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Cite This: Biomacromolecules 2019, 20, 2684−2693

“Tree to Bone”: Lignin/Polycaprolactone Nanofibers for Hydroxyapatite Biomineralization Ding Wang, Jinhyeong Jang, Kayoung Kim, Jinhyun Kim, and Chan Beum Park* Department of Materials Science and Engineering, Korea Advanced Institute of Science and Technology (KAIST), 291 Daehak-ro, Daejeon 34141, Republic of Korea

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ABSTRACT: Bone contains an organic matrix composed of aligned collagen fibers embedded with nanosized inorganic hydroxyapatite (HAp). Many efforts are being made to mimic the natural mineralization process and create artificial bone scaffolds that show elaborate morphologies, excellent mechanical properties, and vital biological functions. This study reports a newly discovered function of lignin mediating the formation of human bone-like HAp. Lignin is the second most abundant organic material in nature, and it exhibits many attractive properties for medical applications, such as high durability, stability, antioxidant and antibacterial activities, and biocompatibility. Numerous phenolic and aliphatic hydroxyl moieties exist in the side chains of lignin, which donate adequate reactive sites for chelation with Ca2+ and the subsequent nucleation of HAp through coprecipitation of Ca2+ and PO43−. The growth of HAp crystals was facilitated by simple incubation of the electrospun lignin/polycaprolactone (PCL) matrix in a simulated body fluid. Multiple analyses revealed that HAp crystals were structurally and mechanically similar to the native bone. Furthermore, the mineralized lignin/PCL nanofibrous films facilitated efficient adhesion and proliferation of osteoblasts by directing filopodial extension. Our results underpin the expectations for this ligninbased biomaterial in future biointerfaces and hard-tissue engineering.

1. INTRODUCTION The synthesis of bone-like ceramic materials for skeletal tissue regeneration is in high demand within clinical medicine. The nucleation and growth of minerals in living organisms are a key process for the development of skeletal frames. The process is typically mediated by bioorganic macromolecules, forming hierarchically well-organized composite materials.1 Osteoblast matrix vesicles accumulate Ca2+ and PO43− from the cytosol and mitochondria which are released toward the newly formed collagen matrix, until the precipitation of amorphous calcium phosphate (ACP) occurs.2 The charged regions of the collagen framework act as nucleation sites for crystal growth by converting the high levels of Ca2+, PO43−, and ACP into crystalline plate-like hydroxyapatite (HAp)a calcium phosphate crystal with an empirical formula of Ca10(PO4)6(OH)2the mineral phase of sclerous tissues (e.g., bone, dental enamel, and dentin).3 Inspired by natural mineralization, various biomolecules (e.g., silk fibroin,4 collagen,5 peptide,6 alginate,7 chitosan,8 and cellulose9) and synthetic polymers (e.g., polydopamine,10 poly(lactic acid),11 polyacrylamide,12 and polyethylene glycol13) have been considered as biomimetic mineralization templates, but these systems have some shortcomings such as the expensive ingredients, harsh deposition conditions, limited mechanical properties, and weak mineralization capability.14 Moreover, © 2019 American Chemical Society

compared to direct addition of bioactive materials (e.g., HAp, tricalcium phosphate, and biphasic calcium phosphate ceramics) into biodegradable polymers for the synthesis of organic−inorganic hybrid biomaterials, biomineralization often requires fewer chemical reaction steps and greener processing and results in sophisticated materials with relatively high stiffness and affirmative biological functions.15 Therefore, it is critical to design multifunctional biomaterials that can biomimetic induce and assemble bone-like HAps close to human bone. Lignin is the second most abundant organic molecule and mainly exists in the cell walls of vascular plants. It is a randomly cross-linked polymer consisting of monolignols and phenylpropanes in different proportions between plant species. More than 70 million tons of lignin byproducts are generated annually from the pulp industry; however, only 2% of this amount has been commercialized to formulate dispersants, adhesives, and surfactants for plastics and rubbers.16 Many recent studies reported lignin’s high durability, thermal stability, antibacterial, good biocompatibility, and capability of protecting cells from oxidative stress effects, making it Received: March 31, 2019 Revised: May 18, 2019 Published: May 22, 2019 2684

DOI: 10.1021/acs.biomac.9b00451 Biomacromolecules 2019, 20, 2684−2693

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Biomacromolecules

surface area for enhanced cell attachment and nutrient transportation. In situ biomineralization of HAp on the nanofibrous surface can not only enhance its strength and toughness but also provide a favorable substrate for cell proliferation and osteogenic conduction.21 We validated the structural and mechanical properties of lignin-induced bone mineral phase using multiple analytical tools. We further confirmed that osteoblastic cells adhered and proliferated well on the lignin/PCL−HAp composites, showing that the combination of lignin and HAp can promote osteointegration as a bioresorbable bone cement. Compared to the cellulose− HAp composite that often requires a series of special conditions for the synthesis (e.g., acid or basic buffer,1c,22 high concentration of SBF,23 and long-term incubation24) and shows low osteoconductivity,25 the lignin/PCL−HAp composite was synthesized under mild and biocompatible conditions, which exhibited a good osteoconductivity, making them suitable for potential bone therapies.

potentially useful for healthcare applications (e.g., drug carrier for cancer therapy, free-radical scavenger, and antibacterial agent).17 However, the brittle nature of lignin, its highly complex three-dimensional (3D) structure, and its incompatibility with nonpolar polymers18 have led to increasing interest in developing homogeneous and elastic lignin-based biomaterials. A facile strategy to synthesize lignin-based matrix is to blend lignin within a polar polymer because lignin’s phenolic hydroxyl-rich structure offers an opportunity to create intermolecular hydrogen bonds with electronegative groups (e.g., ether and carbonyl groups). Polycaprolactone (PCL), an aliphatic polyester, has been widely utilized for medical purposes because of its unique mechanical properties, ease of processing, biocompatibility, and biodegradability, but it has some limitations because of its hydrophobicity, bulk crystallinity, and triggering oxidant stress in local tissue.19 Thus, PCL is a prime candidate used as the blender with lignin to obtain a miscible structure, which can overcome the shortcomings of the use of PCL and lignin alone including the mechanical property, biocompatibility, and better function (e.g., hydroxyl groups).20 Here, we report a newly discovered function of lignin: to mediate interfacial biomineralization and integrate continuous bone-like HAp onto substrates with affirmative bioactivity and osteoconductivity, as depicted in Figure 1. Lignin contains a

2. EXPERIMENTAL SECTION 2.1. Materials. Kraft lignin (Mw = 28 000; prod. no. 370959), PCL (Mw = 80 000 according to gel permeation chromatographic analysis; prod. no. 440744), chloroform, and dimethylformamide (DMF) were purchased from Sigma-Aldrich Chemical Co., USA, and used without further purification. For cell culture, α-MEM medium, fetal bovine serum (FBS), and antibiotics (penicillin−streptomycin solution containing 5000 units/mL penicillin and 5000 μg/mL streptomycin) were attained from the Welgene Inc. Korea. 2.2. Preparation of Lignin/PCL Nanofibrous Film. Lignin/ PCL nanofibers were synthesized via employing an electrospinning process using a rotating drum collector (NanoNC Co., Korea).26 Lignin and PCL dissolved in chloroform/DMF (2:1 v/v) were mixed together to reach a final concentrations containing PCL, 200 mg/mL; lignin, 10 mg/mL. We poured the solution into a 10 mL syringe connected to a 21-gauge blunt tip needle (Cadence Science Inc., USA). A fixed voltage of 18 kV was applied, and a solution-feeding rate of 10 μL/min was established. Fibers were collected on a grounded wheel covered with aluminum foil that was placed at a distance of 18 cm from the needle end and rotated at a speed of 100 rpm. 2.3. Formation of HAp via Lignin as a Template. We prepared an SBF according to the widely used method designed by Kokubo and Takadama.27 The composition of 1.5× SBF (pH 7.5) is as follows: Na+, 213.0 mM; K+, 7.5 mM; Mg2+, 2.25 mM; Ca2+, 3.75 mM; Cl−, 221.7 mM; HCO3−, 6.3 mM; HPO42−, 1.5 mM; SO42−, 0.75 mM. For the formation of HAp crystals, each substrate was soaked in a 1.5× SBF and incubated at 37 °C in a shaking incubator for a predetermined time. Before characterization, the samples were rinsed with deionized water and dried under ambient conditions. 2.4. Characterization. Morphologies of films were characterized with scanning electron microscopy (SEM) using a 4800 field-emission scanning electron microscope (Hitachi High-technologies Co., Japan) at an acceleration voltage of 10 kV and transmission electron microscopy (TEM) using a JEM-3010 transmission electron microscope (JEOL Ltd., Japan) at an acceleration voltage 300 kV. For the SEM analysis, samples were sputtered with platinum using a SCD005 Pt-coater (Bal-Tec AG., Liechtenstein) before observation. For TEM and selected area electron diffraction (SAED) analysis, we carefully rubbed a carbon/formvar-coated Cu TEM grid with HAp@lignin/ PCL film to transfer HAp onto the grid. Energy-dispersive X-ray (EDX) analysis was performed to identify the elements in each film by using EMAX-7593H (Horiba Co., Japan). Attenuated total reflectance−Fourier transform infrared (ATR−FTIR) spectra were collected using a Nicolet iS50 FTIR (Thermo Scientific Inc., USA) spectrometer with one bounce reflectance diamond ATR crystal accessory. The ATR−FTIR spectra were recorded at room temperature at a 4 cm−1 resolution by averaging 32 scans in the spectral

Figure 1. Schematic description for the biomineralization of bone HAp induced by lignin. Lignin/PCL fibrous platform was prepared by electrospinning, followed by incubation in a SBF. Lignin contains a large number of hydroxyl groups, which can donate adequate reactive sites to bind with metal ions and facilitate the nucleation and growth of HAp through coprecipitation of Ca2+ and PO43−.

large number of phenolic hydroxyl and aliphatic hydroxyl groups, which can donate adequate reactive sites to bind with metal ions and possibly facilitate the nucleation and growth of HAp through coprecipitation of calcium and phosphate ions. We have successfully developed a lignin/PCL fibrous matrix by one-step electrospinning, which further induced the formation of HAp in a simulated body fluid (SBF). Electrospinning is a simple and cost-effective process for producing ultrafine polymeric micro/nanofibers in the form of a nonwoven membrane.18 Electrospun mats closely resemble native extracellular matrices, exhibiting high porosity and specific 2685


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Biomacromolecules range of 600−4000 cm−1. Raman spectra were collected using a LabRAM HR UV/VIS/NIR high-resolution dispersive Raman microscope (HORIBA Jobin Yvon, Co., France) by accumulating 60 scans in the range of 100−3500 cm−1 with a resolution of 5 cm−1. X-ray diffraction (XRD) patterns were attained by using a D/MAXRC thin-film X-ray diffractometer (Rigaku Co., Japan) equipped with a nickel filter under a scan speed of 5°/min and Cu Kα radiation wavelength of 1.5418 Å in the range of 5−60°. The formation of HAp was also investigated with X-ray photoelectron spectra (XPS) measurements by K-Alpha (Thermo VG Scientific Inc., UK). The surface roughness and elastic modulus of the specimens were obtained with atomic force microscopy (AFM) using an MFP-3D Origin atomic force microscope (Oxford Instruments Plc., UK) equipped with a CDT-FMR cantilever (Nanosensors Inc., Switzerland) under ambient conditions at a scan rate of 1.0 Hz/s. The AFM cantilever was made of silicon having an average elastic modulus of ∼150 GPa and spring constant of 80 N/m. The cantilever probe that has an average radius of 100 nm and elastic modulus of 500−1000 GPa was coated with diamond. We estimated the elastic modulus according to the Hertz model using the load−displacement (F−d) curves. The tensile properties of the nanofibers were analyzed using MTS Insight (MTS Systems Co., USA). A segment of the nanofibrous film with 30 mm gauge length and 10 mm width was used for the tests. The strain rate was 10 mm/min, and the tests were performed at room temperature. Five specimens were tested for each scaffold. The tensile strength at break and bulk Young’s modulus were determined from the stress−strain curves. 2.5. Osteoblastic Cell Culture. The MC3T3-E1 (subclone 4) cell line (ATCC, USA) derived from mouse osteoblasts was tested to confirm biocompatibility of the lignin/PCL fibrous platforms. The osteoblastic cells were cultured in a complete growth medium (αMEM medium with 10% FBS, and 1% antibiotics) and subcultured at least twice a week under a humidified environment with 37 °C and 5% CO2. 2.6. Cell Viability Assay. The viabilities of MC3T3-E1 cells grown on the developed platforms were confirmed by both LIVE/ DEAD assay kit (L3224; Thermo Fisher Scientific., USA), consisting of calcein AM and ethidium homodimer-1, and Cell Counting Kit-8 (CCK-8) assay kit (Dojindo Corp., Japan). For the LIVE/DEAD assay, the substrates were cut into small circular discs, fitted inside a 24-well culture dish (SPL Life Sciences Co. Ltd., Korea), sterilized with ultraviolet light exposure, and soaked in 70% ethanol for 2 h. Subsequently, cells were seeded on each substrate at a cell density of 2 × 104. After incubation for 48 h, cells were stained with calcein AM and ethidium homodimer according to the provided manual and observed by using the Eclipse 80i fluorescence microscope (Nikon Inc., Japan). For the CCK-8 assay, cells were cultured on a 96-well plate (SPL Life Sciences Co. Ltd., Korea) with the substrates cut into 5 mm diameter at a density of 3 × 104 cells mL−1 for 1, 3, 5, and 7 days. After that, 100 μL of the fresh medium with 10 μL of CCK-8 solution were added in each well, incubated for additional 4 h, and shaken for 300 s. Finally, the absorbance at 450 nm was measured using a Victor 3 microplate reader (PerkinElmer Inc., USA). 2.7. Cell Morphology. The cell morphology was investigated by using both SEM and fluorescent actin filaments staining. For SEM analysis, cells cultured on each substrate were fixed with 2.5% glutaraldehyde, washed with phosphate buffered saline, and dehydrated with graded ethanol series. For cytoskeleton organization analysis, cells were fixed with 3.7% formaldehyde for 15 min, permeabilized with 0.1% Trion X-100 for 5 min, and stained with phalloidin−tetramethylrhodamine B isothiocyanate (Sigma-Aldrich Chemical Co., USA) for 30 min under dark conditions. Actin filaments were observed by using a LSM 510 laser scanning confocal microscope (Carl Zeiss AG., Germany). The number of live cells was counted at 10 independent sites of each sample, and the projected area of each cell and cell morphology index were analyzed by the Image J software that is freely available at http://rsb.info.nih.gov/ij/. The statistical analysis was carried out by means of one-way analysis of variance (ANOVA, n ≥ 3).

3. RESULTS AND DISCUSSION 3.1. Synthesis and Characterization of Electrospun Lignin Fibers. We used commercially available, waterinsoluble kraft lignin for this study. Kraft lignin from wood sources accounts for approximately 85% of total lignin production in the pulp industry and is usually used as a fuel source.28 We prepared solutions of pure lignin in DMF over a wide range of concentrations (10−50 wt %). Our attempts to produce pure lignin-based fibers were unsuccessful; the lignin solutions were prone to electrospray with beaded forms, rather than into uniform fibrous mats (Figure S1). This phenomenon can be attributed to the relatively low molecular weight of lignin, resulting in the lack of substantial chain structures and/ or molecular entanglements.29 When we further increased the concentration of lignin above 50 wt %, the lignin solution’s jetting became uneven because of high viscosity, causing large droplets to be emitted from the spinneret onto the collector. Inelastic (or unentangled) polymer solutions often form fibers with a beads-on-string morphology or break into droplets and the addition of linear, high-molecular-weight polymers can facilitate the formation of uniform electrospun fibers.30 To address the difficulty of making pure lignin-based fibers, we employed PCL as a blender to promote the miscibility of lignin through hydrogen bonding between the hydroxyl groups of lignin and the carbonyl groups of PCL. As shown in the SEM images (Figure S2), we successfully prepared six lignin/ PCL nanofiber matrices via electrospinning by adding lignin in various concentrations ranging from 0 to 10 mg/mL. The nanofibers exhibited a darker brown color with increasing concentrations of lignin, as shown in the inserted digital images in Figure S2. According to our observation (Figure S2g), the average diameter of the nanofibers varied between 230 and 960 nm, and the diameter decreased with the increasing concentration of lignin. This is caused by the decreasing viscosity of precursor solutions with the rising lignin content because lignin has a lower molecular weight and a higher polydispersity index than PCL.31 With lignin concentration over 15 mg/mL, however, we observed the formation of some insoluble lignin particles in the lignin/PCL precursor solution, and beaded fibers were collected after the electrospinning process (Figure S2h). This is ascribed to the lack of interfacial adhesion between lignin and matrix polymers.20c ATR−FTIR spectroscopy spectrum of lignin/PCL fibers exhibited typical peaks for both lignin and PCL, as displayed in Figure S3. The broad peak at 3413 cm−1 stemmed from the O−H stretching of the hydroxyl groups in lignin, whereas the small peak at 1418 cm−1 corresponds to phenolic O−H bending vibration, and the sharp peaks at 1593, 1508, 1458 cm−1 represent aromatic skeletal vibrations.32 Absorption peaks representative of PCL were also observed at 1722, 1238, and 1160 cm−1. These SEM and FTIR results verify the uniform coexistence of lignin with PCL in the fibrous structure. In addition, we compared the wettability of blank PCL and lignin/PCL nanofibers by measuring their water contact angles (Figure S4). Lignin/PCL nanofibers showed a much lower static contact angle (86.27°) than that of blank PCL nanofibers (141.83°) because of the hydrophilicity of the lignin that has many polar groups (e.g., alcoholic and phenolic moieties). This indicated that lignin enhances surface wettability to allow for complete immersion of the film in an aqueous solution. 3.2. Lignin-Induced Mineralization of HAp. We soaked electrospun lignin/PCL nanofiber films in a 1.5 × SBF solution 2686


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Ca2+ ions bound to the surface of the lignin/PCL in the early stage accelerated the later growth of calcium phosphate minerals. Our results suggest that enriched hydroxyl groups on the interface of the lignin-containing nanofibrous film function as chelation sites for Ca2+ ions to facilitate the nucleation of HAp crystals in SBF solutions. This mode is similar to natural bone and tooth formation. Osteoblasts generate the hierarchical nanocomposite structure of bones by secreting a collagenous extracellular matrix enriched in Ca2+ and PO43−, on which HAp crystals subsequently nucleate.33 We further examined the integration of HAp with lignin/ PCL nanofibers using Raman and ATR−FTIR spectroscopies (Figure 3b,c). After incubation for 2 days, new peaks were observed for P−O vibration at 430 cm−1, for O−P−O vibration at 594 cm−1, and for P−O stretching vibration at 961 cm−1 in Raman spectra. In particular, the strongest peak at 961 cm−1 is a representative demonstration of crystalline HAp.14b A shoulder appeared at 1071 cm−1, which is attributable to the presence of CO32−, indicating that trace amounts of PO43− were substituted with CO32− in the HAp in the lattice. Calcium ions cannot mainly deposit as CaCO3 but instead take part in the growth of HAp because of the much lower solubility product constant (Ksp) of HAp (69.6 × 10−126) compared to that of calcite (4.5 × 10−9).34 In the ATR−FTIR spectra obtained after 2 days of HAp growth, PO43− bands observed for the O−P−O vibrations at 562 (ν4), 600 (ν4), 960 (ν1), and 1048 cm−1 (ν3). The CO32− peak at the 870 cm−1 band represents carbonate ions located in PO43− sitesthe most abundant carbonate species in bone.35 3.3. Structure of Lignin-Induced HAp Crystals. We analyzed the structure of the HAp formed on the lignin/PCL nanofibers using TEM, SAED, and XRD. TEM analysis revealed that the HAp agglomerates consisted of clustered, needle-like nanocrystals (Figures 3d and S8). This agglomerate array has a morphology similar to that of native HAp minerals in dental enamel and bone.36 The rod-like residual HAp crystallites were observed in lesions on the enamel’s surface.37 Our analysis using SAED (Figure 3e) confirmed that the platelet-like nanocrystals were HAps. The diffraction pattern of the calcium phosphate clusters that were formed in direct contact with lignin exhibited an arcing of diffraction rings corresponding to the (002) and (004) planes, implying that HAp crystals directionally grew in the c-axis parallel to the nanofibers.38 Furthermore, the XRD results in Figures 3f and S9 reveal a spectral pattern of the (002), (211), (300), (202), (130), (002), (222), and (213) planes, showing that the mineral was HAp in a hexagonal crystal structure, rather than other calcium phosphates such as octacalcium phosphate or whitlockite.39 3.4. Mechanical Properties of Lignin-Induced HAp. The elastic modulus of the cortical bone and cancellous bone ranges from 1 to 20 GPa.40 It is thus imperative to determine the mechanical strength of the HAp precipitated on the surface of lignin/PCL nanofibers and compare it against that of natural bone. We measured the elastic modulus of the HAp layer using the peak force tapping mode of an atomic force microscope. The surface topography and surface roughness changes are shown in Figures 4a and S10. The blank lignin/PCL exhibited a flat, fibrous surface with an average roughness (Ra) of 210.72 ± 9.22 nm. After incubation in SBF for 2 days, protrusions formed on the surface (Ra: 435.72 ± 20.73 nm). We measured the elastic modulus using the Hertz model to analyze the

for mineralization tests. As shown in Figure 2, agglomerates began to nucleate on the lignin/PCL fibrous film after just 2

Figure 2. Formation of HAp minerals facilitated by lignin. (a) SEM images showing the absence and presence of lignin for the formation of HAp after a two-day culturing. (b) SEM image of lignin/PCL nanofibrous film showing that the entire surface of film was covered with HAp after incubation for 5 days. (c) High-magnification SEM image showing a platelet-like structure, which is typically found in natural HAp. (d−f) Results of EDX elemental analysis showing the distribution of Ca and P.

days of incubation (Figure 2a), which evolved into a complete and uniform coating on the film with prolonged incubation for 5 days in the SBF solution (Figure 2b). In contrast, blank PCL nanofibers did not induce the growth of particulates even after incubation for more than 1 week. The high-magnification SEM images in Figure 2c show that the agglomerates exhibited platelet-like clustered protrusions, a typical morphology of HAp crystals. Our analysis using EDX revealed a Ca/P ratio of 1.66 in the HAp agglomerates formed after 5 days of incubation (Figure 2d,e), which is close to the theoretical chemical formula of calcium phosphate as Ca10(PO4)6(OH)2 (Ca/P = 1.67). In the early phase of the continuous mineralization (after 1 day), we observed spherical, ACP particles (Ca/P = 1.32), which subsequently changed the shape to crystalline platelets with an increased Ca/P ratio of 1.64 after 2 days (Figure S5). Note that the nucleation of CaP began when the lignin concentration was higher than 8 mg/mL (Figure S6), which implies that the amount of free hydroxyl groups of lignin is important for CaP nucleation. Thus, we use 10 mg/mL lignin for further experiments. As recorded by our XPS analysis (Figures 3a and S7), initial lignin/PCL nanofibers exhibited C 1s and O 1s peaks in the survey spectrum, but without a high-resolution Ca 2p peak. After incubation in SBF for 12 h, a weak yet detectable Ca 2p peak appeared without any indication of calcium phosphate crystal growth. After further culturing to form calcium phosphate agglomerates, strong Ca 2p and P 2p peaks appeared in the survey spectrum. This indicated that the 2687


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Figure 3. Structural analysis showing that the calcium phosphate cluster formed on lignin-based nanofibers are HAp crystals. (a) Surface XPS analysis indicates the binding of Ca2+ ions to lignin, facilitating HAp nucleation. After incubating the lignin/PCL film in a 1.5× SBF solution for 12 h, a weak Ca 2p peak (without P 2p peak) was observed, which indicates the binding of Ca2+ ions on the nanofibers at an early stage. After further culturing to 24 h, both Ca 2p and P 2p peaks appeared in the spectra. (b) Raman peaks at 430, 594, 961, 1043, and 1071 cm−1 indicate the formation of HAp. (c) ATR−FTIR spectra of HAp@lignin/PCL and lignin/PCL show 562, 600, 870, 960, and 1048 cm−1 peaks for HAp. (d) TEM image of needle-like HAp nanocrystal cluster. (e) SAED and (f) XRD pattern showing the crystalline structure of HAp.

found an elastic modulus of 5.57 ± 0.73 GPa for the lignininduced HAp, which is in a good agreement with the modulus of cancellous human bone.42 It is noteworthy that cancellous bone shows stronger osteogenic and osteoinductive properties than cortical bone, allowing for rapid revascularization of the graft in hard-tissue engineering.43 Taken together, the HAp induced by lignin/PCL nanofiber exhibited structural and mechanical similarities to natural bone. We further tested bulk mechanical strengths of PCL, lignin/ PCL, and HAp@lignin/PCL nanofibers and summarized the result in Table 1. Typical stress−strain curves of the nanofibers are shown in Figure S11. Evidently, the Young’s modulus, defined as the slope of stress−strain curve in the elastic deformation region (5% of strain), increased from 10.23 ± 0.65 MPa for the neat PCL nanofibrous samples to 21.51 ± 0.17 MPa for the lignin/PCL samples. Nevertheless, the tensile strength and elongation at break decreased by the presence of lignin. This is attributed to the inherent chain rigidity and stiffness of lignin, low strength of lignin, and homogeneous distribution (or miscibility) of lignin in PCL.20c In addition, the Young’s Modulus and ultimate tensile strength of lignin/ PCL after mineralization for 2 days (HAp@lignin/PCL-2d) and 5 days (HAp@lignin/PCL-5d) were significantly higher

load−displacement force/distance (F/d) curves according to the following equation F d

3/2

=

4 E × R1/2 3

where F is the applied force on the cantilever, R is the radius of the cantilever probe, d is the indentation depth of the sample, and E is Young’s modulus of a sample. As shown in Figure 4b,c, a relatively flat area (region 1) was near zero indentation, where the contact area was negligible, so contact between the spherical indenter and sample was minimal. Under this condition, the probe-interaction forces and friction were nonnegligible due to the weak contact force.41 With more force to the probe, the contact region increased elastically (region 2), and the local spherical property of the probe became less important.41 We calculated the elastic modulus by estimating the slope of this region. Region 3 displayed a negative indentation depth, which was caused by plastic deformation occurring under the probe, suggesting that it is unfit for calculating elastic modulus. The fitted F/d curve of region 2 was linear, as shown in Figure 4d, corresponding to the Hertz model. Based on the average R value of 100 nm provided by the probe manufacturer (Nanosensors Inc., Switzerland), we 2688


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Figure 4. AFM analysis for HAp formed on the lignin-based substrate. (a) 3D topography of the lignin/PCL film before and after incubation in a SBF for 2 days. (b) Schematic illustration of the different conditions of the probe under varied applied force. (c) Applied force vs indent depth curve for the sample. (d) Data from region 2 of (c), plotted as force vs indent3/2, used to calculate the elastic modulus from the slope of a linear fit to the data.

indicated that the HAp formed by mineralization was favorable for the cell proliferation. We further analyzed cell viability and morphology changes using the LIVE/DEAD assay method. According to our observations, cells attached to all of the scaffolds after 2 days of cultivation, with minimal amounts of dead cells (Figure 5a), and the number of live cells was approximately 145−200 per 106 μm2 area, with no significant differences between the samples (Figure 5b). However, the cell morphology was highly affected by the scaffold type. While the cells spread well and displayed polygonal morphology on the blank PCL and lignin/PCL nanofibers, narrow and elongated spindle-like shapes were observed on the HAp@lignin/PCL nanofibers, where cells wrapped around the HAp microspheres and interacted through filamentous extensions, as shown in the SEM images (Figure S13). To quantify the differences in cell morphology, we estimated the projected cell area and the cell shape index (CSI) on each scaffold material. Note that the CSI represents the circularity of a cell with a value ranging between 0 (linear) and 1 (circle).45 On the blank PCL and lignin/PCL films, the projected cell areas were approx 1850−2000 μm2, but the area decreased remarkably to approx 1000 μm2 after mineralization with HAp (Figure 5c). The CSI values were approx 0.60 and 0.68 when osteoblastic cells were cultured on blank PCL and lignin/PCL nanofibers, respectively. This result suggests that the surface became more hydrophilic with the addition of lignin, enhancing cell adhesion and spreading.46 In contrast, the CSI value decreased significantly to 0.39 on the HAp@ lignin/PCL nanofiber film, which revealed a more elongated cell morphology (Figure 5d). Cells show an elongated morphology because of larger expenditure of energy for metamorphosing the membrane on a rough surface.47 The HAp microspheres formed on the lignin/PCL nanofibers

Table 1. Tensile Results for PCL, Lignin/PCL, and HAp@ Lignin/PCL Fibers samples PCL lignin/PCL HAp@lignin/PCL-2d HAp@lignin/PCL-7d

Young’s module (MPa) 10.23 21.51 33.47 55.56

± ± ± ±

0.65 0.17 0.72 0.77

ultimate tensile strength (MPa) 9.82 8.73 12.41 16.56

± ± ± ±

0.15 0.21 0.13 0.34

elongation at break (%) 42 37 35 33

± ± ± ±

7 5 3 9

than those of their counterpart nanofibers without HAp, suggesting that mechanical properties of the scaffolds could be effectively improved by mineral coatings. 3.5. Biocompatibility and Osteoconductivity of HAp Grown on Lignin/PCL Nanofiber. We investigated the biocompatibility of the bone-like HAp by culturing mouse preosteoblasts (MC3T3-E1) on the HAp-covered scaffolds. Osteoblasts secrete type I collagen, noncollagenous proteins, enzymes, and the growth factors of the bone matrix and trigger the deposition of calcium salts in bone tissue.44 Thus, the interaction between osteoblasts and supporting scaffolds should be evaluated before in vivo applications. We assessed the cell adhesion and proliferation activities of mouse osteoblastic cells (MC3T3-E1) on three types of scaffolds (blank PCL, lignin/PCL, and HAp@lignin/PCL nanofiber films) using the Cell Counting Kit-8 (CCK-8) to measure the optical density (OD) value at the wavelength of 490 nm (Figure S12). The OD values of the cells cultured on all scaffolds increased slowly with no significant difference at the first day, suggesting that cells were in the lag phase. After culturing for 5 and 7 days, cell colonization and proliferation on the HAp@lignin/PCL increased significantly, as compared with the PCL and lignin/PCL scaffolds (p < 0.05), which 2689


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Figure 5. In vitro biocompatibility of blank PCL, lignin/PCL, and HAp@lignin/PCL films. (a) LIVE/DEAD cell assay results in the MC3T3-E1 cells culture after 2 days on each film, (b) number of live cells per 106 μm2, (c) projected cell area, and (d) CSI were analyzed from the fluorescent images. Results are presented as mean ± SEM, and statistical analysis was carried out by means of one-way ANOVA (n ≥ 3). **p < 0.01, n.s. not significant. (e) Cytoskeleton organization of MC3T3-E1 cells grown on each film.

4. CONCLUSION Lignin-based nanofibers can mineralize bioactive inorganic crystals (i.e., bone-like HAp), create osteoblastic environments similar to hard tissues, and promote excellent osteoconductivity. Abundant hydroxyl groups on the side chains of lignin induce the nucleation and growth of HAp crystals in the c-axis through co-precipitation of Ca2+ and PO43− ions, similar to the natural process of bone and tooth formation. The HAp agglomerates grown on the lignin-based matrix consisted of needle-like nanocrystals, which exhibited a structural morphology and alignment close to those in natural HAp. We characterized the HAp structure using multiple analytical tools, including SEM, EDX, Raman, FTIR, XRD, and so forth. The elastic modulus of HAp reached 5.57 ± 0.73 GPa, which is in good accordance with the modulus of cancellous human bone. In addition, the Young’s Modulus and ultimate tensile strength of lignin/PCL after mineralization for 2 and 5 days were significantly higher than those of their counterpart nanofibers without HAp, suggesting that bulk mechanical properties of the scaffolds could be effectively improved by mineral coatings. Furthermore, our cytotoxicity assays indicated the suitability of HAp-mineralized lignin-based scaffolds for enhanced viability and proliferation of osteoblastic cells, which exhibited elongated morphology, with filopodia spread out and extended to wrap around HAp microspheres. This work shows that naturally abundant lignin possesses a strong capability to mediate biomimetic deposition of HAp crystals. Considering the lignin’s inherent properties, such as its antioxidant and antibacterial activities as well as good biocompatibility, the

should induce a narrower morphology for the adhering cells. The cytoskeletons of the MC3T3-E1 cells were well organized, with sheet-like lamellipodiathe branched network of actin on blank PCL and lignin/PCL film after 2 days of culture (Figures 5e and S14). In contrast, elongated filopodia that are contractile bundles of filamentous-actin, protruded from the actin network when cells were grown on the HAp@lignin/PCL film. Importantly, the formation of the filopodia is responsible for cell migration and many cellular processes, including wound healing, adhesion to the extracellular matrix, and guidance toward chemoattractants.48 Moreover, cell adhesion in a 3D matrix via abundant filopodia extension is more biologically relevant to their native microenvironment (or in vivo), shadowing the lamellipodia formation which is generated on the surface of tissue implants.49 Although nanofibrous scaffolds are considered to be a 3D matrix, the short distances between the fibers limit the cell penetration into the electrospun network, thus inducing cell growth with the predominant formation of lamellipodia on the top surface.50 According to the literature, a number of filopodia would spread out and extend to protuberances when the surface had blade-like nanotips with a height up to 230 nm.51 In this regard, HAp crystals, which have a hundreds of nanometer-scale height, can play the role of filopodia binding site. Taken together, our results suggest that HAp@lignin/PCL provides excellent osteoblast adhesion in the matrix of 3D nanotopographies with good cell viability and osteoconductivity, resulting in an integrate of osteoblastic environments which is similar to that in natural tissues. 2690


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lignin−biomineral hybrid platform presents a new prospect for lignin-based materials toward bone-related therapies.

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ASSOCIATED CONTENT

* Supporting Information S

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.biomac.9b00451. SEM images of electrospun products of pure lignin solutions; SEM and digital images of electrospun nanofibers with different lignin concentration and average diameters; ATR−FTIR spectra of lignin, PCL, and lignin/PCL; water contact angle test for lignin/PCL and PCL; schematic process for lignin-induced HAp crystal formation on lignin/PCL film and SEM images with EDX spectra; effect of lignin concentration on the formation of HAp’s; additional surface XPS analysis for lignin/PCL film before and after incubation in a 1.5× SBF solution; additional TEM micrographs of HAp nanocrystalline; documented XRD patterns; topographies of AFM images; stress−strain curves for PCL, lignin/PCL, and HAp@lignin/PCL; CCK-8 assay, SEM images, and additional fluorescent microscopy images of MC3T3-E1 cells grown on PCL, lignin/PCL, and HAp@lignin/PCL films (PDF)

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Chan Beum Park: 0000-0002-0767-8629 Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS This study was supported by the National Research Foundation (NRF) via the Creative Research Initiative Center (NRF-2015R1A3A2066191), Republic of Korea.

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REFERENCES

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