 MC3T3-E1 subclone 4 cells were injected into four in situ fabricated hydrogel systems with mechanics (Kc and G′) tunable over an order of magnitude (Figure 5). The MC3T3E1 cell line expresses markers for differentiation into either a fully calcified phenotype or collagen type I based on chemical cues,70 mechano-transduction,71 or rigidity of the culture substrate.72 Based on these previous studies, it was anticipated that mineralization would be enhanced as the stiffness of the

Figure 5. (a) Optical images of the control PEO hydrogel without fibers, PEO hydrogel containing 7.8 ± 1.5 vol % undrawn PCL fibers, PEO hydrogel containing 7.2 ± 1.3 vol % drawn PCL fibers, and PEO hydrogel containing 7.5 ± 1.6 vol % undrawn PLLA fibers; (b) stiffness of the three hydrogel samples determined from uniaxial compression; (c) dynamic SAOS sweeps of the four hydrogel samples.

ratio (∼242) between the PLLA fiber phase and PEO hydrogel matrix.65 However, a synergistic reinforcement effect between matrix and fiber was indeed achieved (Figure S14c). When compared with PEO hydrogels reinforced with undrawn PCL fibers the stiffness of PLLA fiber reinforced hydrogels increased 350%. The mechanical stability of PLLA fiber hydrogels also increased to 80.4 ± 5.4% (Figure S27). In addition, the dynamic SAOS shear modulus increased to 27.4 ± 1.4 MPa, representing an increase of 600% over the PEO hydrogel reinforced with undrawn PCL fibers at similar vf (Figure 5c). The ability to tailor hydrogel stiffness and shear modulus over an entire order of magnitude simply by tuning processing parameters and material selection presents an incredibly unique avenue to achieve targeted stem cell differentiation as the rigidity of the environment that cells are exposed to has been shown to influence cell differentiation pathways. H

DOI: 10.1021/acsbiomaterials.7b00229 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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ACS Biomaterials Science & Engineering

Figure 7. MC3T3-E1 preosteoblast differentiation after injection into in situ fabricated PEO hydrogels derived from (a) PEO control; (b) undrawn PCL/PEO composite; (c) drawn PCL/PEO composite; (d) undrawn PLLA/PEO composite.

technique to achieve varying degrees of calcification using murine MC3T3-E1 progenitor cells based on 3D hydrogel mechanical environment, which was controlled strictly by fiber type. The ability to fabricate distributed, submicron-scale fiberreinforced synthetic, biocompatible hydrogels which enhanced cell adhesion and proliferation in a tunable, scalable process presents a powerful technological tool for the cell scaffolding and tissue growth communities.

hydrogel environment increased. Cells were incubated fully surrounded by the different mechanical environments provided by the control PEO hydrogel (Figure 7a), and in situ fabricated fiber-reinforced PEO hydrogels with undrawn PCL (Figure 7b), drawn PCL (Figure 7c), and undrawn PLLA (Figure 7d) fiber phases. It is important to remember that vf, SR, and Gf for all of these in situ systems were similar, and Kc and G′ were influenced strictly by fiber type. The Alazarin Red-s stained the cultured cells for calcium phosphate, which appeared an orange-red under the optical microscope and indicated a high calcification content. This calcification marker is indicative of differentiation down the fully mineralizing phenotype, forming calcified ECM, characteristic of true bone. The lack of calcium phosphate present in the control PEO hydrogel culture suggests that cells surrounded by this mechanically weak environment did not fully mineralize. Although strictly qualitative, it was quite exciting to observe that, as gel strength increased, the preference for cellular osteogenesis was enhanced. This promising finding suggests that the robust, flexible in situ hydrogel fabrication platform can also be tailored with specific mechanical responses to induce cellular behavior for targeted differentiation as 3D growth scaffolds.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsbiomaterials.7b00229. Material characterization, multilayer coextrusion analysis, derivation and calculation of SR and Gf, details of compressive mechanical analysis and modeling, PCL and PLLA fiber orientation, distribution and architecture, and PEO hydrogel morphology analysis (PDF)

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Phone: (216) 368-1421.

CONCLUSIONS An innovation in multilayer coextrusion technology allowed the fabrication of two fiber/matrix composites, PCL/PEO and PLLA/PEO. A straightforward cross-linking strategy was utilized to transform these fiber/matrix composites into PEO hydrogels reinforced with either PCL or PLLA fibers. This in situ fiber-reinforced hydrogel manufacturing process presents an unprecedented modularity to yield tunable hydrogels from commodity polymers with high production volume not possible with complex synthetic routes or expensive components. Exfoliated rectangular fibers with lateral dimensions below 10 μm were shown to be well-distributed and randomly aligned throughout the PEO hydrogel matrix, resulting in synergistic reinforcement between the individual fibers and the gel matrix. By varying both fiber loading and fiber type, hydrogel stiffness was tailored between 0.69 ± 0.04 MPa and 8.76 ± 0.21 MPa, while storage modulus was tailored between 2.5 ± 0.2 and 27.4 ± 1.4 MPa. These fiber reinforced PEO hydrogels fabricated in situ provided an excellent 3D culture environment that supported fibroblast proliferation and adhesion, demonstrating their viability as tissue engineering scaffolds. Finally, we exploited the modular nature of the in situ manufacturing

ORCID

Jonathan K. Pokorski: 0000-0001-5869-6942 LaShanda T. J. Korley: 0000-0002-8266-5000 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors thank Profs. Eric Baer and Gary Wnek for their helpful discussions on gel mechanics. The authors also acknowledge funding from the National Science Foundation (NSF) Center for Layered Polymeric Systems (CLiPS) under Grant DMR-0423914 and NSF funding under Grant CMMI1335276.

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REFERENCES

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DOI: 10.1021/acsbiomaterials.7b00229 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX