Sustainable and Antioxidant Lignin–Polyester ... - ACS Publications

Publication Date (Web): May 26, 2017 ... certifying the great potential of lignin–PCLLA copolymers and nanofibers for biomedical or healthcare appli...
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Research Article pubs.acs.org/journal/ascecg

Sustainable and Antioxidant Lignin−Polyester Copolymers and Nanofibers for Potential Healthcare Applications Dan Kai,*,† Kangyi Zhang,† Lu Jiang,† Hua Zhong Wong,‡ Zibiao Li,† Zheng Zhang,† and Xian Jun Loh*,†,§,∥ †

Institute of Materials Research and Engineering (IMRE), A*STAR, 2 Fusionopolis Way, #08-03 Innovis, Singapore 138634 Department of Chemistry, National University of Singapore, 3 Science Drive 3, Singpaore 117543 § Department of Materials Science and Engineering, National University of Singapore, 9 Engineering Drive 1, Singapore 117576 ∥ Singapore Eye Research Institute, 11 Third Hospital Avenue, Singapore 168751 ‡

S Supporting Information *

ABSTRACT: Lignin polymerization has been considered as an effective approach for lignin valorization. Herein we report the synthesis of a series of new lignin-based copolymers (lignin−poly(ε-caprolactone-co-lactide), lignin−PCLLA) via solvent-free ring-opening polymerization. Lignin−PCLLA copolymers with tunable molecular weights (10 to 16 kDa) and glass transition temperatures (−40 to 40 °C) were obtained. Such copolymers were engineered into ultrafine nanofibers by blending with polyesters (polycaprolactone, PCL and poly(L-lactic acid), PLLA) via electrospinning. Both PCL/lignin−PCLLA and PLLA/ lignin−PCLLA nanofibers displayed uniform and beadless nanofibrous morphology. The size (diameters ranging from 300 to 500 nm) and tensile tests of the obtained nanofibers indicated that the lignin copolymers are miscible with the polyester matrices and can significantly improve the mechanical properties of the nanofibers. Moreover, good antioxidant activity and biocompatibility of the lignin nanofibers were demonstrated in vitro, certifying the great potential of lignin−PCLLA copolymers and nanofibers for biomedical or healthcare applications. KEYWORDS: Lignin, Ring-opening polymerization, PCL, PLLA, Electrospinning

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polymers.11,12 Grafting polymers onto lignin is a common chemical modification method that could enhance its chemical identity (or similarity) to the polymer matrices. The active hydroxyl groups on lignin are good binding sites for grafting polymerization. Many polymerization methods have been developed to graft polymers onto lignin, including atom transfer radical polymerization (ATRP), polyurethane synthesis, ring-opening polymerization (ROP), and reversible addition−fragmentation chain transfer (RAFT).13−17 Various polymers, such as poly(lactic acid) (PLA), poly(methyl methacrylate) (PMMA), polystyrene, and poly(ethylene glycol) (PEG), have been successfully grafted onto lignin.8,18−23

INTRODUCTION Lignin is one of the most abundant biopolymers on earth, and it exhibits many attractive properties, including low density, high carbon content, good stiffness, biodegradability, and antioxidant activity.1−5 However, lignin is always treated as waste from the paper industry and only used in low-value applications. In light of the incessant consumption of fossilbased resources today, the conversion of renewable lignin into high-value products would be a major sustainability milestone. Lignin is considered as a potential biopolymeric filler to improve the mechanical properties of composite materials.1,3,6−8 Unfortunately, unmodified lignin shows poor miscibility in many thermoplastic polymers, resulting in an unsatisfactory impact on strength and elongation.9,10 Therefore, chemical modification of the lignin surface is crucial to improve its compatibility with other thermoplastic © 2017 American Chemical Society

Received: March 20, 2017 Revised: May 15, 2017 Published: May 26, 2017 6016

DOI: 10.1021/acssuschemeng.7b00850 ACS Sustainable Chem. Eng. 2017, 5, 6016−6025

Research Article

ACS Sustainable Chemistry & Engineering

Figure 1. (top) Ring-opening polymerization of ε-caprolactone and L-lactide on lignin using tin(II) 2-ethylhexanoate as the catalyst. (bottom) 1H NMR spectrum of lignin−PCLLA copolymer (LP2) in CDCl3.

However, the poor solubility of lignin in commonly used organic solvent limits the approaches of lignin modification. In this study, we developed a new solvent-free polymerization method for grafting polymers onto lignin. Currently, numerous studies have demonstrated that ligninbased copolymers are able to contribute positively to the mechanical reinforcement in bulk composites.7,24 However, there are few studies on lignin-based porous materials. Compared with bulk materials, lignin materials with porous structures might be more interesting, as they exhibit many

advantages, such as low density, high surface area, high absorbability, and high permeability. With such advantages, lignin-based porous composites can potentially be used in various high-value applications, including catalysts, energy storage, sensors, and biomedical materials. In this study, we synthesized a series of lignin−poly(εcaprolactone-co-lactide) (lignin−PCLLA) copolymers via solvent-free ROP of ε-caprolactone/L-lactide mixtures with different ratios. Moreover, such lignin copolymers were blended with poly(ε-caprolactone) (PCL) or poly(L-lactic acid) (PLLA) 6017

DOI: 10.1021/acssuschemeng.7b00850 ACS Sustainable Chem. Eng. 2017, 5, 6016−6025

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ACS Sustainable Chemistry & Engineering Table 1. Molecular and Thermal Characterization of Lignin−PCLLA Copolymers polymer

lignin:CL:LA mass feed ratio

unreacted lignin (%)

yield (%)

Mw (kDa)a

PDIa

PCL:PLLA chain length ratio in copolymerb

% lignin in copolymerc

Tg (°C)d

LP1 LP2 LP3

2:2.4:5.6 2:4:4 2:5.6:2.4

0.4 1.2 1.8

68 51 59

10.4 15.0 16.4

1.30 1.25 2.41

0.5:1 1.4:1 3.5:1

48 33 30

34 20 −37

a Determined by GPC. bDetermined by NMR spectroscopy. PCL = poly(ε-caprolactone); PLA = poly(L-lactide). cDetermined by GPC on the basis of the molecular weight of lignin (5 kDa). dGlass transition temperature, determined by DSC.

−70 °C at 20 °C/min, and finally reheating from −70 to +150 °C at 20 °C/min. Electrospinning. Mixtures of PCL and lignin copolymers were dissolved in 1,1,1,3,3,3-hexafluoro-2-propanol (HFP). The mass ratio of PCL and each lignin−PCLLA copolymer was 90:10. The total concentration of the solution was 10% (w/v). After 1 day of stirring, the homogeneous solution was loaded into a 5 mL syringe with a blunt 22-gauge needle (inner diameter of 0.413 mm). The solution was pumped out at 0.8 mL/h with 12 kV applied to the needle. Fibers were electrospun onto an aluminum-foil-wrapped collector (20 cm away from the needle tip). After spinning, the obtained fibers were dried overnight in a vacuum oven and used for characterization and cell culture assays. PLLA/lignin−PCLLA nanofibers were fabricated using similar parameters except that the total concentration of the solution was 8% (w/v). Characterization of Electrospun Nanofibers. The surface topography of the electrospun fibers was characterized by scanning electron microscopy (SEM) on a JEOL JSM6700F scanning electron microscope. Samples were sputter-coated with a thin layer of gold before imaging. Fiber diameters were determined from 50 random measurements per image using ImageJ (National Institutes of Health, Bethesda, MD, USA). The hydrophilicities of the electrospun fibers were evaluated by water contact angle measurements using a VCA Optima Surface Analysis System (AST Products, Billerica, MA, USA). For each sample, water droplets (0.5 μL) were placed on the surface of nanofibers at room temperature. At least five independent measurements were taken for each sample. The mechanical properties of the nanofibers were determined using a uniaxial tensile testing technique (model 5943 testing system, Instron, Norwood, MA, USA) with a 10 N load capacity at a rate of 10 mm/min. Nanofibers were cut into a rectangular shape with dimensions of 5 mm × 30 mm for testing; the thickness of the samples was about 50 μm, and the gauge length was 20 mm. At least five samples were prepared and tested for each composition. Tensile strength, Young’s modulus, and elongation at break were calculated from the stress−strain curve of each sample. The thermal behavior of the composite fibers was also investigated by DSC. The following protocol was used: heating from −70 to +200 °C at 20 °C/min, holding at +200 °C for 5 min, cooling from +200 to −70 °C at 20 °C/min, and finally reheating from −70 to +200 °C at 20 °C/min. The cold crystallization temperature (Tcc) and cold crystallization enthalpy change (ΔHcc, negative) of PLLA were determined from the exothermic cold crystallization peak, while the melting temperature (Tm) and melting enthalpy change (ΔHm, positive) were determined from the endothermic melting peak (second heating run). The crystallinity (Xc) was calculated using the following equation:

and then electrospun into porous composite fibers. PCL and PLLA are polyester aliphatic polymers.25−29 Both polymers are mechanically strong, easy to process, biodegradable, and biocompatible in the human body. They have been considered as suitable biomaterials for multiple biomedical applications, including sutures, implants, tissue engineering scaffolds, and drug release systems.27,30−35 Electrospinning is a simple and effective technology to engineer three-dimensional porous materials in the form of micro/nanofibers.31,32,34,36−40 Electrospun nanofibers have been used in many high-value applications, such as filtration, cosmetic masks, sensors, and biomaterial scaffolds. Our aim is to investigate the effect of such lignin copolymers on the material properties (morphology and thermal and mechanical properties) of the composite nanofibers. Furthermore, the antioxidant activity of the nanofibers and their biocompatibility were also evaluated in vitro for potential biomedical or healthcare applications.

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MATERIALS AND METHODS

Materials. All of the chemicals were purchased from Sigma-Aldrich Chemicals and used as received except where noted. Alkali lignin (average molecular weight = 5 kDa) was dried at 105 °C overnight before use. PLLA (REVODE101 grade, melting flow index 2−10) was purchased from Zhejiang Hisun Biomaterials Co. Ltd. ε-Caprolactone was purified by distillation under reduced pressure before use. Synthesis of Lignin−PCLLA Copolymers. Lignin−PCLLA was synthesized by solvent-free ROP (Figure 1). Alkali lignin, εcaprolactone, L-lactide, and tin(II) 2-ethylhexanoate (0.5 wt % of the monomer as a catalyst) were weighed into a round-bottom flask. The feed weight ratios of lignin and monomers are shown in Table 1. The mixture was stirred at 130 °C for 24 h under an atmosphere of N2. After the mixture was cooled to room temperature, 100 mL of chloroform was added. Chloroform would dissolve the synthesized lignin−PCLLA copolymers but not the unreacted lignin. After a further 4 h of stirring, the solution was centrifuged, and the supernatant was poured into ether while the solid left in the bottom was dried and collected as unreacted lignin. The obtained precipitate from the supernatant was further purified by washing with ether/ methanol mixed solvent. The final product was then dried in a vacuum oven at 50 °C for 24 h to yield the lignin−PCLLA copolymer as a dark-brown solid. Characterization of Lignin−PCLLA Copolymers. NMR spectra of lignin−PCLLA copolymers were measured at room temperature using Bruker AV-400 and JEOL 500 MHz NMR spectrometers in CDCl 3 solvent. The molecular weights (M n and M w ) and polydispersity index of polymer samples were analyzed by gelpermeation chromatography (GPC) on a Shimadzu SCL-10A and LC8A system equipped with two Phenogel 5 mm, 50 and 1000 Å columns in series and a Shimadzu RID-10A refractive index detector using HPLC-grade tetrahydrofuran as the eluent at a flow rate of 1.0 mL/min at 25 °C. The average molecular weights were determined with a calibration based on linear PMMA standards. The thermal behaviors of the copolymers were also investigated by differential scanning calorimetry (DSC) on a TA Instruments Q100 calorimeter equipped with an autocool accessory and calibrated using indium. The following protocol was used for testing: heating from −70 to +150 °C at 20 °C/min, holding at +150 °C for 5 min, cooling from +150 to

Xc =

ΔHm × 100% ΔHm0

(1)

where ΔHm is normalized based on the mass percentage of PCL or PLLA segments in the fibers and ΔH0m is 139.5 J/g for 100% crystalline PCL41 or 93.7 J/g for 100% crystalline PLLA.42 The antioxidant activity of the nanofibers was evaluated using the 1,1-diphenyl-2-picrylhydrazyl (DPPH) assay.43,44 Fibers (10 mg) were placed in glass vials. A 60 μM DPPH solution in MeOH was prepared, and 10 mL of this solution was added to each vial. The DPPH free radical content was measured by monitoring the absorbance changes at 6018

DOI: 10.1021/acssuschemeng.7b00850 ACS Sustainable Chem. Eng. 2017, 5, 6016−6025

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Figure 2. SEM images of electrospun (A) PCL, (B) PCL/LP1, (C) PCL/LP2, (D) PCL/LP3, (E) PLLA, (F) PLLA/LP1, (G) PLLA/LP2, and (H) PLLA/LP3 nanofibers. Scale bars = 1 μm. 517 nm at each time point. All of the samples were prepared and tested in triplicate. The antiradical activity is represented as the percent inhibition of free radicals by measuring the decrease in absorbance compared with control solutions. The degradation behavior of the electrospun nanofibers was determined by checking their morphological change in phosphatebuffered saline (PBS). The scaffolds were placed in a 24-well plate containing 1.5 mL of PBS (pH 10) in each well and incubated in vitro at 37 °C for 4 weeks. PBS (pH 10) was used for the accelerated degradation. After the degradation period, samples were washed and subsequently dried in a vacuum oven at room temperature for 24 h. In addition, the morphology of the scaffolds was evaluated by SEM. Cell Viability Study. NIH/3T3 (ATCC) fibroblasts (ATCC CRL1658) were cultivated according to ATCC protocols. Cells in culture flasks were seeded into 24-well microplates at a density of 5000 cells per well. On days 2, 4 and 6, the Promega CellTiter-Blue (resazurin) cell viability assay was conducted according to the manufacturer’s protocol. Readings were normalized to cells grown on tissue culture plastic. The samples were then washed with PBS, and the cell culture was continued for the remaining time points. Higher fluorescence readings indicate higher cell viability and biocompatibility. After 6 days of cell cultivation, nanofiber samples were fixed with 4% paraformaldehyde at room temperature. Cells were then permeabilized in 0.1% Triton X-100 before staining in 4′,6diamidino-2-phenylindole (DAPI)−phalloidin solution. Fluorescently labeled phalloidin (Mo Bi Tech) was used to stain F-actin, and DAPI (Sigma) was used for nuclear staining. This was followed by mounting in ProLong Gold antifade reagent (Thermo Fisher Scientific) and visualization using a fluorescence microscope (Leica DMi8 inverted microscope). Statistical Analysis. All of the data presented are expressed as mean ± standard deviation of the mean. Student’s t test was used, and differences between the groups were considered statistically significant at p < 0.05.

lignin were observed at 3.85 ppm, corresponding to methoxy protons, and 6.85 ppm, representing the protons of the phenyl rings. Strong signals were detected at 2.30, 1.64, 1.38, and 4.06 ppm, corresponding to the methylene protons in the PCL chains (Figure 1 and Figure S1 in the Supporting Information). The characteristic peaks of PLLA were observed at 5.12 and 1.50 ppm. Moreover, the chemical compositions of lignin− PCLLA copolymers were studied by X-ray photoelectron spectroscopy (XPS) (Figure S2). In the lignin samples, C is mainly present as C−C/C−H (284.6 eV) and C−O (286.6 eV) bonds. Additionally, the lignin copolymers present the O−C O peak at 292.2 eV, indicating that lignin is grafted with PCL and PLLA. The molecular weights of the lignin−PCLLA copolymers were determined by GPC. As shown in Table 1, LP1, LP2, and LP3 showed weight-average molecular weights (Mw) of 10.4, 15.0, and 16.4 kDa, respectively. On the basis of the molecular weight of lignin (5 kDa), the mass percentages of lignin in these copolymers were 48% for LP1, 33% for LP2, and 30% for LP3. The PCL:PLLA segment length ratios in the copolymers were 0.5:1 for LP1, 1.4:1 for LP2, and 3.5:1 for LP3 (calculated using 1 H NMR spectroscopy by comparing the areas of peaks a and f in Figure 1). The thermal properties of the lignin−PCLLA copolymers were determined by DSC. The DSC results (Figure S3 and Table 1) showed that all three copolymers were amorphous polymers. The glass transition temperature (Tg) of the copolymers decreased with increasing PCL chain length, from 34 °C for LP1 to −37 °C for LP3. All of the above results confirmed that both PCL and PLLA chains were successfully grafted onto lignin. Electrospinning of Nanofibers. Lignin copolymers were blended with PCL or PLLA at a ratio of 90:10 (w/w) to prepare electrospun nanofibers. Electrospun nanofibers with uniform, beadless, randomly oriented structures were obtained under the optimized spinning conditions. This indicates uniform dispersion of lignin copolymers in the polyester matrixes at the nanoscale level. As shown in Figure 2, neat PCL nanofibers were obtained in 10% solution with a fiber diameter of 354 ± 96 nm. The addition of lignin copolymers increased the diameter of the resulting nanofibers slightly. The fiber diameters of PCL/LP1, PCL/PL2, and PCL/PL3 were 458 ± 84, 436 ± 81, and 378 ± 68 nm, respectively. On the other hand, neat PLLA nanofibers were prepared in 8% solution with

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RESULTS AND DISCUSSION Synthesis and Characterization of Lignin-PCLLA Copolymers. Lignin is not soluble in common organic solvents, and chemical pretreatment is necessary before surface modification or grafting. In this work, instead of using any organic solvent, we used liquid ε-caprolactone as the medium to disperse lignin and L-lactide for polymerization. The synthesized lignin copolymers were able to dissolve well in many common organic solvents, such as tetrahydrofuran and chloroform. Such copolymers were characterized by 1H NMR spectroscopy in CDCl3. The characteristic chemical shifts of 6019

DOI: 10.1021/acssuschemeng.7b00850 ACS Sustainable Chem. Eng. 2017, 5, 6016−6025

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ACS Sustainable Chemistry & Engineering Table 2. Material Characterization of Electrospun Nanofibers fiber

copolymer blendeda

PCL PCL/LP1 PCL/LP2 PCL/LP3 PLLA PLLA/LP1 PLLA/LP2 PLLA/LP3

NAd LP1 LP2 LP3 NA LP1 LP2 LP3

fiber diameter (nm) 354 458 436 378 255 384 485 558

± ± ± ± ± ± ± ±

96 84 81 68 67 73 120 136

water contact angle (deg) 132 137 138 138 134 143 141 140

± ± ± ± ± ± ± ±

5 1 1 3 2 2 1 2

Tcc Tg (°C)b (°C)b − − − − 61 57 61 61

NA NA NA NA 96 115 123 96

ΔHcc (J/g)b

Tm (°C)b

ΔHm (J/g)b

Xc (%)c

NA NA NA NA 2.0 99.7 28.5 2.5

57 57 62 57 174 173 174 174

50.3 59.8 52.9 56.5 49.5 41.9 36.4 49.5

36.1 47.6 42.1 45.0 53.2 50.1 43.5 59.1

tensile strength (MPa) 2.9 10.7 6.8 4.9 3.0 2.3 3.1 2.4

± ± ± ± ± ± ± ±

0.4 0.9e 1.2e 1.5e 0.3 0.3e 0.6 0.3e

Young’s modulus (MPa) 6.1 25.9 19.6 12.8 68.9 73.9 67.9 65.1

± ± ± ± ± ± ± ±

1.0 3.4e 3.6e 2.3e 11.1 5.1 8.8 5.4

elongation at break (%) 112 143 122 119 67 30 70 39

± ± ± ± ± ± ± ±

6 14e 12 24 9 8e 11 2e

a

The ratio of PCL or PLLA to lignin copolymer was 9:1. bThe glass transition temperature (Tg), cold crystallization temperature (Tcc), cold crystallization enthalpy change (ΔHcc), melting temperature (Tm), and melting enthalpy change (ΔHm) were determined by DSC. cThe crystallinity was determined using the equation Xc = (ΔHm/ΔH0m) × 100%, where ΔH0m is 139.5 J/g for 100% crystalline PCL or 93.7 J/g for 100% crystalline PLA. dNA: not applicable. eSignificantly different from the corresponding parameter for the PCL or PLLA fiber (p < 0.05).

Figure 3. Typical stress−strain curves of (A) PCL/lignin−PCLLA and (B) PLLA/lignin−PCLLA nanofibers by tensile test.

a fiber diameter of 255 ± 67 nm. The addition of lignin copolymers into PLLA nanofibers increased the fiber diameter from 384 ± 73 nm for PLLA/LP1 to 558 ± 136 nm for PLLA/ LP3. The hydrophilicity of a biomaterial is a crucial parameter, as it affects cell adhesion and growth on its surface. The surface hydrophilicities of the electrospun nanofibers were characterized by water contact angle measurements, and the results are summarized in Table 2. Both the PCL and PLLA fibers were hydrophobic (132 ± 5° for PCL and 134 ± 2° for PLLA), and the addition of lignin copolymers did not significantly affect the water contact angles of the nanofibers. Lignin shows a low water contact angle, indicating that the surface is hydrophilic, but grafting hydrophobic PCLLA chains onto its surface increased the surface hydrophobicity. Moreover, we added only 5% lignin copolymers into the nanofibers, which could not change their surface wettability remarkably. Mechanical Properties of Electrospun Nanofibers. It has been reported that the mechanical properties of the extracelluar matrix can regulate the cell behavior, including attachment, proliferation, migration, and even differentiation.45,46 A biomaterial with suitable mechanical properties may also provide mechanical cues to guide cell growth. A stiff tissue engineering scaffold might benefit bone regeneration, while a soft biomaterial might favor the growth of skin or muscle cells. Therefore, it was crucial to investigate the mechanical properties of these lignin nanofibers. The mechanical properties of the electrospun nanofibers were evaluated by tensile tests. As shown in Table 2 and Figure 3,

PCL is soft and elastic. The neat PCL nanofibers showed the lowest mechanical properties, with a tensile strength of 2.9 ± 0.4 MPa, Young’s modulus of 6.1 ± 1.0 MPa, and elongation at break of 112 ± 6 MPa. The addition of lignin−PCLLA copolymers into PCL significantly improved the mechanical properties of the resulting nanofibers. Among all of the PCL nanofibers, PCL/LP1 displayed the best mechanical properties (tensile strength of 10.7 ± 0.9 MPa, Young’s modulus of 25.9 ± 3.4 MPa, and elongation at break of 143 ± 14 MPa). The results indicate that lignin−PCLLA copolymers serve as good fillers for the mechanical reinforcement of PCL nanofibers and that the copolymers with higher lignin content made a greater contribution to the higher mechanical properties. On the other hand, the addition of lignin copolymers into PLLA failed to improve the mechanical properties of the resulting nanofibers. All of the PLLA nanofibers show similar Young’s modulus (∼70 MPa). Neat PLLA and PLLA/LP2 display similar tensile strength (∼3 MPa) and elongation at break (∼70%). Addition of LP1 and LP2 decreased the strength and elongation of the PLLA nanofibers, and hence, lignin−PCLLA copolymers cannot reinforce PLLA nanofibers. The native function of lignin is to provide mechanical support for plants. Therefore, lignin has been considered as a potential structural filler for mechanical reinforcement in polymer matrixes. Studies have demonstrated that grafting polymers onto lignin could improve its compatibility with a polymer matrix and further enhance mechanical properties. Lignin−PLA and lignin−PCL−PDLA copolymers have been shown to reinforce the tensile properties of PLA films.18,19,47 6020

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Figure 4. DSC curves (second heating run) of (A) PCL/lignin−PCLLA and (B) PLLA/lignin−PCLLA nanofibers.

Figure 5. Free radical inhibition (antioxidant activity) of (A) PCL/lignin−PCLLA and (B) PLLA/lignin−PCLLA by DPPH assay.

Compared with bulk materials or films, a greater obstacle lies in reinforcing porous materials, especially nanofibers. The use of lignin will be a greater challenge in this respect. It is crucial and also challenging to ensure good affinity between the lignin copolymer and the polymer matrix. A filler with good affinity is able to disperse well in polymer matrix and form strong interface binding, while poor affinity may cause filler aggregation and even phase separation. In our study, lignin− PCLLA copolymers significantly enhanced the PCL nanofibers but caused negative effects on the PLLA nanofibers. As shown in Table 1, the PCL segments in the copolymers are longer than the PLLA segments (except LP1). This might result in better affinity of the copolymers for the PCL matrix than the PLLA matrix. Thermal Properties of Electrospun Nanofibers. DSC was used to determine the glass transition temperatures (Tg) and crystallization properties of the electrospun nanofibers. Figure 4 shows the DSC thermograms of the electrospun PCL/ lignin−PCLLA and PLLA/lignin−PCLLA nanofibers. Numerical values corresponding to the thermal transitions are given in Table 2. As presented in Figure 4, all of the obtained PLLA/ lignin−PCLLA nanofibers exhibit a single Tg with a distinct melting endotherm of PLLA. The presence of a single Tg indicates good miscibility between the lignin−PCLLA fillers and the PLLA matrix in the electrospun nanofibers.48,49 The Tg values of the PLLA/lignin−PCLLA nanofibers are similar to that of PLLA (Table 2), thus demonstrating that the presence of lignin−PCLLA did not have a dramatic effect on the PLLA chain mobility during heating. This finding is also supported by

the relatively unaffected Tm of PLLA in the PLLA/lignin− PCLLA nanofibers (Figure 4). The insignificant alteration of the Tg and Tm values of PLLA/lignin−PCLLA nanofibers is consistent with the previous report on the PLLA/carbon nanotube (CNT) composite system. The presence of the CNT wall did not promote enough interactions at the interface that could induce a change in the behavior during thermal transitions.50 With respect to the crystallization behavior of PLLA/lignin−PCLLA nanofibers, the DSC results do not reveal a consistent trend. However, since the materials do not display a sharp Tcc transition point, subtle changes in the PLLA and lignin−PCLLA interactions may remain undetected. This is considering the broadness of the cold crystallization peaks.51−53 Similar thermal transition behavior was also observed for PCL/ lignin−PCLLA nanofibers. Antioxidant Activity of Electrospun Nanofibers. The antioxidant activities of lignin−PCLLA copolymers and their nanofibers were evaluated by the DPPH assay. The lignin copolymers maintained good antioxidant activity, and those with higher lignin content showed higher antioxidant activity (Figure S4). At 1 h, LP1, LP2, and LP3 exhibited free radical inhibitions of 86.3 ± 2.3%, 77.7 ± 4.3%, and 29.1 ± 4.8% respectively. After 8 h of incubation, the inhibition values of all three copolymers were above 75%. The high content of phenolic moieties in lignin contributes to its good antioxidant activity. Our results indicate that ring-opening polymerization grafts PCL and PLLA chains onto only some of the phenol groups of lignin. The remaining phenols groups can still be functionalized for radical termination. 6021

DOI: 10.1021/acssuschemeng.7b00850 ACS Sustainable Chem. Eng. 2017, 5, 6016−6025

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Figure 6. Cell viability results for (A) day 2, (B) day 4, (C) day 6, and (D) all days. Readings are normalized to tissue culture plastic. In (B) and (C), the line annotations are a selected subset of all significant means comparison after one-way ANOVA (N = 6; solid line, p ≤ 0.05; dotted line, p ≤ 0.01; dashed line, p ≤ 0.001).

acrylate, lignin−PMMA, and lignin−PLA) by using different cell types.38,51,56 Here we cultured NIH/3T3 fibroblasts on PLLA/ and PCL/lignin−PCLLA nanofibers to evaluate their biocompatibility. As shown in Figure 6, while there was no significant difference in cell viability between samples after 2 days of cell cultivation, interesting trends could be observed at day 4. Within both the PLLA and PCL materials, introducing lignin at the LP3 concentration caused the best improvement in cell proliferation (Figure 6B). Significant differences can be observed when PLLA is compared with PLLA/LP2 and PLLA/ LP3. These differences do not continue until day 6 because the group of PLLA scaffolds would have attained maximum cell density (Figure 6C). In general, the PLLA group of scaffolds demonstrated superior cell viability compared with the PCL group at days 4 and 6 (Figure 6B,C). Moreover, the morphology of NIH/3T3 fibroblasts was examined by F-actin staining. F-actin is an important protein to form microfilaments in cells, and it also participates in many cellular functions, including cell attachment, growth, division, and signaling. When visualized under a fluorescence microscope (Figure 7), the NIH/3T3 fibroblasts showed the desirable elongated, spindlelike morphology for all nanofiber types. The cells were well spread out and oriented along fibers, indicating good interaction between the cells and nanofibers. Together with the resazurin assay, we can conclude that the material is noncytotoxic and biocompatible.

Figure 5A shows the antioxidant activity of PCL/lignin− PCLLA nanofibers. The neat PCL nanofibers exhibited low antioxidant activity (12.9 ± 1.0% at 24 h). The incorporation of lignin copolymers improved the antioxidant activity of the nanofibers, and PCL/LP1 nanofibers showed the highest free radical inhibition (99.4 ± 0.5% at 24 h). Similarly, the free radical inhibition of neat PLLA nanofibers was only 8.6 ± 1.6% after 24 h of incubation (Figure 5B), but all of the PLLA/ lignin−PCLLA nanofibers exhibited excellent antioxidant activity (>70%). The good antioxidant activities of these nanofibers may enable their application for food packaging and biomedical materials to address issues of oxidative stress. Biodegradation Behavior of Electrospun Nanofibers. The biodegradability of the nanofibers was evaluated by examining their morphological changes after soaking in PBS. The accelerated biodegradation test was applied by using PBS (pH 10). As shown in Figure S5, after soaking for 4 weeks, neat the PCL and PLLA nanofibers retained their morphology and exhibited very little swelling or degradation. However, the nanofibers with lignin copolymers were found to undergo certain degradation, and the fibers swelled, softened, collapsed, and merged together, especially for PCL/LP1 and PLLA/LP1. It is interesting to observe that the addition of lignin copolymers could influence the degradation rate of the polyester nanofibers. These interesting results indicate that a longer-term study of the degradation of these lignin copolymers is needed to understand their lifetime behavior. Biocompatibility of Electrospun Nanofibers. The biocompatibility of lignin and its copolymers is still unclear, and few studies have assessed their cytotoxicity.54,55 In our previous studies, we demonstrated the biocompatibility of various lignin copolymers (lignin−PEG methyl ether meth-

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CONCLUSION

In this study, we synthesized a series of lignin−PCLLA copolymers via solvent-free ROP. The obtained lignin copolymers showed tunable molecular weights and thermal properties. Upon blending with PCL or PLLA, uniform 6022

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Figure 7. Fluorescence microscopy images of NIH/3T3 cells on nanofibers. The cell nuclei were stained with DAPI (blue), and the F-actin filaments were stained with phalloidin (green). Images of PLLA/lignin nanofibers are presented in the left column [(A) PLLA/LP1, (C) PLLA/LP2, (E) PLLA/LP3] and images of PCL/lignin nanofibers in the right column [(B) PCL/LP1, (D) PCL/LP2, (F) PCL/LP3]. Scale bars = 100 μm.

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nanofibers were generated via electrospinning. The incorporation of the copolymers reinforced the PCL nanofibers, but exhibited a negative influence on the mechanical properties of the PLLA nanofibers. Moreover, these lignin-based nanofibers exhibited good antioxidant activity and biocompatibility and thus have potential for biomedical or healthcare applications.

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected]. ORCID

Zibiao Li: 0000-0002-0591-5328 Zheng Zhang: 0000-0003-3127-8710 Xian Jun Loh: 0000-0001-8118-6502

ASSOCIATED CONTENT

S Supporting Information *

Notes

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acssuschemeng.7b00850. NMR spectra of LP1 and LP3 (Figure S1), XPS spectra of lignin−PCLLA copolymers (Figure S2), DSC curves of lignin−PCLLA copolymers (Figure S3), free radical inhibition of lignin−PCLLA copolymers (Figure S4), and biodegradation of nanofibers (Figure S5) (PDF)

The authors declare no competing financial interest.

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