Better explicating the strengths and shortcomings of these models will help refine projections and improve transparency in the years ahead.
Image credit: Witsawat.S.
Edited by Alexis T. Bell, University of California, Berkeley, CA, and approved June 3, 2019 (received for review March 17, 2019)
Significance
Deep eutectic solvents (DESs) have gained increasing attention due to their application-friendly properties, including universal solvating capabilities and wide tunability. Additionally, ease of synthesis and broad availability from inexpensive chemical components could render DESs more versatile solvents for biomass pretreatment, as compared with traditional ionic liquids. Because the long-term success of the biorefinery depends on the development of sustainable processes to convert lignocellulosics into biofuels, DESs derived from renewable sources such as lignin are highly desirable. We herein present our innovative process that integrates the use of low-recalcitrant engineered biomass with its pretreatment using lignin-derived DESs. The promising results described by near-theoretical sugar yield demonstrate the effectiveness of the integrated process, opening up opportunities toward a sustainable and circular bioeconomy.
Abstract
Despite the enormous potential shown by recent biorefineries, the current bioeconomy still encounters multifaceted challenges. To develop a sustainable biorefinery in the future, multidisciplinary research will be essential to tackle technical difficulties. Herein, we leveraged a known plant genetic engineering approach that results in aldehyde-rich lignin via down-regulation of cinnamyl alcohol dehydrogenase (CAD) and disruption of monolignol biosynthesis. We also report on renewable deep eutectic solvents (DESs) synthesized from phenolic aldehydes that can be obtained from CAD mutant biomass. The transgenic Arabidopsis thaliana CAD mutant was pretreated with the DESs and showed a twofold increase in the yield of fermentable sugars compared with wild type (WT) upon enzymatic saccharification. Integrated use of low-recalcitrance engineered biomass, characterized by its aldehyde-type lignin subunits, in combination with a DES-based pretreatment, was found to be an effective approach for producing a high yield of sugars typically used for cellulosic biofuels and biobased chemicals. This study demonstrates that integration of renewable DES with plant genetic engineering is a promising strategy in developing a closed-loop process.
The modern lignocellulosic biorefinery strives to develop new processes and products to achieve a sustainable energy future. Although renewable fuels from lignocellulosic biomass have proven to be alternatives to fossil fuels, innovative technologies are still required to build economically viable processes for converting biomass to fuels, chemicals, and materials (1). Recent efforts to overcome such barriers include (i) developing a feedstock-agnostic biomass pretreatment, (ii) engineering microorganisms that can catabolize both monosaccharides and lignin, and (iii) understanding biosynthesis of plant cell walls to develop engineered biomass with improved properties for biofuels and bioproducts. In lignocellulosic biomass-to-ethanol processes, researchers have endeavored to develop a biocompatible and scalable biomass pretreatment process, design new microbial strains that can convert both pentose and hexose with enhanced resistance to inhibitors, and engineer feedstocks to provide high yields of sugars and readily processable lignin.
In contrast to first-generation ethanol, which has been studied in great depth and is considered to be mature, the production of cellulosic ethanol from lignocellulosic biomass still requires overcoming technical and economic hurdles. In particular, lignin represents one of the primary factors contributing to the recalcitrance of biomass as its presence restricts enzymatic hydrolysis by nonproductive binding enzymes (2). Although recent studies have unlocked lignin’s potential for various applications (3), it is still one of the most challenging biopolymers to work with because of its inherent recalcitrant structural characteristics. In this regard, reducing the total amount of lignin in lignocellulosic biomass has been a widely adopted strategy to improve saccharification and the extractability of biomass components (4). In addition, there have been many recent efforts to alter lignin monomeric composition in plants to render lignin more amenable to extraction or chemical depolymerization. For example, by targeting the monolignol biosynthetic pathway (Fig. 1), genetic down-regulation of caffeic acid 3-O-methyltransferase (COMT) (5), hydroxycinnamoyl-CoA shikimate hydroxycinnamoyl transferase (HCT) (6), cinnamoyl-CoA reductase (CCR) (7), ferulate 5-hydroxylase (F5H) (8, 9), caffeoyl shikimate esterase (CSE) (10), and cinnamyl alcohol dehydrogenase (CAD) (11) alter lignin content and/or composition, thus reducing biomass recalcitrance and resulting in an enhancement of saccharification efficiency. Recently, introducing an exotic feruloyl-CoA monolignol transferase (FMT) gene in poplar resulted in the production of monolignol ferulate ester conjugates that are incorporated into lignin (“Zip lignin”) (12). Structurally altered lignin with ester linkages in transgenics was found to be less recalcitrant, which led to improved cell wall digestibility and liberated more monosaccharides from biomass compared with the wild type (WT) (12, 13). Such strategic modification of lignin structure can certainly enhance its overall processability by lowering the energy and chemicals required for biomass conversion (13). Herein, a previously characterized CAD-deficient transgenic Arabidopsis line was used as a feedstock. CAD is an enzyme that catalyzes the last step of the lignin monomer biosynthetic pathway, converting p-coumaryl aldehyde, coniferyl aldehyde, and sinapyl aldehyde to p-coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol, respectively (Fig. 1). Thus, in a CAD mutant expression system, cinnamaldehydes in the cell wall participate in the lignification process instead of conventional monolignols, producing mainly coniferyl aldehyde- and sinapyl aldehyde-derived lignin units (14).
Fig. 1.
The main pathway involved in monolignol biosynthesis. The three monolignol precursors shown with a gray background accumulate upon down-regulation of the CAD genes. PAL, phenylalanine ammonia lyase; C4H, cinnamate 4-hydroxylase; 4CL, 4-coumarate-CoA ligase; HCT, hydroxycinnamoyl-CoA shikimate hydroxycinnamoyl transferase; C3H, coumarate 3-hydroxylase; CSE, caffeoyl shikimate esterase; CCR, cinnamoyl-CoA reductase; CCOMT, caffeoyl-CoA O-methyltransferase; F5H, ferulate 5-hydroxylase; COMT, caffeic acid O-methyltransferase; CAD, cinnamyl alcohol dehydrogenase.
Recently, deep eutectic solvents (DESs) have been introduced to various biomass processes. DES is a solvent system formed from a eutectic mixture of Lewis or Brønsted acids and bases (15). Like ionic liquids (ILs), DESs exhibit desirable chemical and physical properties such as low vapor pressure, tunability, and high thermal stability (16). Also, choline chloride (ChCl)-based DESs have drawn significant attention due to their ability to solvate a wide range of compounds including metal oxides (17), CO2 (18), and lignin (19, 20). Furthermore, DESs are very simple to synthesize, biodegradable, and cheaper to prepare compared with conventional ILs (15, 21). Although ChCl-based DESs are typically hygroscopic, they are relatively unreactive with water, making DESs versatile solvents (22).
DESs are typically prepared from the complexation of quaternary ammonium salts (e.g., ChCl) and hydrogen bond donors (HBDs) including amide (e.g., urea), alcohols (e.g., glycerol), and carboxylic acids (e.g., lactic acid and oxalic acid). ChCl has been widely used as a hydrogen bond acceptor due to its capability of forming an intermolecular hydrogen bond, biocompatibility, and relative inexpensiveness (23, 24). It is also noteworthy that ChCl can be produced biologically, although the current industrial production of ChCl is via reaction of trimethylamine hydrochloride with ethylene oxide (25). DESs have been widely employed as reagents for biomass pretreatment (19, 26⇓–28). More recently, renewable DESs have been prepared from lignin-derived phenolic compounds and proven to be effective in biomass pretreatment, especially when pretreated with p-coumaric acid-derived DES (24). More importantly, a closed-loop biorefinery concept was also proposed by using lignin-derived DESs (24).
In this work, we report the integration of renewable DESs with biomass genetic engineering to move closer toward achieving a closed-loop biorefinery (Fig. 2). Previously, Socha et al. (29) described a closed-loop process using ILs synthesized with lignin and hemicellulose derived aromatic aldehydes. Down-regulation of CAD genes results in the formation of an abnormal and potentially valuable aldehyde-rich lignin (4, 14). As a proof-of-concept, biomass from CAD–down-regulated plants is pretreated with DESs derived from phenolic aldehydes. Then, the residual lignin recovered after pretreatment can be further processed to extract phenolic aldehydes for the production of renewable biomass-based DESs. The synthesized DESs can refill the pretreatment reactor while released monomeric sugars can be converted into biofuels and other value-added products. It is believed that the integrated strategy introduced in the present work could provide future biorefineries with a closed-loop process to achieve a sustainable bioeconomy.
Fig. 2.
A closed-loop biorefinery that can be achieved by integration of renewable deep eutectic solvent (DES) from lignin-derived phenolic aldehydes with engineered biomass.
Results
NMR Analysis of Lignin Structures in WT and CAD Mutant Biomass.
We first characterized the lignin structures in Arabidopsis WT and the CAD double mutant (cad-c cad-d) (30) using 2D heteronuclear single-quantum coherence spectroscopy (HSQC) NMR analysis. Fig. 3 shows partial HSQC spectra of lignins extracted from the WT and CAD transgenic line. The HSQC spectra were divided broadly into aromatic (δH/δC 6.0–8.0/90–160), aliphatic (δH/δC 2.5–6.0/50–90), and aldehyde regions (δH/δC 9.0-10.0/184-196) and analyzed separately. In the aromatic regions of the lignin from the WT (Fig. 3A), typical guaiacyl (G) and syringyl (S) units and cinnamyl alcohol-end groups (substructure X1) were observed. In addition, naturally occurring aldehyde units [cinnamaldehydes (X2) and benzaldehydes (X3)] were also present in the WT lignin, although signal intensities were relatively lower. In contrast, the CAD mutant showed a very low level of typical G/S lignins (Fig. 3B). Instead, new signals from uncommon guaiacyl (G′) and syringyl (S′) units derived from polymerization of coniferyl aldehyde and sinapyl aldehyde were observed, respectively (31, 32). Also, correlations from cinnamaldehydes (X2) and benzaldehydes (X3) appeared in the spectra from the CAD mutant. The recession of the normal G and S units is obvious and the signals from G′ and S′ units are dominant, which is also demonstrated by the semiquantitative analysis. As can be seen in Fig. 3B, the HSQC contour intensity ratio of the typical G and S units from the mutant only accounts for 11% of the total lignin aromatics observed, while G′ and S′ account for 80%. The S/G ratio in the WT was determined to be 0.31, which was nearly identical to the S′/G′ and (S + S′)/(G + G′) ratios in the CAD mutant.
Fig. 3.
HSQC spectra of isolated lignins from Arabidopsis WT (A) and transgenic CAD mutant (B). Main structures of conventional lignin subunits (C) and unique lignin subunits from CAD mutant (D).
In the aliphatic regions of the WT, as expected, typical lignin side-chain hydrogen and carbon resonances were identified. This includes β–O–4 aryl ether (I), β–5 phenylcoumaran (II), and β–β resinol (III) structures. As for the CAD mutant lignin isolates, the HSQC spectra differed markedly from the WT lignin. The common lignin interunit linkages such as β–O–4, β–5, and β–β (I, II, and III) were not detected. The lower molecular weight of lignin in the CAD mutant compared with that of the WT (SI Appendix, Fig. S1) can be explained by this observation. Interestingly, some unique aldehyde signals derived from the polymerization of hydroxycinnamaldehyde appeared. As previously reported, the presence of the aldehydes-derived 8–O–4 (I′), 8–5 (II′), and 8–8 (III′) substructures are evident from the analysis of the HSQC spectra (31, 32). The application of NMR spectroscopy clearly provides useful structural information on lignins from both the WT and CAD mutant.
DES Synthesis from Lignin-Derived Phenolic Aldehydes.
As an initial screen test, vanillin (VAN) and 4-hydroxybenzaldehyde (HBA) were tested as an HBD at three different molar ratios (2:1, 1:1, and 1:2) with ChCl. As shown in SI Appendix, Fig. S2, both ChCl-VAN and ChCl-HBA formed clear and homogeneous DESs at 1:2 molar ratio, respectively. A representation of the possible complex of the ChCl-VAN and ChCl-HBA is given in Scheme 1. A strong hydrogen bond between ChCl and phenolic aldehyde formed under the synthesis conditions, which resulted in the depressed melting point of the mixture (24). It has been demonstrated that HBDs capable of forming strong hydrogen bonds with a chloride ion can exhibit a significant depression of the melting point (33). As shown in Table 1, the eutectic point of each DES is significantly reduced. The decrease in the melting point of ChCl-HBA was more dramatic than that of ChCl-VAN. It is believed that the phenolic hydroxyl group plays a crucial role in forming a strong hydrogen bond between ChCl and the phenolic compound for DES formation. Based on the computational analysis, a chloride ion tends to strongly interact with the phenolic hydroxyl group upon the DES synthesis (SI Appendix, Fig. S3). The presence of a methoxy group in the ortho position was previously reported to interrupt the formation of a DES by steric hindrance (24), which resulted in relatively less depression of melting point of the eutectic mixture.
Scheme. 1.
Possible complex formation in ChCl-VAN and ChCl-HBA.
View this table:
Table 1.
Two DESs used for biomass pretreatment
Biomass Pretreatment Using DESs and Enzymatic Saccharification.
Two DESs synthesized from lignin-derived phenolic aldehydes were tested for biomass pretreatment. The pretreatment of WT and CAD mutant was conducted at 80 °C, which is a relatively mild condition considering that the organosolv pretreatment of biomass is typically carried out at 100–250 °C (34). In addition to the efficiency of DESs as pretreatment solvents, the impact of mild DES pretreatment severity on plant cell wall modifications is discussed.
After pretreatment, the residual DES was washed out, and the solid was recovered for compositional analysis to determine the efficacy of each DES in terms of lignin removal. Compositional analysis of pretreated samples indicates that DES pretreatment of the CAD mutant removed a substantial amount of lignin (SI Appendix, Table S1). Lignin removals from WT biomass were ∼5% and 10% when pretreated with ChCl-VAN and ChCl-HBA, respectively, and those from CAD mutant significantly increased to 25% and 32%, respectively. Previously, the lignin removal was reported to be 6.5% and 10.1% from corncob when pretreated with ChCl-glycerol and ChCl-urea at 80 °C, respectively (26). Clearly, more lignin was removed from biomass of the CAD mutant compared with WT, regardless of the DES used for pretreatment.
It has been reported that lignins from CAD-deficient biomass are more readily solubilized and extracted in alkaline conditions due to their higher content of free-phenolic end groups (11, 35, 36). Based on the principles of hydrogen bond interaction, DES could provide a mild acid-base catalysis mechanism that initiates the cleavage of aryl-ether linkages, leading to lignin separation from the biomass (20). Also, lignin removal by DESs is possibly correlated to the hydrogen bond basicity of the constituent part (29, 37). In addition, gel permeation chromatography (GPC) analysis determined that the lignin macromolecule in the CAD mutant sample has lower molecular weight compared with the WT (SI Appendix, Fig. S1). This also suggests that the reduced degree of polymerization of lignins from CAD mutant could render them more prone to solubilization in DES (38), resulting in greater removal during pretreatment.
Interestingly, in both samples, ChCl-HBA showed higher pretreatment performance regarding lignin removal than ChCl-VAN. It is speculated that the interaction between ChCl-HBA and lignin is relatively stronger than ChCl-VAN and lignin complexes, which led to higher lignin removal from biomass. The pretreated biomass was then subjected to enzymatic hydrolysis to analyze the digestibility of polysaccharides. Fig. 4 shows the sugar yields after ChCl-VAN and ChCl-HBA pretreatment of biomass from both WT and CAD mutant followed by enzymatic hydrolysis. A significant increase in saccharification yield after DES pretreatment was apparent in transgenic biomass. Yields of glucose released from WT were 10.0 and 10.5 wt% (based on initial dried biomass) when pretreated with ChCl-VAN and ChCl-HBA, respectively. Regardless of the DESs employed, glucose yields almost doubled in the case of the CAD mutant. The glucose yield from biomass of the transgenic plant was 19.4 and 20.8 wt% with ChCl-VAN and ChCl-HBA, respectively. The higher digestibility of the CAD mutant results from the strategic down-regulation of CAD genes involved in lignin biosynthesis. This result is in accordance with previous saccharification results obtained from CAD–down-regulated poplar (+39.6–52.0% glucose and +34.2–63.8% xylose after 62.5 mM NaOH pretreatment at 90 °C) (11), switchgrass (+15.0–35.0% glucose after 1.5% H2SO4 pretreatment at 121 °C) (39), alfalfa (+16.0% total sugars after 1.5% H2SO4 pretreatment at 130 °C) (40), and Brachypodium (+44.0–46.0% total sugars after 0.5 M NaOH pretreatment at 90 °C) (41). Clearly, the use of DESs as reagents for biomass pretreatment under mild temperature resulted in comparable amount of sugar without using any conventional acid or base catalyst. Glucan conversions were also calculated based on the biomass compositional analysis (SI Appendix, Table S1). Taking into account the slightly higher glucan content in the raw CAD mutant biomass (∼11.7%), the glucan conversion to glucose was significantly higher from CAD mutant compared with the WT (70% vs. 40% with ChCl-HBA and 66% vs. 38% with ChCl-VAN). It is noted that the enzyme mixture used in this work primarily consists of cellulase, and that overall xylose yield was relatively low. However, the yields of xylose from CAD mutant were considerably higher than those from WT. When pretreated with ChCl-VAN, the xylose yield increased from 2.0% (WT) to 7.9% (CAD mutant). Also, the pretreatment with ChCl-HBA yielded 8.5% xylose from the transgenic biomass, which is more than fourfold increase compared with WT. The discrepancy in xylose yield could be attributed to the enhanced enzyme accessibility to xylan chains in the CAD mutant because of higher lignin removal by DES-assisted pretreatment.
Fig. 4.
Sugar yield from raw biomass (A) and glucan conversion (B) for WT and CAD mutant pretreated with ChCl-VAN and ChCl-HBA, respectively.
The Fate of Aldehyde-Rich Lignin.
The CAD mutant released more fermentable sugars than the corresponding WT control when pretreated with ChCl-VAN and ChCl-HBA, which could be attributed to its modified lignin. Thus, it was hypothesized that DESs effectively remove CAD mutant lignin from the whole biomass structure during pretreatment. To prove this hypothesis, solid residues recovered after pretreatment followed by enzymatic hydrolysis were characterized by 2D HSQC NMR analysis. As shown in SI Appendix, Fig. S4, each residual solid obtained from the 2 DES pretreatments did not show peaks of phenolic aldehydes, which were observed in the raw material. SI Appendix, Fig. S4 shows 2D HSQC NMR spectra of the residual lignin obtained from the 2 DES pretreatments. Although the lignin residue from the ChCl-HBA pretreatment shows weak correlations from benzaldehydes (X3, δH/δC 9.7/191), most of the cross-peaks from aldehydes disappeared. This indicates that DESs effectively removed aldehyde-derived lignin substructures during pretreatment. Also, it reveals that the structures in the aldehyde-rich lignin from the CAD mutant are more chemically reactive than those found in normal lignin. To further confirm this hypothesis, density functional theory (DFT)-based computational study was conducted to calculate a kinetic quantity of lignin structure (29). Model dimers typically found in lignin from WT (β–O–4) and CAD mutant (8–O–4) were used to understand the chemical reactivity. Fig. 5 illustrates the optimized geometry and electrophilicity index of β–O–4 and 8–O–4 dilignols. An electrophilicity index, which determines a quantitative classification of the global electrophilic nature of a molecule (42), was calculated by the global hardness and electronic chemical potential. As shown in this figure, the electrophilicity index of β–O–4 is 1.88 eV, whereas that of the 8–O–4 model is 3.00 eV. It is obvious that 8–O–4 dimer has a higher electrophilicity index, indicating that this structure is chemically more reactive than the normal β–O–4 structure (43). The results from HSQC NMR analysis and computational study support the hypothesis that lignin in CAD mutant biomass is more amenable to processing, thus being readily removed by DES-based pretreatment in comparison with WT.
Fig. 5.
Optimized geometry of β–O–4 dimeric model compounds from WT (A), 8–O–4 dimeric model compounds from CAD mutant (B), and electrophilicity index as a descriptor of chemical reactivity.
Recovery of Phenolic Aldehydes.
As hypothesized earlier, DESs were successfully synthesized using phenolic aldehydes and proved to be effective for biomass pretreatment. Another experiment was carried out to demonstrate that lignin from CAD-deficient plants can supply more phenolic aldehydes for designing DESs. Simply, both WT and CAD mutant biomass were hydrothermally depolymerized at 200 °C without catalyst. The resulting liquid fraction was analyzed by gas chromatography and three primary phenolic aldehydes were identified and quantified (see SI Appendix, Fig. S5 for gas chromatograms). Fig. 6 shows the amount of three phenolic aldehydes obtained from WT and CAD mutant. As shown, the overall yield of phenolic aldehydes produced from transgenic biomass was significantly higher than that from WT (+317%). For individual compounds, the yield of vanillin was 1,345 µg/g lignin from WT, while the CAD mutant yielded 4,142 µg/g lignin. Additionally, CAD mutant produced over 3,000 µg/g lignin of syringyl aldehyde and coniferyl aldehyde, respectively, whereas WT biomass yielded only a small amount of those phenolic aldehydes. A simple hydrothermal cracking of lignin that has aldehyde-rich structures (i.e., as in CAD-deficient plants) was found to be effective at extracting phenolic aldehydes, although more in-depth tests are required to find optimal reaction conditions to maximize yields.
Fig. 6.
The amount of phenolic aldehydes released from hydrothermal depolymerization of WT and CAD mutant lignin.
Discussion
Altering the fundamental composition of lignin has proven to be effective to make biomass more processable during pretreatments. The strategic down-regulation of CAD during lignification results in aldehyde-rich lignin units. As observed from the HSQC NMR analysis, the lignin extracted from CAD mutant shows a significant amount of aldehyde-derived substructures (8–O–4, 8–5, and 8–8) and cinnamaldehyde- and benzaldehyde-end units. In this work, VAN and HBA were used as representative phenolic aldehydes because the CAD mutant has potential for producing such compounds upon depolymerization. The pretreatment of WT and CAD mutant using lignin-derived DESs revealed that lignin removal was more significant from biomass of CAD-deficient plants, which resulted in higher sugar release upon enzymatic saccharification.
The improved sugar yield obtained from CAD transgenic biomass is attributed to several factors, which include (i) the slight decrease in total lignin amount in biomass cell walls reducing the overall physical barrier; (ii) significant structural modification in lignin that may loosen the interactions between lignin and carbohydrates, thus enhancing the accessibility of enzymes (44⇓–46); and (iii) the higher reactivity of aldehyde-rich units in lignin that are chemically more amenable. Considering that only a modest reduction of total lignin content was observed in the CAD transgenic biomass, the effect associated with lignin content is likely to be marginal. Instead, the higher concentration of phenolic aldehydes due to CAD down-regulation reduces the interconnectivity within the macromolecular structure (45). Also, altering lignin composition and the corresponding lignin structures with increased reactivity are likely to reduce cell wall recalcitrance and to result in higher sugar yield.
DESs synthesized from phenolic aldehydes were found to be effective at biomass pretreatment. Interestingly, ChCl-HBA was more effective than ChCl-VAN for lignin removal during pretreatment and sugar release upon saccharification, which is likely attributed to the strong intermolecular interaction in ChCl-HBA compared with ChCl-VAN. Although more supporting parameters (e.g., Kamlet–Taft solvent parameters) are required to compare the pretreatment efficiency of DESs, a better pretreatment performance is anticipated for ChCl-HBA considering its larger depression of melting point compared with that from ChCl-VAN. The strategic down-regulation of CAD during lignification produced biomass with aldehyde-rich lignin, which has the potential to serve as feedstock for the production of aldehyde-derived DESs or other value-added chemicals. The synthesized DESs from phenolic aldehydes proved to be promising solvents for biomass pretreatment resulting in a high yield of fermentable sugars under mild conditions.
There have been reports on enhanced saccharification from CAD mutant biomasses (11, 39⇓–41). It is challenging to directly compare the results from these different studies with the present study because the biomass is derived from different plant species that have various levels of both lignin and CAD deficiency. However, it would be interesting to test and scale-up DES-based pretreatment process with biomass from CAD-deficient bioenergy crops such as poplar (11), bm1 maize (47), bm6 sorghum (48), and switchgrass (49). It is worth noting that, unlike certain mutants affected in other steps of the lignin biosynthetic pathway, several CAD mutants do not show any visible phenotype or biomass yield penalty. This is the case for greenhouse-grown switchgrass (39, 49) and field-grown corn (50, 51), poplar (52), and sorghum (53), although genotype-dependent variations have been observed for the latter (54, 55). It is also important that DES-based catalyst-free pretreatment of transgenic biomass could lower chemical use and energy inputs typically required for the production of intermediate sugars, which can be further processed into value-added products. Although the computational study proved higher reactivity of CAD lignin, more in-depth studies are necessary to obtain a more comprehensive understanding of the effect of compositional alteration of lignin. In actual chemical systems, many atoms concertedly interact with other compounds, contributing to the stability of the whole reaction system (56). Also, the effect of a solvent can be significant in electrophile interactions. Despite some technical challenges, this study clearly demonstrates the potential of developing sustainable biorefinery benefiting from looped production of renewable DESs.
Taken together, down-regulation of CAD in biomass produces lignin with aldehyde-rich units. This increases the overall saccharification efficiency and gives the potential for producing phenolic aldehyde-derived DESs, which can be used for biomass pretreatment. Integration of renewable DESs with biomass genetic engineering is a step closer toward a closed-loop biorefinery and developing a sustainable energy future.
Materials and Methods
Biomass Material.
Arabidopsis thaliana (ecotype Wassilewskija) seeds from WT and CAD double mutant (cad-c and cad-d) (14) were germinated directly on soil. Growing conditions were 14-h light/day at 100 µmol⋅m−2⋅s−1, 22 °C, and 55% humidity. For analyses, stems from mature senesced dried plants harvested without siliques and leaves were ball-milled to a fine powder using a Mixer Mill MM 400 (Retsch) and stainless-steel balls for 2 min. All chemicals used in this work were purchased from Sigma-Aldrich and used without any purification.
DES Synthesis.
DESs used in this work were synthesized using vanillin (ChCl-VAN) and 4-hydroxybenzaldehyde (ChCl-HBA), respectively (Table 1). ChCl and two hydrogen-bonding donor molecules were mixed in the three different molar ratios (2:1, 1:1, and 1:2), respectively, and heated to 100 °C with continuous stirring until a clear liquid was formed. As shown in SI Appendix, Fig. S2, both ChCl-VAN and ChCl-HBA formed a homogeneous DES only at the 1:2 (ChCl:HBD) molar ratio under the synthesis conditions tested in this work. 1H and 13C NMR spectra of VAN, HBA, ChCl, ChCl-VAN, and ChCl-HBA were obtained on an AVANCE III HD 800-MHz instrument in D2O (SI Appendix, Fig. S6).
ChCl-VAN. 1H NMR: 3.22 (s, 9H), 3.54 (t, 2H), 3.87 (m, 6H), 4.09 (m, 2H), 6.97 (m, 2H), 7.34 (m, 2H), 7.44 (m, 2H), 9.62 (s, 2H); 13C NMR: 54.00 (3C), 55.71 (1C), 55.90 (2C), 67.55 (1C), 111.30 (2C), 115.45 (2C), 127.54 (2C), 129.10 (2C), 147.92 (2C), 152.31 (2C), 194.73 (2C).
ChCl-HBA. 1H NMR: 3.22 (s. 9H), 3.54 (t, 2H), 4.07 (m, 2H), 7.03 (m, 4H), 7.88 (m, 4H), 9.73 (s, 2H); 13C NMR: 54.01 (3C), 55.71 (1C), 67.55 (1C), 116.15 (4C), 128.84 (2C), 133.20 (4C), 162.67 (2C), 194.99 (2C).
Pretreatment and Enzymatic Saccharification.
For DES-mediated biomass pretreatment, ∼5 wt% biomass solution was prepared by mixing 0.20 g of biomass with 3.80 g of each DES in a 20-mL pressure tube (Ace Glass). Although typical solids loading in biomass pretreatment is 10 wt% or greater, 5 wt% solids loading was used in this work for the purpose of demonstrating a closed-loop biorefinery. Pretreatment of each biomass with two different DESs was conducted in an oil bath at 80 °C for 3 h. After pretreatment, biomass was transferred to a 15-mL centrifuge tube and washed with 50 mL (10 mL × 5) of an ethanol/DI water mixture [2:1 (vol/vol)] to remove any residual DESs, and the solid fraction was completely dried for digestibility assays. For saccharification of the pretreated biomass, enzymatic hydrolysis was performed using a commercial enzyme blend Cellic CTec2 (Sigma-Aldrich). Briefly, 5 mL of 50 mM citrate buffer (pH 5.0) was added to the pretreated biomass with an enzyme dosage of 10 mg protein per gram biomass. Protein content of the enzyme mixture used in this study was determined from the ninhydrin-based assay (161.5 mg/mL). The hydrolysis was conducted in a rotating hybridization oven at 50 °C for 72 h.
Analytical Methods.
After saccharification, the hydrolysate was separated from the substrate and filtered through a 0.45-μm syringe filter. Glucose and xylose release was measured using a YL 9100 high-performance liquid chromatography (Young-Lin) equipped with a refractive index detector and a Bio-Rad Aminex HPX-87H ion-exchange column. The mobile phase used was 4 mM H2SO4 at a constant flow rate of 0.6 mL/min, and the column temperature was set at 60 °C.
To elucidate the structural characteristics of lignin, a 2D HSQC NMR analysis was conducted with biomass samples. Before the NMR analysis, cellulolytic enzyme lignin (CEL) was isolated as described in the previous study (57). In brief, extractives-free biomass was prepared from the ball-milled samples using Soxhlet extraction with a toluene-ethanol mixture [2:1 (vol/vol)] for 12 h followed by water extraction for an additional 8 h. The extractives-free samples were ball-milled using a Retsch Ball Mill PM 100 in a ZrO2 jar (50-mL internal volume) with 10 ZrO2 balls. The ball milling was conducted at 600 rpm at a frequency of 5 min with 5-min pauses in-between for 2.5 h. The ball-milled samples were treated with a mixture of Cellic CTec2 in the sodium acetate buffer solution (pH 4.8) at 50 °C and 200 rpm for 48 h. The residual solids were centrifuged and treated with fresh enzyme (CTec2) one more time under the same conditions. The residual solids after two-step enzymatic hydrolysis were washed with DI water and freeze-dried. The dried solid residues were extracted with dioxane [96% (vol/vol)] for 24 h. The extracted supernatant was collected and dioxane extraction was repeated. Dioxane and water in the collected supernatant were evaporated by rotary evaporator at 45 °C, and the recovered lignins (CEL) were freeze-dried for the further analysis. The prepared lignin samples were dissolved in a 5-mm NMR tube with DMSO-d6. The NMR analysis was performed using a Bruker Avance III HD 500-MHz spectrometer equipped with a N2 Cryoprobe (BBO 1H and 19F-5 mm) with the following acquisition parameters: spectra width 12 ppm in F2 (1H) dimension with 1,024 time of domain (acquisition time, 85.2 ms), 220 ppm in F1 (13C) dimension with 256 time of domain (acquisition time, 6.1 ms), a 1.0-s delay, a 1JC–H of 145 Hz, and 128 scans. HSQC experiments were carried out with a Bruker pulse sequence (hsqcetgpspsi2.2). Assignment of the HSQC spectra is described elsewhere (11, 30).
Molecular-weight distribution of lignin was analyzed by GPC. The analysis was conducted using the Agilent GPC SECurity 1200 system equipped with three Walters Styragel columns (HR1, HR2, and HR6) and an UV detector (270 nm). The analysis was conducted with tetrahydrofuran as mobile phase at 1.0 mL/min. Polystyrene standards were used for calibration. Polymer Standards Service WinGPC Unity software was used for data collection and processing. All lignin samples were acetylated using acetic anhydride and pyridine [1:1 (vol/vol)] at 60 °C for 2 h before analysis.
Hydrothermal Depolymerization.
Hydrothermal depolymerization of WT and CAD mutant was performed in a 50-mL batch Parr reactor (Parr Instrument Company). For the test, 0.40 g of each biomass sample was placed in the reactor and 25 mL of deionized water was then added. The reactor was sealed, purged, and pressurized to 300 psi with He. The reaction mixture was heated and maintained at 200 °C for 1 h under continuous stirring. After reaction, the reactor was cooled in an ice bath. The resulting solution was transferred into a separatory funnel with ethyl acetate (25 mL) added. The mixture was vigorously mixed, and the organic layer was separated. The water phase was extracted once more with ethyl acetate (25 mL). The combined ethyl acetate fractions were dried over sodium sulfate, and the filtrate was evaporated under reduced pressure. Approximately 50 mg of the products were then dissolved in 2 mL of ethyl acetate and used for gas chromatography (GC) analysis. Identification and quantification of the phenolic aldehydes were conducted using an Agilent 7820A GC equipped with 5975 mass spectrometry detector. The capillary column used was an Agilent HP-Innowax (30 m × 0.25 mm × 0.25 μm). Injection temperature was 250 °C and oven temperature was programmed to hold at 70 °C for 5 min, ramp to 260 °C at 5 °C/min, and then hold for additional 5 min. In this work, three phenolic aldehydes were identified and quantified, namely vanillin, syringaldehyde, and coniferylaldehyde.
Computational Analysis.
Considering the complex structure of lignin, computational studies typically use lignin model compounds to gain mechanistic understanding. Therefore, lignin model compounds with representative interunit linkages, such as a β–O–4 dimer, are widely used because it accounts for 50–80% of whole interunit structures. In this work, a β–O–4 dimer [1-(4-hydroxyphenyl)-2-phenoxypropane-1,3-diol] from normal lignin and the corresponding 8–O–4 dimer [3-(4-hydroxyphenyl)-2-phenoxyacrylaldehyde] from lignin of the CAD mutant were used for mechanistic study. For geometry optimizations, two model dimers were analyzed using DFT with the M06-2X hybrid exchange-correlation functional method and the 6-31+G(2d,2p) basis set in Gaussian 09 program (58). In this work, the DFT-based global reactivity descriptor, electrophilicity index (ω) was calculated using the electronegativity (χ) and the chemical hardness (η) to compare the reactivity of chemical system (29, 43, 56, 59). Equations for the calculation of the electronegativity (χ) and the hardness (η) are given as follows:
where
Acknowledgments
This work is supported, in part, by the Korea Institute of Science and Technology–The University of British Columbia Biorefinery on-site laboratory project. In addition, support was provided in part, by UT-Battelle, LLC, under Contract DE-AC05-00OR22725 with the U.S. Department of Energy. This study was supported and performed, in part, as part of the Center for Bioenergy Innovation (CBI). CBI is a U.S. Department of Energy (DOE) Bioenergy Research Centers supported by the Office of Biological and Environmental Research in the DOE Office of Science. This work was part of the DOE Joint BioEnergy Institute (http://www.jbei.org) supported by the U.S. DOE, Office of Science, Office of Biological and Environmental Research, through Contract DE-AC02-05CH11231 between Lawrence Berkeley National Laboratory and the U.S. DOE. The publisher, by accepting the article for publication, acknowledges that the U.S. Government retains a nonexclusive, paid-up, irrevocable, worldwide license to reproduce the published form of this manuscript, or allow others to do so, for U.S. Government purposes. The DOE will provide public access to these results of federally sponsored research in accord with the DOE Public Access Plan (https://www.energy.gov/downloads/doe-public-access-plan). The views and opinions of the authors expressed herein do not necessarily state or reflect those of the U.S. Government or any agency thereof. The views and opinions of the authors expressed herein do not necessarily state or reflect those of the U.S. Government or any agency thereof. Neither the U.S. Government nor any agency thereof, nor any of their employees, makes any warranty, expressed or implied, or assumes any legal liability or responsibility for the accuracy, completeness, or usefulness of any information, apparatus, product, or process disclosed, or represents that its use would not infringe privately owned rights. We thank Jie Wu of the Department of Wood Science at the University of British Columbia for measuring the protein content. The NIH Shared Instrumentation Grant 1S10OD012254 for the 800-MHz NMR spectrometer is also acknowledged.
Footnotes
- ↵1To whom correspondence may be addressed. Email: kwanghokim{at}kist.re.kr.
Author contributions: K.H.K. and A.E. designed research; K.H.K., A.E., K.J., and C.G.Y. performed research; K.H.K., K.J., C.G.Y., C.S.K., and A.R. analyzed data; and K.H.K., A.E., K.J., C.G.Y., C.S.K., and A.R. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1904636116/-/DCSupplemental.
Published under the PNAS license.
References
Integration of renewable deep eutectic solvents with engineered biomass to achieve a closed-loop biorefinery
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