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Hydrothermal-process-based direct extraction of polydisperse lignin microspheres from black liquor and their physicochemical characterization
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Hydrothermal-process-based direct extraction of polydisperse lignin microspheres from black liquor and their physicochemical characterization ⁎
Young-Lok Cha, Al-Mahmnur Alam , Sung-Min Park, Youn-Ho Moon, Kwang-Soo Kim, Ji-Eun Lee, Da-Eun Kwon, Yong-Gu Kang Bioenergy Crop Research Institute, National Institute of Crop Science, Rural Development Administration, Muan 58545, Republic of Korea
G R A P H I C A L A B S T R A C T
A R T I C LE I N FO
A B S T R A C T
Keywords: Black liquor Extraction Microspheres FESEM FTIR
Lignin nano-/microstructures are widely employed for agricultural drug delivery and heavy metal removal from wastewater, and facile low-cost methods of their large-scale production are therefore highly sought after. Herein, uniform-morphology polydisperse lignin microspheres were directly extracted from black liquor by lowering its pH to < 4 followed by hydrothermal treatment and featured several lignin-typical characteristics, e.g., functional groups, thermal stability, amorphousness, and monolignol units. It was assumed that lignin rearranged and assembled into microspheres of various size, shape, and uniformity depending on pH, temperature, and hydrothermal treatment time. Lignin microsphere extraction efficiency was estimated as 15.87–21.62 g L−1, and the average size of microspheres obtained under different conditions was calculated as ∼1 µm, while the C, H, O, and N contents equaled 48–62, 5–6, 30–36, and 0.2–1.5%, respectively. Thus, our method was deemed suitable for direct large-scale extraction of lignin microspheres from black liquor.
1. Introduction Together with cellulose and hemicellulose, lignin is a major component of the cell walls of vascular plants such as grasses, agricultural crops, and trees (Xu et al., 2006). This polyphenolic compound features an amorphous structure and is produced by enzymatic oxidative polymerization of monolignols such as sinapyl, coniferyl, and p-coumaryl
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alcohols (Thakur and Thakur, 2015), thus comprising syringyl (S), guaiacyl (G), and p-hydroxyphenyl (H) moieties, respectively (Sasaki et al., 2004). The hydrophobicity oflignin precludes (hemi)cellulose contained in plant cell walls from absorbing water and thus permits efficient water transport. Moreover, lignin offers structural and antimicrobial support, providing structural rigidity and absorbing ultraviolet radiation to protect plant cells from pest and pathogen attack
Corresponding author. Tel.: +1 657 256 8194. E-mail addresses: [email protected], [email protected] (A.-M. Alam).
https://doi.org/10.1016/j.biortech.2019.122399 Received 11 September 2019; Received in revised form 6 November 2019; Accepted 7 November 2019 0960-8524/ © 2019 Elsevier Ltd. All rights reserved.
Please cite this article as: Young-Lok Cha, et al., Bioresource Technology, https://doi.org/10.1016/j.biortech.2019.122399
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of directly extracting uniform LMSs from BL collected at the stage of twin screw–based alkaline pretreatment of Miscanthus sacchariflorus during bioethanol production was developed. Specifically, BL pH was lowered to 1–4 via the dropwise addition of 25, 50, or 98% H2SO4 in different batches followed by autoclaving at 121 °C. Several experimental conditions were tested, and LMSs were obtained in all cases with sufficiently high extraction efficiency and characterized in terms of morphology/distribution, lignin-like texture, organic functional groups, thermal stability, crystallinity and structural components, and oxygen/ carbon content.
(Ithal et al., 2007; Schuetz et al., 2014). Recently, much attention has been directed at the development and utilization of lignin as a potential biomaterial for the production of biobased chemicals/high-performance polymer nanocomposites as well as for biological and pharmaceutical applications (Calvo-Flores and Dobado, 2010). For example, lignin can be employed as a sustainable and biodegradable alternative to petroleum and other fossil fuels such as coal and natural gas in a wide range of industrial applications (Kumar et al., 2019). However, lignin is strongly attached to cellulose, hemicellulose, and other organic components of lignocellulosic plant and agricultural biomass (Calvo-Flores and Dobado, 2010), and its isolation therefore requires delignification and purification procedures. The production of paper, pulp, and biofuel involves the separation of lignin from lignocellulosic biomass during alkaline pretreatment, which concomitantly affords highly alkaline black liquor (BL; pH 13–14) containing dissolved lignin and other organic/inorganic components (Wallberg et al., 2003). The exact composition of BL depends on pretreatment conditions and biomass source. On average, BL features a solid content of 12–18% and a lignin content of 25–35%, with aliphatic carboxylic acids, other organics, and inorganics accounting for 30–35, 5–10, and 30–40% of the total solids, respectively (Alén, 2015; Marchessault, 1994; Wallberg et al., 2003). Conventionally, soluble lignin in BL is precipitated at pH of acidic range and is obtained as a brownish solid composed of irregular-shaped aggregated particles. Lignin has a complex organic structure and (in its native form) is prone to clustering in solvents (e.g., water), which affects surface area–dependent physicochemical properties (Humpert et al., 2016). Moreover, native lignin shows low metal ion/drug molecule adsorption capacities and response rates, which hinders its applications as an agricultural drug (pesticide, herbicide, and fertilizer) delivery agent, biomedical agent, and adsorbent for heavy metal removal from wastewater (Demirbas, 2004; Kumar et al., 2019; Šćiban et al., 2011). Nevertheless, lignin is still a promising adsorbent and drug carrier because of its abundant oxygenated functional groups, thermal stability, biocompatibility, and low cost (Ge et al., 2014). The properties of nano-/microstructured lignin are considerably different from those of bulk lignin of the same composition because of the larger accessible surface area available for binding of guest material in the former case with or without functionalization. Although the preparation and functionalization of lignin-based microspheres for encapsulating to-be-delivered hydrophobic/hydrophilic drugs has been extensively researched (Tortora et al., 2014; Yiamsawas et al., 2014), lignin microsphere (LMS) preparation by emulsification of aqueous lignin solutions in olive oil or cyclohexane containing toluene diisocyanate and a surfactant has received little attention. Lignin acetate microparticles for the controlled release of agricultural chemicals such as fertilizers, herbicides, and pesticides can be synthesized by a solvent evaporation method (Witzler et al., 2018). Moreover, sub-nanometerscale lignin spheres can be prepared by carbonization of commercial lignin, gradual addition of hydrochloric acid to a solution of lignin and ethylene glycol, gradual addition of non-solvent (water) to a solution of lignin acetate in tetrahydrofuran, or the addition of alkaline lignin solution to a solution of a cationic polyelectrolyte such as poly (diallyldimethylammonium chloride) (Antunes et al., 2014; Frangville et al., 2012; Gonugunta et al., 2012; Jiang et al., 2013; Jongpaiboonkit et al., 2009; Qian et al., 2014). Notably, all previous reports employed either laboratory-extracted or commercially available lignin as a precursor for the synthesis of lignin nano-/microspheres through the dissolution and reassembly of lignin molecules, utilizing expensive chemicals and laboratory facilities. Moreover, these methods are not upscalable and are thus poorly suited for the commercial production of uniform LMSs required for agricultural drug delivery to plants, removal of heavy metals from waste water, and other applications. Finally, the synthesized microspheres should be uniform to ensure the constant release of agricultural drugs or sufficient adsorption of heavy metals. To address the above problems, an upscalable and low-cost method
2. Materials and methods 2.1. Lignin microsphere preparation The raw Miscanthus sacchariflorus (an agricultural biomass) was composed of 40.3 wt% cellulose, 24.1 wt% hemicellulose, and 24.1 wt% lignin. The pretreatment of Miscanthus was prepared by a previous study with some modifications (Cha et al., 2016). Briefly, Miscanthus was chopped and ground to obtain 3 mm sized particles. The sieved samples were driven into the continuous twin screw extrusion pretreatment system with 0.5 M NaOH as a catalyst, 100 ℃, 8 min residence time and 50 rpm screw speed. After pretreatment, the black liquor was collected using a screw press with a screening mesh size of 1 mm. The amount of BL was 5 L/kg Miscanthus. In this case, the mixture ratio to the hydrolysis was 5 kg/h of Miscanthus and 30 L/h of 0.5 M NaOH as a catalyst. Collected BL was centrifuged at 12000 rpm for 30 min to remove insoluble debris and stored in a glass bottle for further use. The pH measured with a calibrated pH meter for 13 different BL samples considered for LMS extraction lied in the range of 13–14. Aqueous stock solutions of 25 and 50% H2SO4 were prepared from 98% H2SO4, and extraction was performed using a slight modification of a previously reported technique (Mussatto et al., 2007). In brief, the pH of highly alkaline BL was lowered to 1, 2, or 4 by the dropwise addition of 25, 50, or 98% H2SO4 in different batches, which caused a distinct black-to-brown color change. The acidified mixture was autoclaved at 121 °C for 1 or 3 h, and the resulting light-brown samples were centrifuged to afford pellets that were thoroughly washed 5–6 times with deionized water until supernatant pH reached a value of 6–7. The solids were dried at 60 °C for 12 h, pulverized to obtain LMSs as a brown powder, and weighed. In total, 13 different reaction conditions were tested for LMS extraction, and extraction efficiency was calculated from the extracted LMS amount and BL working volume as 15.87–21.62 g L−1. 2.2. Material characterization LMS morphology and structure were probed by field-emission scanning electron microscopy (FESEM; S-2460N, Hitachi, Japan). The size distribution of each sample was obtained by measuring the diameter of 400 particles of each set of samples, presented as a histogram, and fitted by a normal distribution. Fourier transform infrared (FTIR) spectra (Nicolet 6700 FT-IR, Thermo Nicolet Corp., USA) of solid samples were recorded in attenuated total reflectance (ATR) mode in the range of 4000–500 cm−1. LMS thermal stability was probed by thermogravimetric analysis (TGA; DTG-60H, Shimadzu, Japan) in an atmosphere of N2 at a heating rate of 10 °C min−1 in the temperature range of 20–900 °C. X-ray diffraction (XRD; D/MAX Ultima III, Rigaku, Japan) patterns were recorded using Ni-filtered Cu Kα radiation (λ = 0.1541 nm) generated at 40 kV/40 mA. A scan speed of 0.02° s−1 and a 2θ range of 5–80° were employed. Gas chromatography coupled with mass spectrometry (GC–MS; 7890GC and 5977B MSD, Agilent Technologies, Inc., USA) was used to analyze LMS components. Data acquisition was performed using ChemStation software, and the analyzed compounds were identified with the help of the NIST Tandem Mass Spectral Library (version 2.3, build May 2, 2017). Elemental 2
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the cleavage of β-O-4 ether bonds during delignification. The broad peak at 2800–2980 cm−1 was ascribed to C–H stretching vibrations of methyl and methylene units, and the wide absorption at 1703 cm−1 indicated the presence of C]O bonds not conjugated to aromatic rings. The sharp absorptions at 1590, 1507, 1458, and 1420 cm−1 were attributed to the skeletal vibrations of C]C aromatic units forming the fundamental structure of lignin. Absorptions at 1400–800 cm−1 were assigned to coniferyl (G), p-coumaryl (H), and sinapyl (S) alcohol moieties (Thakur et al., 2014; Upton and Kasko, 2016). The typical aliphatic C–H stretch of acetate methyl groups was observed at 1367 cm−1 and indicated the natural acetylation of the obtained lignin. The peak at 1326 cm−1 was ascribed to syringyl ring breathing with C–O stretching vibration. All LMSs featured a sharp absorption peak at 1121 cm−1 (in the fingerprint region), which revealed the presence of guaiacyl-syringyl (GS) moieties. Peaks at 1030 and 832 cm−1 were assigned to aromatic in-plane bending and C–H out-of-plane deformation, respectively.
compositions were determined using an elemental analyzer (Thermo Scientific, FLASH 2000 series). 2.3. Sample preparation and analysis Dried and well-ground LMS powder was used for FESEM, FTIR, TGA, XRD and elemental composition analyses. In the case of FESEM, well-ground LMS powder was placed on the sample holder and sputtercoated with Au prior to imaging. As liquid fractionation was required for GC–MS analysis, LMSs were dispersed in a 1:1 (v/v) mixture of ethanol and acetone, sonicated for 5 h, and then centrifuged at 12,000 rpm for 10 min. Finally, 2.0-mL extract aliquots were filtered through a 0.20-μm membrane for GC–MS analysis. Separation was performed on an HP-5MS (30 m × 250 μm × 0.25 μm i.d.) fused silica capillary column. The oven temperature was held at 60 °C for 1 min, then raised to 280 °C at 15 °C min−1, and held constant for 2 min. The injector temperature equaled 300 °C, and split mode was employed. Helium was used as the carrier gas at a flow rate of 2 mL min−1.
3.3. X-ray diffraction analysis 3. Results and discussion XRD analysis was performed to examine sample crystallinity, demonstrating that the patterns of all LMSs were similar to that of lignin. The representative XRD patterns of LMSs and lignin were compared with the dried biomass. The dried biomass pattern featured two peaks at ∼16 and 23° which were ascribed to amorphous carbon and the (0 0 2) reflection of carbon nanocrystallites present in the cellulose of dried biomass (Newman, 1999; Park et al., 2004; Wang et al., 2012). In contrast, a broad diffraction peak at 10–30° was observed for LMSs and lignin. In the LMS pattern, the peak at ∼16° disappeared, and that at ∼23° became wider because of chemical pretreatment, which was indicative of particle amorphousness (Park et al., 2004).
3.1. Lignin microsphere morphology and distribution FESEM analysis revealed that LMSs were formed in the autoclave under all experimental conditions and featured a reaction condition–dependent morphology. SEM images of LMSs extracted by 98% H2SO4 at different pH and autoclaving times were captured. It was found that LMSs with an average size of 3.7 ± 0.18 µm and a rough surface were formed at pH 1 after 1 h autoclaving. When the autoclaving time was increased to 3 h, the LMS surface became smoother, but the average size hardly changed. In contrast, LMS morphology was noticeably affected by increasing the pH of BL from 1 to 2, in which case LMSs with uniform size and shape were formed after 1 and 3 h autoclaving. When BL pH was further increased to 4, LMSs extracted after 1 and 3 h autoclaving exhibited a morphology similar to that observed for pH 2. Considering environmental safety, the consumption of H2SO4 was reduced as much as possible by using more dilute H2SO4 (50 and 25%) to extract LMSs from BL at pH 4 while maintaining other conditions constant. The corresponding size distributions of LMSs extracted from BL at pH 4 by 25 and 50% H2SO4 are presented in Fig. 1, which reveals that these particles featured a similar morphology and a narrow size distribution. The average diameter of LMSs extracted after 1 and 3 h autoclaving were determined as 0.8 ± 0.14 and 0.9 ± 0.18 µm, respectively. In contrast, LMSs extracted by 50% H2SO4 at pH 4 after 1 and 3 h autoclaving featured an average diameter of 1.0 ± 0.18 µm. However, no LMSs were detected in the extracted powder obtained at pH 4 under ambient conditions (room temperature) after BL acidification with 25, 50, or 98% H2SO4. This finding highlighted the crucial role of hydrothermal treatment in LMS formation, which was assumed to involve the rearrangement of lignin molecules through bond breakage followed by reassembly into large spheres.
3.4. Thermogravimetric analysis TGA and differential thermogravimetry (DTG) analysis of LMSs were performed to assess their thermal stability and degradation at high temperature. Typically, lignin is composed of different kinds of aromatic rings and organic functional groups linked through chemical bonds, which leads to the variation of degradation temperature in the range of 25–1000 °C. Herein, three stages of degradation were observed for all LMS samples except for dried biomass. Fig. 2(a) and (b) show representative TGA and DTG curves of LMSs isolated from BL at pH 4 using 98, 50, and 25% H2SO4 and 3 h autoclaving at 121 °C. The initial weight loss at 25–150 °C was ascribed to the evaporation of absorbed moisture, while the gradual decline between 150 and 600 °C was attributed to the degradation of lignin carbohydrates to afford volatiles such as CO, CO2, and CH4 (Watkins et al., 2015). The final degradation phase was slow and occurred at 600–1000 °C. On average, 30–36 wt% of LMSs remained non-volatilized even at 800–1000 °C (Fig. 2(a)), which was ascribed to the formation of highly compacted aromatic structures that could be converted into char (Yang et al., 2007), in agreement with previously reported results (Dhyani and Bhaskar, 2018; Fang et al., 2008; Jin et al., 2013; Park et al., 2004; Sangchoom and Mokaya, 2015; Tejado et al., 2007; Watkins et al., 2015; Yang et al., 2006). The TGA curve of dried biomass exhibited slightly different features (Fig. 2(a)), revealing moisture loss below 130 °C followed by sharp decomposition between 200 and 380 °C and slow decomposition at 400–1000 °C to afford a solid residue amounting to < 20 wt% of the original sample. This behavior was explained by the co-presence of cellulose, hemicellulose, and lignin in the dried biomass. Cellulose is known to undergo fast decomposition at 315–400 °C to leave only small amounts of residue, while the decomposition of lignin occurs slowly in a temperature range from ambient to 1000 °C (Dhyani and Bhaskar, 2018). The DTG profile in Fig. 2(b) indicates that the maximum weight loss rate of cellulose components occurred at 330 °C for dried biomass, while the presence of lignin in LMSs resulted in a weight loss over a
3.2. Fourier transform infrared spectroscopy analysis LMS functional groups were probed by ATR-FTIR spectroscopy, which revealed that the absorptions bands associated with lignin functional groups were observed in the range of 4000–400 cm−1, with the most informative bands lying in the region of 1800–800 cm−1 for all samples. The representative FTIR spectra of LMSs were compared with the spectrum of dried biomass. The peak maxima of the fundamental absorption bands of lignin, assigned to the corresponding functional groups present in LMSs were in good agreement with the results previously reported for lignin (Bui et al., 2015; Manara et al., 2014; Sun et al., 2013). The broad absorption peak at 3700–3100 cm−1 observed for LMSs and lignin clearly indicated the existence of abundant hydroxyl groups, some of which may have been produced through 3
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Fig. 1. Size distributions of LMSs obtained using 25 (a, b) and 50% (c, d) H2SO4 at autoclaving times of 1 and 3 h (particle sets with 100 nm-sized bins). Solid curves represent normal fits to the distributions. Scale bar = 5 µm.
building blocks of lignin, (Jiang et al., 2019; Wang et al., 2019; Xu et al., 2018). Moreover, compared to those of dried biomass, the TICs of LMSs featured several hydrocarbon peaks with different retention times that disappeared or shifted. The TIC peak intensity of selected compounds such as coniferyl (at 5.6 min), and sinapyl (at 9.5 min) alcohols indicated that the concentration and distribution of these compounds was higher for pure lignin. In contrast, the concentration of p-coumaryl (at 8.5 min) alcohol was found to be lower in lignin than in LMSs, according to TIC peak intensity. The obtained results indicated the complete or partial conversion of one organic component to another one or its derivatization during the thermal treatment of lignin in an acidic environment at 121 °C, and the conversion of lignin components was concluded to play a vital role in the bulk-to-microsphere change of lignin morphology.
wide temperature range of 175–385 °C (Dhyani and Bhaskar, 2018). 3.5. Gas chromatography-mass spectrometry analysis The total ion chromatograms (TICs) of the selected liquid fraction from LMSs obtained by hydrothermal acidic hydrolysis and of lignin extracted under the same acidic conditions without autoclaving were recorded. Several compounds were identified in liquids extracted from lignin using 98, 50 and 25% H2SO4 without autoclaving. The TIC data indicated the coexistence of three organic molecules at the retention time of 5.6, 8.5 and 9.5 min respectively, in all the selected samples before and after autoclaving along with other phenolic compounds and aromatic hydrocarbons. The structure of the three molecules found in MS scan data during GC–MS analysis in the sample resembles the structure of coniferyl, paracoumaryl, and sinapyl alcohol, which are
Fig. 2. (a) TGA and (b) DTG profiles of dried biomass and LMSs extracted by 25, 50 and 98% H2SO4 at pH 4 after 3 h autoclaving at 121 °C. 4
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Table 1 C, N, H, and O contents of dried biomass, extracted LMSs, and lignin. Sample
C (%)
Dried biomass (Miscanthus)
H (%)
O (%)
N (%)
44.78 ± 0.18
5.69 ± 0.03
41.33 ± 0.01
0.13 ± 0.01
98% H2SO4
pH1_1 h pH1_3 h pH2_1 h pH2_3 h pH4_1 h pH4 _3 h
60.60 62.20 56.95 60.80 50.56 50.82
5.77 5.85 5.81 5.84 5.63 5.31
30.68 29.32 33.51 30.28 30.25 30.24
0.64 0.69 0.87 0.93 1.41 1.50
50% H2SO4
pH4_1 h pH4_3 h
50.31 ± 0.68 48.88 ± 0.11
5.53 ± 0.14 5.11 ± 0.05
35.16 ± 0.002 30.25 ± 0.17
1.09 ± 0.01 0.32 ± 0.01
25% H2SO4
pH4_1 h pH4_3 h
49.12 ± 0.09 49.86 ± 0.01
4.99 ± 0.01 5.64 ± 0.05
31.04 ± 0.01 32.43 ± 0.03
0.34 ± 0.01 0.56 ± 0.06
98% H2SO4
pH4 Ambient
50.82 ± 0.15
5.26 ± 0.06
34.45 ± 0.01
0.24 ± 0.05
50% H2SO4
pH4 Ambient
54.47 ± 0.24
6.07 ± 0.03
35.34 ± 0.25
0.99 ± 0.01
25% H2SO4
pH4 Ambient
53.18 ± 0.18
5.92 ± 0.07
36.06 ± 0.02
0.88 ± 0.04
± ± ± ± ± ±
0.07 0.13 0.001 0.43 0.23 0.45
± ± ± ± ± ±
0.08 0.05 0.01 0.02 0.04 0.05
3.6. Elemental analysis
Korea.
The C, H, O, and N contents of LMSs (Table 1) were in the ranges of 48–62, 5–6, 30–36, and 0.2–1.5%, respectively, and barely influenced by reaction conditions. In contrast, dried biomass featured C, H, O, and N contents of 44.78, 0.13, 5.69, and 41.33%, respectively. The C content of LMSs exceeded that of dried biomass by 5–16%, while the O content of LMSs was lower than that of dried biomass by 5–10%. The C and O content of LMSs varied by 3.5–4.5 and 2.0–2.8%, respectively, according to the total amount of LMSs obtained under each reaction condition.
Appendix A. Supplementary data
± ± ± ± ± ±
0.003 0.07 0.04 0.03 0.06 0.01
± ± ± ± ± ±
0.04 0.01 0.01 0.18 0.08 0.13
Supplementary data to this article can be found online at https:// doi.org/10.1016/j.biortech.2019.122399. References Alén, R., 2015. Pulp mills and wood-based biorefineries, in: Industrial Biorefineries and White Biotechnology. pp. 91-126. doi: 10.1016/B978-0-444-63453-5.00003-3. Antunes, P.V., Ramalho, A., Carrilho, E.V.P., 2014. Mechanical and wear behaviours of nano and microfilled polymeric composite. effect of filler fraction and size. Mater. Des. https://doi.org/10.1016/j.matdes.2014.04.056. Bui, N.Q., Fongarland, P., Rataboul, F., Dartiguelongue, C., Charon, N., Vallée, C., Essayem, N., 2015. FTIR as a simple tool to quantify unconverted lignin from chars in biomass liquefaction process: application to SC ethanol liquefaction of pine wood. Fuel Process. Technol. https://doi.org/10.1016/j.fuproc.2015.02.020. Calvo-Flores, F.G., Dobado, J.A., 2010. Lignin as renewable raw material. ChemSusChem. https://doi.org/10.1002/cssc.201000157. Cha, Y.L., Yang, J.W., Seo, S.L., An, G.H., Moon, Y.H., You, G.D., Lee, J.E., Ahn, J.W., Lee, K.B., 2016. Alkaline twin-screw extrusion pretreatment of Miscanthus with recycled black liquor at the pilot scale. Fuel 164, 322–328. Demirbas, A., 2004. Adsorption of lead and cadmium ions in aqueous solutions onto modified lignin from alkali glycerol delignication. J. Hazard. Mater. 109, 221–226. https://doi.org/10.1016/j.jhazmat.2004.04.002. Dhyani, V., Bhaskar, T., 2018. A comprehensive review on the pyrolysis of lignocellulosic biomass. Renew. Energy. https://doi.org/10.1016/j.renene.2017.04.035. Fang, Z., Sato, T., Smith, R.L., Inomata, H., Arai, K., Kozinski, J.A., 2008. Reaction chemistry and phase behavior of lignin in high-temperature and supercritical water. Bioresour. Technol. https://doi.org/10.1016/j.biortech.2007.08.008. Frangville, C., Rutkevičius, M., Richter, A.P., Velev, O.D., Stoyanov, S.D., Paunov, V.N., 2012. Fabrication of environmentally biodegradable lignin nanoparticles. ChemPhysChem. https://doi.org/10.1002/cphc.201200537. Ge, Y., Xiao, D., Li, Z., Cui, X., 2014. Dithiocarbamate functionalized lignin for efficient removal of metallic ions and the usage of the metal-loaded bio-sorbents as potential free radical scavengers. J. Mater. Chem. A 2, 2136–2145. https://doi.org/10.1039/ c3ta14333c. Gonugunta, P., Vivekanandhan, S., Mohanty, A.K., Misra, M., 2012. A study on synthesis and characterization of biobased carbon nanoparticles from lignin. World J. Nano Sci. Eng. https://doi.org/10.4236/wjnse.2012.23019. Humpert, D., Ebrahimi, M., Czermak, P., 2016. Membrane technology for the recovery of lignin: a review. Membranes (Basel). https://doi.org/10.3390/membranes6030042. Ithal, N., Recknor, J., Nettleton, D., Maier, T., Baum, T.J., Mitchum, M.G., 2007. Developmental transcript profiling of cyst nematode feeding cells in soybean roots. Mol. Plant-Microbe Interact. https://doi.org/10.1094/mpmi-20-5-0510. Jiang, C., He, H., Jiang, H., Ma, L., Jia, D.M., 2013. Nano-lignin filled natural rubber composites: preparation and characterization. Express Polym. Lett. https://doi.org/ 10.3144/expresspolymlett.2013.44. Jiang, P., Li, Q., Gao, C., Lu, J., Cheng, Y., Zhai, S., An, Q., Wang, H., 2019. Fractionation of alkali lignin by organic solvents for biodegradable microsphere through self-assembly. Bioresour. Technol. https://doi.org/10.1016/j.biortech.2019.121640. Jin, W., Singh, K., Zondlo, J., 2013. Pyrolysis kinetics of physical components of wood and wood-polymers using isoconversion method. Agriculture. https://doi.org/10. 3390/agriculture3010012. Jongpaiboonkit, L., Franklin-Ford, T., Murphy, W.L., 2009. Growth of hydroxyapatite coatings on biodegradable polymer microspheres. ACS Appl. Mater. Interfaces.
4. Conclusions Herein, a new technique for direct lignin microsphere extraction from highly alkaline BL based on pH control and hydrothermal processing was presented. The obtained microspheres featured a polydisperse morphology, smooth surface, narrow size distribution (0.8–1.0 µm), typical lignin-like thermal stability, significant carbon content (50–60%), and an extraction efficiency of 15.87–21.62 g L−1, which reflected the uniqueness of our method and its upscalability. The high carbon content, non-toxicity, and biodegradability of LMSs make them well suited for use as agricultural drug delivery agents, heavy metal adsorbents, and UV absorbers for sun cream formulations. CRediT authorship contribution statement Young-Lok Cha: . : Conceptualization, Writing - review & editing, Project administration. Al-Mahmnur Alam: Conceptualization, Methodology, Writing - original draft. Sung-Min Park: Software, Data curation, Formal analysis. Youn-Ho Moon: Resources, Investigation. Kwang-Soo Kim: Resources, Formal analysis. Ji-Eun Lee: Data curation, Investigation. Da-Eun Kwon: Formal analysis, Visualization. Yong-Gu Kang: . : Conceptualization. Declaration of Competing Interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. Acknowledgments This research was supported by the Cooperative Research Program for Agriculture Science & Technology Development (Project No. PJ012575022019), Rural Development Administration, Republic of 5
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https://doi.org/10.1021/am9001716. Kumar, A., Anushree, Kumar, J., Bhaskar, T., 2019. Utilization of lignin: a sustainable and eco-friendly approach. J. Energy Inst. https://doi.org/10.1016/j.joei.2019.03.005. Manara, P., Zabaniotou, A., Vanderghem, C., Richel, A., 2014. Lignin extraction from Mediterranean agro-wastes: Impact of pretreatment conditions on lignin chemical structure and thermal degradation behavior. Catal. Today. https://doi.org/10.1016/ j.cattod.2013.10.065. Marchessault, R.H., 1994. Wood chemistry, fundamentals and applications. Carbohydr. Res. https://doi.org/10.1016/0008-6215(94)90030-2. Mussatto, S.I., Fernandes, M., Roberto, I.C., 2007. Lignin recovery from Brewer’s spent grain black liquor. Carbohydr. Polym. https://doi.org/10.1016/j.carbpol.2007.03. 021. Newman, R.H., 1999. Estimation of the lateral dimensions of cellulose crystallites using 13C NMR signal strengths. Solid State Nucl. Magn. Reson. https://doi.org/10.1016/ S0926-2040(99)00043-0. Park, C.H., Kang, Y.K., Im, S.S., 2004. Biodegradability of cellulose fabrics. J. Appl. Polym. Sci. https://doi.org/10.1002/app.20879. Qian, Y., Deng, Y., Qiu, X., Li, H., Yang, D., 2014. Formation of uniform colloidal spheres from lignin, a renewable resource recovered from pulping spent liquor. Green Chem. https://doi.org/10.1039/c3gc42131g. Sangchoom, W., Mokaya, R., 2015. Valorization of lignin waste: carbons from hydrothermal carbonization of renewable lignin as superior sorbents for CO2 and hydrogen storage. ACS Sustain. Chem. Eng. https://doi.org/10.1021/acssuschemeng.5b00351. Sasaki, S., Nishida, T., Tsutsumi, Y., Kondo, R., 2004. Lignin dehydrogenative polymerization mechanism: a poplar cell wall peroxidase directly oxidizes polymer lignin and produces in vitro dehydrogenative polymer rich in β-O-4 linkage. FEBS Lett. 562, 197–201. https://doi.org/10.1016/S0014-5793(04)00224-8. Schuetz, M., Benske, A., Smith, R.A., Watanabe, Y., Tobimatsu, Y., Ralph, J., Demura, T., Ellis, B., Samuels, A.L., 2014. Laccases direct lignification in the discrete secondary cell wall domains of protoxylem. Plant Physiol. https://doi.org/10.1104/pp.114. 245597. Šćiban, M.B., Klašnja, M.T., Antov, M.G., 2011. Study of the biosorption of different heavy metal ions onto Kraft lignin. Ecol. Eng. 37, 2092–2095. https://doi.org/10. 1016/j.ecoleng.2011.08.006. Sun, Y., Qiu, X., Liu, Y., 2013. Chemical reactivity of alkali lignin modified with laccase. Biomass Bioenergy. https://doi.org/10.1016/j.biombioe.2013.02.006. Tejado, A., Peña, C., Labidi, J., Echeverria, J.M., Mondragon, I., 2007. Physico-chemical characterization of lignins from different sources for use in phenol-formaldehyde resin synthesis. Bioresour. Technol. https://doi.org/10.1016/j.biortech.2006.05.042. Thakur, V.K., Thakur, M.K., 2015. Recent advances in green hydrogels from lignin: a
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Hydrothermal-process-based direct extraction of polydisperse lignin microspheres from black liquor and their physicochemical characterization ⁎
Young-Lok Cha, Al-Mahmnur Alam , Sung-Min Park, Youn-Ho Moon, Kwang-Soo Kim, Ji-Eun Lee, Da-Eun Kwon, Yong-Gu Kang Bioenergy Crop Research Institute, National Institute of Crop Science, Rural Development Administration, Muan 58545, Republic of Korea
G R A P H I C A L A B S T R A C T
A R T I C LE I N FO
A B S T R A C T
Keywords: Black liquor Extraction Microspheres FESEM FTIR
Lignin nano-/microstructures are widely employed for agricultural drug delivery and heavy metal removal from wastewater, and facile low-cost methods of their large-scale production are therefore highly sought after. Herein, uniform-morphology polydisperse lignin microspheres were directly extracted from black liquor by lowering its pH to < 4 followed by hydrothermal treatment and featured several lignin-typical characteristics, e.g., functional groups, thermal stability, amorphousness, and monolignol units. It was assumed that lignin rearranged and assembled into microspheres of various size, shape, and uniformity depending on pH, temperature, and hydrothermal treatment time. Lignin microsphere extraction efficiency was estimated as 15.87–21.62 g L−1, and the average size of microspheres obtained under different conditions was calculated as ∼1 µm, while the C, H, O, and N contents equaled 48–62, 5–6, 30–36, and 0.2–1.5%, respectively. Thus, our method was deemed suitable for direct large-scale extraction of lignin microspheres from black liquor.
1. Introduction Together with cellulose and hemicellulose, lignin is a major component of the cell walls of vascular plants such as grasses, agricultural crops, and trees (Xu et al., 2006). This polyphenolic compound features an amorphous structure and is produced by enzymatic oxidative polymerization of monolignols such as sinapyl, coniferyl, and p-coumaryl
⁎
alcohols (Thakur and Thakur, 2015), thus comprising syringyl (S), guaiacyl (G), and p-hydroxyphenyl (H) moieties, respectively (Sasaki et al., 2004). The hydrophobicity oflignin precludes (hemi)cellulose contained in plant cell walls from absorbing water and thus permits efficient water transport. Moreover, lignin offers structural and antimicrobial support, providing structural rigidity and absorbing ultraviolet radiation to protect plant cells from pest and pathogen attack
Corresponding author. Tel.: +1 657 256 8194. E-mail addresses: [email protected], [email protected] (A.-M. Alam).
https://doi.org/10.1016/j.biortech.2019.122399 Received 11 September 2019; Received in revised form 6 November 2019; Accepted 7 November 2019 0960-8524/ © 2019 Elsevier Ltd. All rights reserved.
Please cite this article as: Young-Lok Cha, et al., Bioresource Technology, https://doi.org/10.1016/j.biortech.2019.122399
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of directly extracting uniform LMSs from BL collected at the stage of twin screw–based alkaline pretreatment of Miscanthus sacchariflorus during bioethanol production was developed. Specifically, BL pH was lowered to 1–4 via the dropwise addition of 25, 50, or 98% H2SO4 in different batches followed by autoclaving at 121 °C. Several experimental conditions were tested, and LMSs were obtained in all cases with sufficiently high extraction efficiency and characterized in terms of morphology/distribution, lignin-like texture, organic functional groups, thermal stability, crystallinity and structural components, and oxygen/ carbon content.
(Ithal et al., 2007; Schuetz et al., 2014). Recently, much attention has been directed at the development and utilization of lignin as a potential biomaterial for the production of biobased chemicals/high-performance polymer nanocomposites as well as for biological and pharmaceutical applications (Calvo-Flores and Dobado, 2010). For example, lignin can be employed as a sustainable and biodegradable alternative to petroleum and other fossil fuels such as coal and natural gas in a wide range of industrial applications (Kumar et al., 2019). However, lignin is strongly attached to cellulose, hemicellulose, and other organic components of lignocellulosic plant and agricultural biomass (Calvo-Flores and Dobado, 2010), and its isolation therefore requires delignification and purification procedures. The production of paper, pulp, and biofuel involves the separation of lignin from lignocellulosic biomass during alkaline pretreatment, which concomitantly affords highly alkaline black liquor (BL; pH 13–14) containing dissolved lignin and other organic/inorganic components (Wallberg et al., 2003). The exact composition of BL depends on pretreatment conditions and biomass source. On average, BL features a solid content of 12–18% and a lignin content of 25–35%, with aliphatic carboxylic acids, other organics, and inorganics accounting for 30–35, 5–10, and 30–40% of the total solids, respectively (Alén, 2015; Marchessault, 1994; Wallberg et al., 2003). Conventionally, soluble lignin in BL is precipitated at pH of acidic range and is obtained as a brownish solid composed of irregular-shaped aggregated particles. Lignin has a complex organic structure and (in its native form) is prone to clustering in solvents (e.g., water), which affects surface area–dependent physicochemical properties (Humpert et al., 2016). Moreover, native lignin shows low metal ion/drug molecule adsorption capacities and response rates, which hinders its applications as an agricultural drug (pesticide, herbicide, and fertilizer) delivery agent, biomedical agent, and adsorbent for heavy metal removal from wastewater (Demirbas, 2004; Kumar et al., 2019; Šćiban et al., 2011). Nevertheless, lignin is still a promising adsorbent and drug carrier because of its abundant oxygenated functional groups, thermal stability, biocompatibility, and low cost (Ge et al., 2014). The properties of nano-/microstructured lignin are considerably different from those of bulk lignin of the same composition because of the larger accessible surface area available for binding of guest material in the former case with or without functionalization. Although the preparation and functionalization of lignin-based microspheres for encapsulating to-be-delivered hydrophobic/hydrophilic drugs has been extensively researched (Tortora et al., 2014; Yiamsawas et al., 2014), lignin microsphere (LMS) preparation by emulsification of aqueous lignin solutions in olive oil or cyclohexane containing toluene diisocyanate and a surfactant has received little attention. Lignin acetate microparticles for the controlled release of agricultural chemicals such as fertilizers, herbicides, and pesticides can be synthesized by a solvent evaporation method (Witzler et al., 2018). Moreover, sub-nanometerscale lignin spheres can be prepared by carbonization of commercial lignin, gradual addition of hydrochloric acid to a solution of lignin and ethylene glycol, gradual addition of non-solvent (water) to a solution of lignin acetate in tetrahydrofuran, or the addition of alkaline lignin solution to a solution of a cationic polyelectrolyte such as poly (diallyldimethylammonium chloride) (Antunes et al., 2014; Frangville et al., 2012; Gonugunta et al., 2012; Jiang et al., 2013; Jongpaiboonkit et al., 2009; Qian et al., 2014). Notably, all previous reports employed either laboratory-extracted or commercially available lignin as a precursor for the synthesis of lignin nano-/microspheres through the dissolution and reassembly of lignin molecules, utilizing expensive chemicals and laboratory facilities. Moreover, these methods are not upscalable and are thus poorly suited for the commercial production of uniform LMSs required for agricultural drug delivery to plants, removal of heavy metals from waste water, and other applications. Finally, the synthesized microspheres should be uniform to ensure the constant release of agricultural drugs or sufficient adsorption of heavy metals. To address the above problems, an upscalable and low-cost method
2. Materials and methods 2.1. Lignin microsphere preparation The raw Miscanthus sacchariflorus (an agricultural biomass) was composed of 40.3 wt% cellulose, 24.1 wt% hemicellulose, and 24.1 wt% lignin. The pretreatment of Miscanthus was prepared by a previous study with some modifications (Cha et al., 2016). Briefly, Miscanthus was chopped and ground to obtain 3 mm sized particles. The sieved samples were driven into the continuous twin screw extrusion pretreatment system with 0.5 M NaOH as a catalyst, 100 ℃, 8 min residence time and 50 rpm screw speed. After pretreatment, the black liquor was collected using a screw press with a screening mesh size of 1 mm. The amount of BL was 5 L/kg Miscanthus. In this case, the mixture ratio to the hydrolysis was 5 kg/h of Miscanthus and 30 L/h of 0.5 M NaOH as a catalyst. Collected BL was centrifuged at 12000 rpm for 30 min to remove insoluble debris and stored in a glass bottle for further use. The pH measured with a calibrated pH meter for 13 different BL samples considered for LMS extraction lied in the range of 13–14. Aqueous stock solutions of 25 and 50% H2SO4 were prepared from 98% H2SO4, and extraction was performed using a slight modification of a previously reported technique (Mussatto et al., 2007). In brief, the pH of highly alkaline BL was lowered to 1, 2, or 4 by the dropwise addition of 25, 50, or 98% H2SO4 in different batches, which caused a distinct black-to-brown color change. The acidified mixture was autoclaved at 121 °C for 1 or 3 h, and the resulting light-brown samples were centrifuged to afford pellets that were thoroughly washed 5–6 times with deionized water until supernatant pH reached a value of 6–7. The solids were dried at 60 °C for 12 h, pulverized to obtain LMSs as a brown powder, and weighed. In total, 13 different reaction conditions were tested for LMS extraction, and extraction efficiency was calculated from the extracted LMS amount and BL working volume as 15.87–21.62 g L−1. 2.2. Material characterization LMS morphology and structure were probed by field-emission scanning electron microscopy (FESEM; S-2460N, Hitachi, Japan). The size distribution of each sample was obtained by measuring the diameter of 400 particles of each set of samples, presented as a histogram, and fitted by a normal distribution. Fourier transform infrared (FTIR) spectra (Nicolet 6700 FT-IR, Thermo Nicolet Corp., USA) of solid samples were recorded in attenuated total reflectance (ATR) mode in the range of 4000–500 cm−1. LMS thermal stability was probed by thermogravimetric analysis (TGA; DTG-60H, Shimadzu, Japan) in an atmosphere of N2 at a heating rate of 10 °C min−1 in the temperature range of 20–900 °C. X-ray diffraction (XRD; D/MAX Ultima III, Rigaku, Japan) patterns were recorded using Ni-filtered Cu Kα radiation (λ = 0.1541 nm) generated at 40 kV/40 mA. A scan speed of 0.02° s−1 and a 2θ range of 5–80° were employed. Gas chromatography coupled with mass spectrometry (GC–MS; 7890GC and 5977B MSD, Agilent Technologies, Inc., USA) was used to analyze LMS components. Data acquisition was performed using ChemStation software, and the analyzed compounds were identified with the help of the NIST Tandem Mass Spectral Library (version 2.3, build May 2, 2017). Elemental 2
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the cleavage of β-O-4 ether bonds during delignification. The broad peak at 2800–2980 cm−1 was ascribed to C–H stretching vibrations of methyl and methylene units, and the wide absorption at 1703 cm−1 indicated the presence of C]O bonds not conjugated to aromatic rings. The sharp absorptions at 1590, 1507, 1458, and 1420 cm−1 were attributed to the skeletal vibrations of C]C aromatic units forming the fundamental structure of lignin. Absorptions at 1400–800 cm−1 were assigned to coniferyl (G), p-coumaryl (H), and sinapyl (S) alcohol moieties (Thakur et al., 2014; Upton and Kasko, 2016). The typical aliphatic C–H stretch of acetate methyl groups was observed at 1367 cm−1 and indicated the natural acetylation of the obtained lignin. The peak at 1326 cm−1 was ascribed to syringyl ring breathing with C–O stretching vibration. All LMSs featured a sharp absorption peak at 1121 cm−1 (in the fingerprint region), which revealed the presence of guaiacyl-syringyl (GS) moieties. Peaks at 1030 and 832 cm−1 were assigned to aromatic in-plane bending and C–H out-of-plane deformation, respectively.
compositions were determined using an elemental analyzer (Thermo Scientific, FLASH 2000 series). 2.3. Sample preparation and analysis Dried and well-ground LMS powder was used for FESEM, FTIR, TGA, XRD and elemental composition analyses. In the case of FESEM, well-ground LMS powder was placed on the sample holder and sputtercoated with Au prior to imaging. As liquid fractionation was required for GC–MS analysis, LMSs were dispersed in a 1:1 (v/v) mixture of ethanol and acetone, sonicated for 5 h, and then centrifuged at 12,000 rpm for 10 min. Finally, 2.0-mL extract aliquots were filtered through a 0.20-μm membrane for GC–MS analysis. Separation was performed on an HP-5MS (30 m × 250 μm × 0.25 μm i.d.) fused silica capillary column. The oven temperature was held at 60 °C for 1 min, then raised to 280 °C at 15 °C min−1, and held constant for 2 min. The injector temperature equaled 300 °C, and split mode was employed. Helium was used as the carrier gas at a flow rate of 2 mL min−1.
3.3. X-ray diffraction analysis 3. Results and discussion XRD analysis was performed to examine sample crystallinity, demonstrating that the patterns of all LMSs were similar to that of lignin. The representative XRD patterns of LMSs and lignin were compared with the dried biomass. The dried biomass pattern featured two peaks at ∼16 and 23° which were ascribed to amorphous carbon and the (0 0 2) reflection of carbon nanocrystallites present in the cellulose of dried biomass (Newman, 1999; Park et al., 2004; Wang et al., 2012). In contrast, a broad diffraction peak at 10–30° was observed for LMSs and lignin. In the LMS pattern, the peak at ∼16° disappeared, and that at ∼23° became wider because of chemical pretreatment, which was indicative of particle amorphousness (Park et al., 2004).
3.1. Lignin microsphere morphology and distribution FESEM analysis revealed that LMSs were formed in the autoclave under all experimental conditions and featured a reaction condition–dependent morphology. SEM images of LMSs extracted by 98% H2SO4 at different pH and autoclaving times were captured. It was found that LMSs with an average size of 3.7 ± 0.18 µm and a rough surface were formed at pH 1 after 1 h autoclaving. When the autoclaving time was increased to 3 h, the LMS surface became smoother, but the average size hardly changed. In contrast, LMS morphology was noticeably affected by increasing the pH of BL from 1 to 2, in which case LMSs with uniform size and shape were formed after 1 and 3 h autoclaving. When BL pH was further increased to 4, LMSs extracted after 1 and 3 h autoclaving exhibited a morphology similar to that observed for pH 2. Considering environmental safety, the consumption of H2SO4 was reduced as much as possible by using more dilute H2SO4 (50 and 25%) to extract LMSs from BL at pH 4 while maintaining other conditions constant. The corresponding size distributions of LMSs extracted from BL at pH 4 by 25 and 50% H2SO4 are presented in Fig. 1, which reveals that these particles featured a similar morphology and a narrow size distribution. The average diameter of LMSs extracted after 1 and 3 h autoclaving were determined as 0.8 ± 0.14 and 0.9 ± 0.18 µm, respectively. In contrast, LMSs extracted by 50% H2SO4 at pH 4 after 1 and 3 h autoclaving featured an average diameter of 1.0 ± 0.18 µm. However, no LMSs were detected in the extracted powder obtained at pH 4 under ambient conditions (room temperature) after BL acidification with 25, 50, or 98% H2SO4. This finding highlighted the crucial role of hydrothermal treatment in LMS formation, which was assumed to involve the rearrangement of lignin molecules through bond breakage followed by reassembly into large spheres.
3.4. Thermogravimetric analysis TGA and differential thermogravimetry (DTG) analysis of LMSs were performed to assess their thermal stability and degradation at high temperature. Typically, lignin is composed of different kinds of aromatic rings and organic functional groups linked through chemical bonds, which leads to the variation of degradation temperature in the range of 25–1000 °C. Herein, three stages of degradation were observed for all LMS samples except for dried biomass. Fig. 2(a) and (b) show representative TGA and DTG curves of LMSs isolated from BL at pH 4 using 98, 50, and 25% H2SO4 and 3 h autoclaving at 121 °C. The initial weight loss at 25–150 °C was ascribed to the evaporation of absorbed moisture, while the gradual decline between 150 and 600 °C was attributed to the degradation of lignin carbohydrates to afford volatiles such as CO, CO2, and CH4 (Watkins et al., 2015). The final degradation phase was slow and occurred at 600–1000 °C. On average, 30–36 wt% of LMSs remained non-volatilized even at 800–1000 °C (Fig. 2(a)), which was ascribed to the formation of highly compacted aromatic structures that could be converted into char (Yang et al., 2007), in agreement with previously reported results (Dhyani and Bhaskar, 2018; Fang et al., 2008; Jin et al., 2013; Park et al., 2004; Sangchoom and Mokaya, 2015; Tejado et al., 2007; Watkins et al., 2015; Yang et al., 2006). The TGA curve of dried biomass exhibited slightly different features (Fig. 2(a)), revealing moisture loss below 130 °C followed by sharp decomposition between 200 and 380 °C and slow decomposition at 400–1000 °C to afford a solid residue amounting to < 20 wt% of the original sample. This behavior was explained by the co-presence of cellulose, hemicellulose, and lignin in the dried biomass. Cellulose is known to undergo fast decomposition at 315–400 °C to leave only small amounts of residue, while the decomposition of lignin occurs slowly in a temperature range from ambient to 1000 °C (Dhyani and Bhaskar, 2018). The DTG profile in Fig. 2(b) indicates that the maximum weight loss rate of cellulose components occurred at 330 °C for dried biomass, while the presence of lignin in LMSs resulted in a weight loss over a
3.2. Fourier transform infrared spectroscopy analysis LMS functional groups were probed by ATR-FTIR spectroscopy, which revealed that the absorptions bands associated with lignin functional groups were observed in the range of 4000–400 cm−1, with the most informative bands lying in the region of 1800–800 cm−1 for all samples. The representative FTIR spectra of LMSs were compared with the spectrum of dried biomass. The peak maxima of the fundamental absorption bands of lignin, assigned to the corresponding functional groups present in LMSs were in good agreement with the results previously reported for lignin (Bui et al., 2015; Manara et al., 2014; Sun et al., 2013). The broad absorption peak at 3700–3100 cm−1 observed for LMSs and lignin clearly indicated the existence of abundant hydroxyl groups, some of which may have been produced through 3
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Fig. 1. Size distributions of LMSs obtained using 25 (a, b) and 50% (c, d) H2SO4 at autoclaving times of 1 and 3 h (particle sets with 100 nm-sized bins). Solid curves represent normal fits to the distributions. Scale bar = 5 µm.
building blocks of lignin, (Jiang et al., 2019; Wang et al., 2019; Xu et al., 2018). Moreover, compared to those of dried biomass, the TICs of LMSs featured several hydrocarbon peaks with different retention times that disappeared or shifted. The TIC peak intensity of selected compounds such as coniferyl (at 5.6 min), and sinapyl (at 9.5 min) alcohols indicated that the concentration and distribution of these compounds was higher for pure lignin. In contrast, the concentration of p-coumaryl (at 8.5 min) alcohol was found to be lower in lignin than in LMSs, according to TIC peak intensity. The obtained results indicated the complete or partial conversion of one organic component to another one or its derivatization during the thermal treatment of lignin in an acidic environment at 121 °C, and the conversion of lignin components was concluded to play a vital role in the bulk-to-microsphere change of lignin morphology.
wide temperature range of 175–385 °C (Dhyani and Bhaskar, 2018). 3.5. Gas chromatography-mass spectrometry analysis The total ion chromatograms (TICs) of the selected liquid fraction from LMSs obtained by hydrothermal acidic hydrolysis and of lignin extracted under the same acidic conditions without autoclaving were recorded. Several compounds were identified in liquids extracted from lignin using 98, 50 and 25% H2SO4 without autoclaving. The TIC data indicated the coexistence of three organic molecules at the retention time of 5.6, 8.5 and 9.5 min respectively, in all the selected samples before and after autoclaving along with other phenolic compounds and aromatic hydrocarbons. The structure of the three molecules found in MS scan data during GC–MS analysis in the sample resembles the structure of coniferyl, paracoumaryl, and sinapyl alcohol, which are
Fig. 2. (a) TGA and (b) DTG profiles of dried biomass and LMSs extracted by 25, 50 and 98% H2SO4 at pH 4 after 3 h autoclaving at 121 °C. 4
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Table 1 C, N, H, and O contents of dried biomass, extracted LMSs, and lignin. Sample
C (%)
Dried biomass (Miscanthus)
H (%)
O (%)
N (%)
44.78 ± 0.18
5.69 ± 0.03
41.33 ± 0.01
0.13 ± 0.01
98% H2SO4
pH1_1 h pH1_3 h pH2_1 h pH2_3 h pH4_1 h pH4 _3 h
60.60 62.20 56.95 60.80 50.56 50.82
5.77 5.85 5.81 5.84 5.63 5.31
30.68 29.32 33.51 30.28 30.25 30.24
0.64 0.69 0.87 0.93 1.41 1.50
50% H2SO4
pH4_1 h pH4_3 h
50.31 ± 0.68 48.88 ± 0.11
5.53 ± 0.14 5.11 ± 0.05
35.16 ± 0.002 30.25 ± 0.17
1.09 ± 0.01 0.32 ± 0.01
25% H2SO4
pH4_1 h pH4_3 h
49.12 ± 0.09 49.86 ± 0.01
4.99 ± 0.01 5.64 ± 0.05
31.04 ± 0.01 32.43 ± 0.03
0.34 ± 0.01 0.56 ± 0.06
98% H2SO4
pH4 Ambient
50.82 ± 0.15
5.26 ± 0.06
34.45 ± 0.01
0.24 ± 0.05
50% H2SO4
pH4 Ambient
54.47 ± 0.24
6.07 ± 0.03
35.34 ± 0.25
0.99 ± 0.01
25% H2SO4
pH4 Ambient
53.18 ± 0.18
5.92 ± 0.07
36.06 ± 0.02
0.88 ± 0.04
± ± ± ± ± ±
0.07 0.13 0.001 0.43 0.23 0.45
± ± ± ± ± ±
0.08 0.05 0.01 0.02 0.04 0.05
3.6. Elemental analysis
Korea.
The C, H, O, and N contents of LMSs (Table 1) were in the ranges of 48–62, 5–6, 30–36, and 0.2–1.5%, respectively, and barely influenced by reaction conditions. In contrast, dried biomass featured C, H, O, and N contents of 44.78, 0.13, 5.69, and 41.33%, respectively. The C content of LMSs exceeded that of dried biomass by 5–16%, while the O content of LMSs was lower than that of dried biomass by 5–10%. The C and O content of LMSs varied by 3.5–4.5 and 2.0–2.8%, respectively, according to the total amount of LMSs obtained under each reaction condition.
Appendix A. Supplementary data
± ± ± ± ± ±
0.003 0.07 0.04 0.03 0.06 0.01
± ± ± ± ± ±
0.04 0.01 0.01 0.18 0.08 0.13
Supplementary data to this article can be found online at https:// doi.org/10.1016/j.biortech.2019.122399. References Alén, R., 2015. Pulp mills and wood-based biorefineries, in: Industrial Biorefineries and White Biotechnology. pp. 91-126. doi: 10.1016/B978-0-444-63453-5.00003-3. Antunes, P.V., Ramalho, A., Carrilho, E.V.P., 2014. Mechanical and wear behaviours of nano and microfilled polymeric composite. effect of filler fraction and size. Mater. Des. https://doi.org/10.1016/j.matdes.2014.04.056. Bui, N.Q., Fongarland, P., Rataboul, F., Dartiguelongue, C., Charon, N., Vallée, C., Essayem, N., 2015. FTIR as a simple tool to quantify unconverted lignin from chars in biomass liquefaction process: application to SC ethanol liquefaction of pine wood. Fuel Process. Technol. https://doi.org/10.1016/j.fuproc.2015.02.020. Calvo-Flores, F.G., Dobado, J.A., 2010. Lignin as renewable raw material. ChemSusChem. https://doi.org/10.1002/cssc.201000157. Cha, Y.L., Yang, J.W., Seo, S.L., An, G.H., Moon, Y.H., You, G.D., Lee, J.E., Ahn, J.W., Lee, K.B., 2016. Alkaline twin-screw extrusion pretreatment of Miscanthus with recycled black liquor at the pilot scale. Fuel 164, 322–328. Demirbas, A., 2004. Adsorption of lead and cadmium ions in aqueous solutions onto modified lignin from alkali glycerol delignication. J. Hazard. Mater. 109, 221–226. https://doi.org/10.1016/j.jhazmat.2004.04.002. Dhyani, V., Bhaskar, T., 2018. A comprehensive review on the pyrolysis of lignocellulosic biomass. Renew. Energy. https://doi.org/10.1016/j.renene.2017.04.035. Fang, Z., Sato, T., Smith, R.L., Inomata, H., Arai, K., Kozinski, J.A., 2008. Reaction chemistry and phase behavior of lignin in high-temperature and supercritical water. Bioresour. Technol. https://doi.org/10.1016/j.biortech.2007.08.008. Frangville, C., Rutkevičius, M., Richter, A.P., Velev, O.D., Stoyanov, S.D., Paunov, V.N., 2012. Fabrication of environmentally biodegradable lignin nanoparticles. ChemPhysChem. https://doi.org/10.1002/cphc.201200537. Ge, Y., Xiao, D., Li, Z., Cui, X., 2014. Dithiocarbamate functionalized lignin for efficient removal of metallic ions and the usage of the metal-loaded bio-sorbents as potential free radical scavengers. J. Mater. Chem. A 2, 2136–2145. https://doi.org/10.1039/ c3ta14333c. Gonugunta, P., Vivekanandhan, S., Mohanty, A.K., Misra, M., 2012. A study on synthesis and characterization of biobased carbon nanoparticles from lignin. World J. Nano Sci. Eng. https://doi.org/10.4236/wjnse.2012.23019. Humpert, D., Ebrahimi, M., Czermak, P., 2016. Membrane technology for the recovery of lignin: a review. Membranes (Basel). https://doi.org/10.3390/membranes6030042. Ithal, N., Recknor, J., Nettleton, D., Maier, T., Baum, T.J., Mitchum, M.G., 2007. Developmental transcript profiling of cyst nematode feeding cells in soybean roots. Mol. Plant-Microbe Interact. https://doi.org/10.1094/mpmi-20-5-0510. Jiang, C., He, H., Jiang, H., Ma, L., Jia, D.M., 2013. Nano-lignin filled natural rubber composites: preparation and characterization. Express Polym. Lett. https://doi.org/ 10.3144/expresspolymlett.2013.44. Jiang, P., Li, Q., Gao, C., Lu, J., Cheng, Y., Zhai, S., An, Q., Wang, H., 2019. Fractionation of alkali lignin by organic solvents for biodegradable microsphere through self-assembly. Bioresour. Technol. https://doi.org/10.1016/j.biortech.2019.121640. Jin, W., Singh, K., Zondlo, J., 2013. Pyrolysis kinetics of physical components of wood and wood-polymers using isoconversion method. Agriculture. https://doi.org/10. 3390/agriculture3010012. Jongpaiboonkit, L., Franklin-Ford, T., Murphy, W.L., 2009. Growth of hydroxyapatite coatings on biodegradable polymer microspheres. ACS Appl. Mater. Interfaces.
4. Conclusions Herein, a new technique for direct lignin microsphere extraction from highly alkaline BL based on pH control and hydrothermal processing was presented. The obtained microspheres featured a polydisperse morphology, smooth surface, narrow size distribution (0.8–1.0 µm), typical lignin-like thermal stability, significant carbon content (50–60%), and an extraction efficiency of 15.87–21.62 g L−1, which reflected the uniqueness of our method and its upscalability. The high carbon content, non-toxicity, and biodegradability of LMSs make them well suited for use as agricultural drug delivery agents, heavy metal adsorbents, and UV absorbers for sun cream formulations. CRediT authorship contribution statement Young-Lok Cha: . : Conceptualization, Writing - review & editing, Project administration. Al-Mahmnur Alam: Conceptualization, Methodology, Writing - original draft. Sung-Min Park: Software, Data curation, Formal analysis. Youn-Ho Moon: Resources, Investigation. Kwang-Soo Kim: Resources, Formal analysis. Ji-Eun Lee: Data curation, Investigation. Da-Eun Kwon: Formal analysis, Visualization. Yong-Gu Kang: . : Conceptualization. Declaration of Competing Interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. Acknowledgments This research was supported by the Cooperative Research Program for Agriculture Science & Technology Development (Project No. PJ012575022019), Rural Development Administration, Republic of 5
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