Research Article pubs.acs.org/journal/ascecg
Valorizing Recalcitrant Cellulolytic Enzyme Lignin via Lignin Nanoparticles Fabrication in an Integrated Biorefinery Dong Tian,†,‡ Jinguang Hu,*,‡ Richard P. Chandra,‡ Jack N. Saddler,‡ and Canhui Lu*,† †
State Key Laboratory of Polymer Materials Engineering, Polymer Research Institute of Sichuan University, Chengdu 610065, China Forest Products Biotechnology/Bioenergy Group, Department of Wood Science, Faculty of Forestry, University of British Columbia,2424 Main Mall, Vancouver, British Columbia V6T 1Z4, Canada
‡
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S Supporting Information *
ABSTRACT: Conversion of condensed lignin into valueadded products in current lignocellulosic biorefineries has been challenging due to its structure recalcitrance. However, this work showed a technically feasible route to valorize recalcitrant cellulolytic enzyme lignin (CEL: lignin residue after enzymatic hydrolysis) via “high-quality” lignin nanoparticles (LNPs) fabrication. Three representative CELs obtained from hydrolysis of industrial relevant, steam-pretreated, agriculture reside corn stover, hardwood poplar, and softwood lodgepole pine were evaluated for their potential to produce LNPs through the prevalent dialysis method, which gave a LNPs yield of 81.8%, 90.9% and 41.0% with a corresponding average particle size of 218, 131, and 104 nm, respectively. The obtained “high-quality” LNPs were in sphere-like shapes, abundant with functional groups, and highly stable from pH 4 to 10, which showed tremendous promise for the applications in the emerging nanomaterial fields. When the substructures of these three LNPs were further characterized using prevalent 13C and 2D-HSQC NMR techniques, they showed that their structure recalcitrance followed the order of lodgepole pine LNPs > poplar LNPs > corn stover LNPs. It was also apparent that the biomass lignin condensation occurring during steam pretreatment could be considered as a “hydrophobic modification”, which benefits the self-assembling of LNPs to small particle sizes and regular shapes. KEYWORDS: Lignin nanoparticles, Enzymatic hydrolysis, Condensation, Biorefinery
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INTRODUCTION Biorefinery, serving as a potential alternative to the traditional oil-based refinery, means the production of a wide range of biobased fuels, chemicals, and materials from renewable lignocellulosic biomass.1,2 Converting lignin into value-added products provides an additional revenue for this nascent biorefinery concept. However, unlike the prevalent bioconversion routes for cellulose and hemicellulose utilization where various hydrolytic enzymes have been employed to breakdown these polysaccharides into a sugar platform for fuels/chemicals production,3 upgrading lignin to a usable form is much more challenging due to its complexity and random structures given by both the nature of biomass species and the pretreatment methods applied.4 The nature of lignin is a highly branched, three-dimensional polymer derived from three phenylpropane units (monolignols), namely, guaiacyl (G, conniferyl alcohol), syringyl (S, sinapyl alcohol), and p-hydroxyphenyl (H, p-coumaryl alcohol).1 The structure and composition of lignin is known to be very dependent on the source of lignocellulosic biomass. For example, softwood lignins are primarily composed of G units, whereas hardwood lignins are dominant in G and S units, and grass lignins contain all three monolignols.5 Besides the © 2017 American Chemical Society
complexity of the nature of the biomass lignin structure, the pretreatment technologies currently applied to “open-up” the biomass structure in many cases further increase the structure intricacy and inhomogeneity of the biomass lignin,6 thereby adding extra challenges for its downstream utilization.5 The currently reported possible conversion routes for lignin valorization often require some unique lignin properties.7,8 For example, breaking the lignin down to produce platform chemicals usually requires low molecular weight lignins with high β−O−4′ content, while the high reactivity of C−H bonds on the aromatic ring and/or the high content of aliphatic hydroxyl groups are favored when lignins were used as material precursors to produce biobased material such as resins (phenol formaldehyde and polyurethane respectively).9−11 However, it has been challenging to obtain these “high-value” lignin feedstock from pretreated biomass due to the extensive lignin fragmentation (through the cleavage of β−O−4′ bonds) and condensation (through the free radical-induced polymerization Received: December 13, 2016 Revised: January 10, 2017 Published: February 2, 2017 2702
DOI: 10.1021/acssuschemeng.6b03043 ACS Sustainable Chem. Eng. 2017, 5, 2702−2710
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Figure 1. Flowchart of integrating lignin nanoparticles production into lignocellulosic biorefinery. Enzymatic hydrolysis was conducted for 72 h at a solids loading of 2% w/v using Cellic Ctec3 (40 mgenzyme g−1glucan). Lignin extraction was conducted at a very mild condition (2% w/v, 3h, 80 °C) by DMSO before subsequent dialysis at room temperature. The dialysis was stopped until no DMSO trace was checked in the wastewater.
lignin (CEL: lignin residue after enzymatic hydrolysis), their valorization would be one of the determinant factors for the economically feasible cellulosic bioethanol production. The current biochemically based second generation biofuel plants have been using a variation of a steam pretreatment process.23 Although steam pretreatment could greatly “open-up” the biomass structure and improve substrate hydrolyzability toward cellulase enzymes, it also caused lignin condensation which, as mentioned above, is the major technical barrier for further lignin valorization.23−25 However, this condensed lignin (greatly enhanced lignin hydrophobicity) might be a good candidate for creating high-quality LNPs since the condensation occurred among aromatics could potentially enhance their hydrophobic aggregation toward formation of the core of lignin nanoparticles even without additional hydrophobic modifications.21 In the work reported here, the technical feasibility of fabricating LNPs from three typical, commercially relevant steam- pretreated lignocellulosic substrates (hardwood, softwood, and agricultural residue) was assessed. The main goal of this study is to extract higher value from the recalcitrant cellulolytic enzyme lignins by integrating LNPs production into current lignocellulosic biorefineries. Furthermore, the characteristics of the obtained LNPs, including morphology, chemical structure, particle size, and stability in acidic and alkaline aqueous conditions were further assessed by prevalent techniques used in lignin chemistry and nanoscience for their downstream use. The relationship between the degree of lignin condensation and the ease of formation of LNPs was also extensively assessed.
among aromatics) happening during the pretreatment processes.12 Several recent studies have shown that producing lignin nanoparticles (LNPs) could be a promising alternative approach for lignin valorization.13−15 Lessons can be learned from the successful production and application of nanomaterials derived from other bioresources such as cellulose, silk, chitosan, starch, etc.16−18 Similar to other natural polymeric nanomaterials, LNPs could play an important role in fabricating engineered nanocomposites as value-added applications in material fields.19,20 Another advantage is that, unlike other techniques for lignin valorization, fabricating LNPs is a very controllable process with the relevant uniform products, even if the starting lignin materials are complex and heterogeneous. Currently, LNPs are mainly produced via a solution-based micellization process where the lignin feedstock has been first dissolved in the selected organic solvent system, and the dissolved lignin then precipitated in a controlled manner (dropwise addition or dialysis) in an antisolvent (usually water) system.21 The mechanism of micellization is generally acknowledged as phase separation due to the amphiphilic property of the lignin. Briefly, similar to a synthetic amphiphilic diblock copolymer, the hydrophobic part of lignin (phenylpropanoid units) aggregates in the water to form the micelle core, while the hydrophilic part of lignin (hydroxyl and carboxyl groups) forms the micelle shell simultaneously.22 Although the direct micellization of some lignin feedstock such as commercial Kraft and organosolv lignin have been achieved with different amounts of success in recent reports,14,15 the authors have claimed that an additional acetylation step prior to micellization is usually required to enhance the hydrophobicity of the starting lignin material if high-quality lignin colloidal nanoparticles products (e.g., smaller particle size, more regular particle shape, higher stability, etc.) are expected.20,21 The lignin “by-products” of the emerging cellulosic bioethanol plants will add to the already enormous pile of underutilized technical lignin derived from the pulp and paper industry.4 Considering the abundance of cellulolytic enzyme
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RESULTS AND DISCUSSION Three industrial relevant, steam-pretreated lignocellulosic substrates, namely, hardwood poplar (POP), softwood lodgepole pine (LPP), and agricultural residue corn stover (CS), were employed in this study to assess the potential of lignin nanoparticle (LNP) production from the enzymatic hydrolysis residues. All the steam pretreatments were executed in 2703
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hydrophilic groups such as the hydroxyl groups associated with carbohydrate “contamination”.21 When the purity of these three LNP samples were assessed, it appeared that, as expected, there was a considerable amount of carbohydrates “contamination” within CS LNPs (∼10%, Table 1), while POP LNPs and LPP LNPs were quite “pure” lignin
compromised conditions to maximize the carbohydrates recovery and to create the substrates with relatively good hydrolyzability.26 The flowchart of LNPs production from these steam-pretreated substrates and the material mass balance at each crucial stage are summarized in Figure 1. As expected, most of the polysaccharides (glucan and xylan) were harvested in the water-soluble fraction after enzymatic hydrolysis, leaving a considerable amount of lignin-rich solid fraction (about half of the original steam-pretreated substrates) as the residue (Figure 1). These hydrolysis residues were usually directly burned as the industrial “waste” in the industrial cellulosic ethanol production process;1 their valorization, therefore, could significantly improve the revenue of the biofuel/biorefinery sector. As dimethyl sulfoxide (DMSO) has a Hildebrand solubility parameter (δ value) close to various types of lignin (high lignin solubility) and can also be easily recycled/recovered at the industrial scale,27 DMSO was selected as the demonstrating lignin extraction solvent for subsequent LNPs production. It was apparent that the solubility of the lignin residue in the DMSO was highly substrate dependent. For example, a large amount of lignin (∼76%) derived from steam-pretreated POP hydrolysis residue was dissolved after solvent extraction (Figure 1a), while only about 37% and 50% of lignin could be extracted from steam-pretreated LPP and CS hydrolysis residue, respectively (Figure 1b and 1c). When the final LNPs production yields were calculated based on the lignin content of the corresponding substrate hydrolysis residues, it appeared that the highest yield of LNP was from steam-pretreated POP (90.9%), followed by CS (81.8%), and LPP (41.0%) (Figure 1). Although previous studies have shown that a considerable amount of biomass lignin condensation could happen during the steam pretreatment process which significantly reduced its value for the further application,28−31 the results reported here indicated that such lignin condensation might not limit the ability to produce LNPs for steam-pretreated substrates. The general properties (e.g., particle morphology and size, residual carbohydrates content, pH stability, etc.) of these LNP products were further examined. Transmission electron microscopy (TEM) images showed that all three LNPs products in water solution were nanosize sphere-like structures, where CS LNPs had the largest spherical particle size, followed by POP LNPs and LPP LNPs (Figure 2). When these LNPs were further freeze-dried and observed under scan electron microscopy (SEM), it was apparent that only CS LNPs suffered a strong aggregation while POP LNPs and LPP LNPs still have their sphere-like morphological structures (Figure S1). This phenomenon indicated that CS LNPs might contain more
Table 1. Residul Carbohydrates Content in Lignin Nanoparticles
a
carbohydrates (%)
POP LNPs
LPP LNPs
CS LNPs
arabinose galactose glucose xylose mannose total
0.29 0.32 0.54 0.48 bdl 1.63
0.28 0.29 0.11 bdla bdl 0.68
1.23 0.63 2.58 6.13 bdl 10.57
bdl: below detection limit.
products (only containing ∼1% carbohydrate). In addition, among the major five biomass sugar components, xylose was the dominant sugar “contamination” within CS LNPs (Table 1), which was likely due to the strong association of corn stover xylan with lignin.32 Recent studies have also indicated that about 20−30% of recalcitrant xylan within the pretreated corn stover could not be hydrolyzed even with extremely high enzyme loading.33 It was also worth noting that although all of the enzymatically hydrolyzed residues contained a considerable amount of glucan (∼30%, Figure 1), their corresponding LNPs exhibited negligible glucose contamination (Table 1). This indicated that the tailored technique route for LNPs production could selectively extract lignin from the biomass enzymatic hydrolysis residues. Since the potential LNPs application might cover a wide range of pH and the pH value is a key variable affecting the particle stability,14 the stability of the LNPs products by dynamic light scattering analysis at various pH was further assessed. The electrical double layer repulsion resulting from the phenolic hydroxyl groups and/or the carboxyl groups were likely the main drivers for stabilizing LNPs;15,21 thus, the zeta potential values of these LNPs (a key indicator of the electrical double layer repulsion) were employed to gain better insights to the dynamic stability of LNP dispersions at various pH. The strong stability of these three LNPs dispersions was observed between pH 4 and 10, as evidenced by the negligible particle size changes (Figure 3a) and the appreciable zeta potential values (Figure 3b) over this pH range. In general, the dynamic light scattering analysis showed that the average particle sizes of CS LNPs, POP LNPs, and LPP LNPs were about 218, 131, and 104 nm, respectively, from pH 4 to 10 (Figure 3a), which was consistent with the previous TEM observations (Figure 2). The zeta potential values of three LNPs were between −20 and −50 mV at the pH value above 4 (Figure 3b), which indicated that these LNPs indeed had appreciable stability at this pH range.15 As also reported before, the effect of pH on the electrical double layer repulsion existed without any clear correlation, which was likely due to the complex protonation and deprotonation reactions occurring on the related hydroxyl groups at various pH.15 Although an increase in particle size was observed at both lower pH (10), the relevant mechanisms behind it might be quite different. For example, at lower pH, LNPs tended to aggregate due to decreased electrical double layer repulsion (Figure 3b) resulting
Figure 2. TEM images of lignin nanoparticles prepared from steampretreated poplar cellulolytic enzyme lignin (a), lodgepole pine cellulolytic enzyme lignin (b), and corn stover cellulolytic enzyme lignin (c). 2704
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The above analysis showed the great potential to valorize these “low-value” cellulolytic enzyme lignin (CEL) residues through the production of “high-quality” LNPs. Earlier work has shown that if a given commercial Kraft lignin substrate was directly subjected to LNPs fabrication using the dialysis method, the corresponding LNPs exhibited quite large particle size (200−500 nm).15 An additional hydrophobic modification of the Kraft lignin (to enhance the hydrophobic aggregation of the lignin fragments) was needed to further reduce the lignin nanoparticle size to around 100 nm.21 However, in the work reported here, it appeared that the direct fabrication of LNPs from the recalcitrant CELs was achieved with decent nanoparticle size (∼100 and 200 nm for woody biomass and agriculture residue, respectively), which was comparable to that of LNPs resulting from hydrophobic modification of Kraft lignin. The enhanced hydrophobicity of the CEL was likely a result of the extensive lignin condensation occurred during steam pretreatment.31 Although the morphology, particle size, and zeta potential are the crucial physical properties of LNPs dispersed in water, further understanding their chemical structure could provide important information to elucidate their formation mechanism and also help to tailor synthetic methodology to produce downstream LNPs-based products. It should be noted that the characterization of these lignin products/materials has been challenging due to their complex and random structure; therefore, a combination of several techniques (e.g., FTIR, 13 C NMR, and 2D-HSQC NMR) was employed in this study to better understand the major chemical structure of these LNP products.34 Due to the high efficiency and easy complement with other sophisticated analysis methods, FTIR analysis was first carried out to acquire some basic information about the chemical structure of these LNPs. A number of absorption peaks shown in the FTIR spectra reflected the structural inhomogeneity and complexity of these LNPs samples (Figure 4), where the hydroxyl groups (strong absorption at 3420 cm−1) were quite abundant in all of the three LNPs. Generally, POP LNPs and CS LNPs were of the HGS (hydroxy−guaiacyl−syringyl)-type lignin with strong bands at 1329 cm−1 (S ring breathing plus G
Figure 3. Effect of pH on particle size (a) and zeta potential (b) of the three lignin nanoparticle samples. The measurement was conducted using the dynamic light scattering method. The standard error was negligible; thus, the error bar is not shown.
from protonation of charged functional groups, while at higher pH LNPs started to disassemble toward dissolution thus showing larger particle size.14,15
Figure 4. FTIR spectra of the fabricated lignin nanoparticles. 2705
DOI: 10.1021/acssuschemeng.6b03043 ACS Sustainable Chem. Eng. 2017, 5, 2702−2710
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Table 2. Assignment and Quantification of Signals of 13C NMR Spectra (Results Expressed as Number of Moieties per Aromatic Ring)
a
chemical shift (ppm)
assignment
POP LNPs
LPP LNPs
CS LNPs
57−54 125−103 140−125 160−140 195−190 176−163 125−103 171−168.5 168.5−166
−OCH3 CAr−H CAr−C CAr−O carbonyl carbon carboxyl carbon degree of condensationa aliphatic −OH phenolic −OH
1.92 2.08 1.49 2.55 0.14 0.40 0.92 0.09 0.04
1.12 1.87 1.71 2.55 0.27 0.52 1.13 0.11 0.08
0.81 2.66 2.05 1.40 0.22 0.70 0.34 0.22 0.22
Calculated from 3.00−I125−103
Figure 5. 2D-HSQC spectra and the main structures of the three LNPs samples. Side-chain linkages (δC/δH 50−90/2.5−6.0): (A) β−O−4′ structures, (A′) γ-acylated β−O−4′ substructures, (B) β−5′ phenylcoumaran substructures, (C) β−β′ resinol substructures, and (C′) γ-acylated β−β′ tetrahydrofuran structures and aromatic units (δC/δH 100−150/5.0−8.5): (G) guaiacyl units, (S) syringyl units, (S′) α-oxidized syringyl units, (PB) p-hydroxybenzoate units, (H) p-hydroxyphenyl units, (FA) ferulate units, and (pCA) p-coumarate. The uncolored cross peaks are signals of sugars or unidentified lignin substructures.
ring substituted in position 5) and 834 cm−1 (C−H out-ofplane in positions 2 and 6 of S units, and in all position of H units), while LPP LNPs showed typical G type lignin bands at 1270 cm−1 (G ring plus C−O stretch), 858 cm−1 (C−H out-ofplane in positions 2 and 5), and 817 cm−1 (C−H out-of-plane in position 6 of G).35 Aromatic C−H in-plane deformations in S units appeared in all of the three LNPs samples but located at
different wavenumbers, which was 1139 cm−1 for LPP LNPs but shifted to 1122 and 1126 cm−1 for POP LNPs and CS LNPs, respectively (Figure 4). This result further supported the higher S units content of POP LNPs and CS LNPs since the shift of aromatic C−H in-plane deformations toward lower wavenumbers indicated an increasing content of S units.36 It was also apparent that both conjugated and unconjugated 2706
DOI: 10.1021/acssuschemeng.6b03043 ACS Sustainable Chem. Eng. 2017, 5, 2702−2710
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ACS Sustainable Chemistry & Engineering carboxyl groups (CO stretching, 1696 and 1633 cm−1, respectively) were presented in the CS LNPs sample, while conjugated carboxyl groups (1650 cm−1) were predominant in the POP LNPs and LPP LNPs samples.35 Further analysis of the location and quantification of the hydroxyl groups in LNPs using the prevalent quantitative 31P NMR technique failed due to the poor solubility of these LNPs in the testing solvent (anhydrous pyridine and deuterated chloroform, 1.6:1.0, v/v), even with the supplementation of the cosolvent (N,N-dimethylformamide).37 This phenomenon indicated that the chemical structure of these LNPs was indeed quite recalcitrant especially when compared to that of representative Kraft and organosolv lignins, which are usually soluble in the above solvent system for the 31P NMR test.37 When the three LNPs samples were analyzed by quantitative 13 C NMR, it was apparent that the three spectra were quite different for their locations and intensities (Figure S2), which indicated that the wide structural differences existed among these LNPs. When the moieties of the major functional groups were further calculated (expressed as number of moieties per aromatic ring, Ar in Table 2),38,39 it appeared that the three LNP products had a much lower content of total hydroxyl groups (aliphatic −OH and phenolic −OH) compared to that of other representative milled wood lignins.40,41 According to the content of protonated aromatics (CAr−H) along with the corresponding methoxyl groups (−OCH3) in each LNPs sample, it could be deduced that the S and G units were the predominant building units for POP LNPs and LPP LNPs, respectively, while H units were much more abundant in CS LNPs (Table 2), which was also consistent with the previous FTIR results (Figure 4). In addition, the major ether linkage β−O−4′ within the three biomasses was likely cleaved to various extents during steam pretreatment, and their corresponding LNPs products were heavily condensed especially for LPP LNPs (degree of condensation, 1.13), followed by POP and CS LNPs (0.92 and 0.34, respectively) (Table 2). When the main linkages and substructure units of these LNPs were identified by using 2D-HSQC NMR (Figure 5 and Table 3, assigned 13C−1H correlation signals in the 2D-HSQC
recalcitrance. In addition, the oxidation at the α-carbon on the S units was observed for POP LNPs (S′ substructure, Figure 5). These three LNPs also contained a certain amount of carboxylic acid or carboxylate ester in different substructures, where it was mainly located at the α-carbon on the H units for POP LNPs (carboxylate ester, PB substructures), at the γcarbon on the G units for LPP LNPs (carboxylic acid, FA substructures), and at the γ-carbon on the G and H units for CS LNPs (FA and pCA substructures). These results were consistent with previous FTIR and 13C NMR analyses (Figure 4 and Table 2). From the analysis above, it seemed like the structure recalcitrance of the three LNPs have followed the order of LPP LNPs > POP LNPs > CS LNPs. Even though various extents of lignin condensation were accompanied by the cleavage of β− O−4′ linkages during the steam pretreament process, as hypothesized earlier, these condensed lignin structures helped the LNPs production likely due to their increased hydrophobicity. More specifically, when the LNPs particle size was plotted against its degree of condensation as well as its content of β−O−4′ linkages, it was apparent that the lignin condensation was favorable to the small-size LNPs formulation (R2 = 0.998), while had a reverse correlation with the β−O−4′ content (R2 = 0.964) (Figure 6). Earlier work has suggested
Table 3. Major Covalent Bond Content Quantified by 2DHSQC Spectra (Results Expressed as Number of Moieties per 100 Aromatic Rings) sample
β−O−4′
β−β′
β−5′
S/G
POP LNPs LPP LNPs CS LNPs
22.9 3.5 57.5
5.7 5.6 25.3
4.6 8.5 5.7
1.7
