Article pubs.acs.org/JAFC
Cite This: J. Agric. Food Chem. XXXX, XXX, XXX−XXX
Structural Characterization of Lignins from Willow Bark and Wood Jinze Dou,†,‡ Hoon Kim,‡ Yanding Li,‡ Dharshana Padmakshan,‡ Fengxia Yue,‡ John Ralph,‡ and Tapani Vuorinen*,† †
Department of Bioproducts and Biosystems, School of Chemical Engineering, Aalto University, Post Office Box 16300, FI-00076 Aalto, Finland ‡ Department of Biochemistry and United States Department of Energy Great Lakes Bioenergy Research Center, Wisconsin Energy Institute, University of WisconsinMadison, Madison, Wisconsin 53726, United States Downloaded via UNIV OF SUNDERLAND on July 8, 2018 at 17:33:42 (UTC). See https://pubs.acs.org/sharingguidelines for options on how to legitimately share published articles.
S Supporting Information *
ABSTRACT: Understanding the chemical structure of lignin in willow bark is an indispensable step to design how to separate its fiber bundles. The whole cell wall and enzyme lignin preparations sequentially isolated from ball-milled bark, inner bark, and wood were comparatively investigated by nuclear magnetic resonance (NMR) spectroscopy and three classical degradative methods, i.e., alkaline nitrobenzene oxidation, derivatization followed by reductive cleavage, and analytical thioacidolysis. All results demonstrated that the guaiacyl (G) units were predominant in the willow bark lignin over syringyl (S) and minor phydroxyphenyl (H) units. Moreover, the monomer yields and S/G ratio rose progressively from bark to inner bark and wood, indicating that lignin may be more condensed in bark than in other tissues. Additionally, major interunit linkage substructures (β-aryl ethers, phenylcoumarans, and resinols) together with cinnamyl alcohol end groups were relatively quantitated by twodimensional NMR spectroscopy. Bark and inner bark were rich in pectins and proteins, which were present in large quantities and also in the enzyme lignin preparations. KEYWORDS: willow, bark, lignin, nuclear magnetic resonance spectroscopy, structural characterization
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INTRODUCTION Willow has been investigated over time as a potential crop for integrated biorefinery processes, in addition to its use in heat and power generation.1 Willow species possess several characteristics, such as high productivity, short rotation time, and ability to grow on marginal land, that make them ideal feedstocks for pulping, bioethanol production, and biomedicinal applications.2−4 Previous studies revealed that the highquality fibers and aqueous aromatic extracts from the bark could produce a higher value than burning these residues.5 Therefore, a deeper understanding of the chemical structure of bark lignin and its linking to other cell wall polymers is the key for developing appropriate methods for the separation of sclerenchyma fiber bundles from willow bark as a part of the whole willow biorefinery scheme.5 Lignin is a major natural substance in the plant world and the largest potential source of aromatic compounds.6 Lignins are amorphous polymers arising from enzyme-mediated oxidation of three main precursors, p-coumaryl, coniferyl, and sinapyl alcohols, and subsequent polymerization of the formed phenoxy radicals in a purely chemical fashion.7 These monolignols give rise to p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) phenylpropanoid units when incorporated into the lignin polymer. The dominant types of linkages that connect these units are β−O−4 (β-aryl ether), β−5 (phenylcoumaran), β−β (resinol), 5−5 (biphenyl), and 4−O−5 (biaryl ether), along with low levels of β−1 linkages. The lignin chemical composition and the yields of monomers derived from them by classical degradation methods vary among botanical species, plant tissues, and cell types.8 The monolignol ratio, especially S/G ratio, and the distribution of the linkage © XXXX American Chemical Society
types and functional groups, determine the polymer properties and chemical reactivity of lignin, e.g., in chemical pulping processes.7 Therefore, an improved understanding of the composition and structure of lignin would provide an asset to tailor conditions for processing of bark for various end products. Although the chemical structure of wood lignin has been elucidated recently,9,10 the composition and structure of bark lignin have not been investigated earlier. In this study, whole cell wall (WCW) and enzyme lignin (EL) from bark, inner bark, and wood were prepared and analyzed by the classical degradative methods, including alkaline nitrobenzene oxidation (NBO), derivatization followed by reductive cleavage (DFRC), and thioacidolysis. These wet chemical methods allow for qualitative and quantitative analyses of the monolignol composition, expressed as H/G and S/G ratios. Nuclear magnetic resonance (NMR) spectroscopy can provide supporting and additional information on lignin chemistry and allow for the detection of even new lignin substructures. Therefore, solution-state two-dimensional (2D) NMR spectroscopy of EL and WCW was also performed.11,12
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MATERIALS AND METHODS
Materials. All chemicals and solvents used in this study were purchased from Aldrich (Milwaukee, WI, U.S.A.) and used as supplied. Four-year-old trees of willow hybrid ‘Karin’ were harvested Received: Revised: Accepted: Published: A
April 18, 2018 June 20, 2018 June 22, 2018 June 22, 2018 DOI: 10.1021/acs.jafc.8b02014 J. Agric. Food Chem. XXXX, XXX, XXX−XXX
Article
Journal of Agricultural and Food Chemistry from the plantation at the VTT Technical Research Centre of Finland, Ltd., located in central Finland (Kyyjärvi), on May 13, 2015. The whole stems were stored at −20 °C until use. For separation of bark, the frozen stems were immersed in water at 20 °C overnight before debarking. The bark, inner bark, and wood were manually separated with a scalpel and stored at 20 °C until further use. For the degradative and Klason lignin analyses, the bark, inner bark, and wood fractions were ground for 2 min at 30 Hz using a MM400 mixer mill (Retsch, Newtown, PA, U.S.A.) with corrosion-resistant stainless-steel screw-top grinding jars (50 mL) containing a single steel ball bearing (30 mm). The extractive-free samples were stored at 20 °C. Klason Lignin Determination. The Klason lignin content of bark, inner bark, and wood was determined on the solid residue after consecutive extractions with distilled water (3 times), 80% ethanol (3 times), and acetone (3 times) using ultrasonication. The Klason lignin content was measured as freeze-dried solid material by following the standard method applied on 300 mg of extractive-free sample.13 Klason lignin data are reported as the average of triplicate runs. WCW and EL Preparation. The extractive-free isolated cell walls were finely milled at 600 rpm using a Pulverisette 7 planetary ball mill (Retsch) with zirconium dioxide (ZrO2) vessels (50 mL) and ZrO2 ball bearings (10 mm × 10). Each sample was ground in 23 cycles of 10 min, with 5 min breaks between the cycles. To prepare EL, the polysaccharide content of the ground WCW was reduced by shaking the slurry in pH 5 buffer with Cellulysin crude cellulase (Calbiochem, San Diego, CA, U.S.A.) in an incubator for 72 h at 40 °C. The cellulase treatment was repeated twice. The final slurry was centrifuged at 3000 rpm for 20 min with a high-speed centrifuge. The solid obtained was washed 3 times with distilled water and then freeze-dried for approximately 72 h to obtain the EL preparation.14 NMR Spectroscopy for WCW and EL. NMR spectra were acquired on an AVANCE 700 MHz instrument (Bruker Biospin, Billerica, MA, U.S.A.) equipped with a cryogenically cooled 5 mm QCI 1H/31P/13C/15N cryoprobe with inverse geometry (1H coils closest to the sample). Phase-sensitive 2D 1H−13C heteronuclear single quantum coherence (HSQC) spectra were acquired with spectral widths of 12 ppm for 1H (1682 data points and a F2 acquisition time of 100 ms) and 220 ppm for 13C with 620 increments (F1 acquisition time of 8 ms) using 800 scans, with a 500 ms interscan delay. An adiabatic version of the HSQC experiment was used for both WCW and EL samples (hsqcetgpsisp.2 pulse sequence from the Bruker Library; phase-sensitive gradient-edited 2D HSQC using adiabatic pulses for inversion and refocusing).15 The central dimethyl sulfoxide (DMSO) solvent peak was used as an internal reference (δC, 39.5 ppm; δH, 2.49 ppm). Processing used typical matched Gaussian apodization in the 1H dimension and squared cosine bell apodization in the 13C dimension. The volume integration of contours used TopSpin 3.2 software (Bruker). The image contours were colorized using Adobe Illustrator. Thioacidolysis. Thioacidolysis of the WCW and EL samples was performed according to a published standard procedure.16,17 WCW (5 mg) or EL (2 mg) was mixed with 4.0 mL of freshly prepared thioacidolysis reagent (2.5 mL of ethanethiol + 0.7 mL of BF3 etherate + 21.8 mL of freshly distilled 1,4-dioxane) in a tube fitted with a Teflon-lined screw cap. The cap was screwed on the top not so tightly before settling into a heating block at 100 °C for 4 h with occasional shaking. After cooling with ice water, 1 mg/mL 4,4′ethylidenebisphenol (EBP) in distilled 1,4-dioxane (150 μL for wood and 20 μL for bark) and 1 mg/mL tetracosane (C24) in CH2Cl2 (180 μL for wood and 25 μL for bark) were added as an internal standard (IS). After closing, the vial was vigorously shaken to homogenize the mixture. The mixture was then transferred into a separatory funnel, and 7 mL of 0.4 mol/L NaHCO3 was added to neutralize BF3. After acidifying (pH < 3) the aqueous phase with 2 mL of 1 mol/L HCl, the solution was further extracted with CH2Cl2 (10 mL × 3 times). The combined CH2Cl2 extracts were washed with saturated NH4Cl, dried over anhydrous Na2SO4, and filtered. The filtrate was collected in a 100 mL pear-shaped flask, and the solvent was evaporated under reduced pressure at 40 °C.
For the trimethylsilane (TMS) derivatization, the dry thioacidolysis product mixture was transferred into a gas chromatography (GC) vial using 1 mL of CH2Cl2 (200 μL × 5). N,O-Bis(trimethylsilyl)trifluoroacetamide (BSTFA) (100 μL) was then added to the GC vial and heated at 50 °C for 30 min. The trimethylsilylated products were analyzed by gas chromatography−mass spectroscopy (GC−MS), consisting of a GC2010 instrument connected with a PARVUM2 mass spectrometer (Shimadzu Co., Ltd., Kyoto, Japan). The fusedsilica capillary column used was a 300 × 0.25 mm inner diameter, 0.25 μm, SHR5XLB and identified by comparison to the retention times and mass spectra of the corresponding authentic compounds. GC oven temperature was ramped from 140 °C with a hold of 1 min to 280 °C at 3 °C/min with a hold of 1 min and then raised at 30 °C/ min to 300 °C (held for 10 min), with a total running time of 59.33 min. Retention times: 26.27 min, EBP (IS); 31.92 min, C24 (IS); 33.77 min, H (p-hydroxyphenyl monomer); 36.10 and 36.21 min, G (guaiacyl monomer); and 38.47 and 38.62 min, S (syringyl monomer). The flame ionization detector (FID) response factor (RF) values relative to EBP (IS) (1.46, H; 1.95, G; and 2.03, S) and C24 (IS) (1.06, H; 1.44, G; and 1.51, S) were determined according to earlier literature.17 Quantitative measurements were performed in triplicate applying a FID. Alkaline Nitrobenzene Oxidation (NBO). Nitrobenzene oxidation of WCW and EL samples was performed according to the literature.18 The sample (40 mg) was placed in a steel Parr reactor (10 mL volume) with 2 mol/L NaOH (7 mL) and nitrobenzene (0.4 mL), and then the vessel was heated at 170 °C for 2 h in an oil bath with continuous shaking. As an IS, 1 mL of 5 μmol/mL 3-ethoxy-4hydroxybenzaldehyde (EV) in 0.1 mol/L NaOH was added to the reaction mixture. Then, the vessel was closed again and vigorously shaken to guarantee the homogeneity of the mixture. The reaction mixture were transferred to a 100 mL separatory funnel and washed with CH2Cl2 (15 mL × 3). The remaining aqueous layer was acidified with 2 mol/L HCl, and the oxidation products were extracted with CH2Cl2 (20 mL × 2) and diethyl ether (20 mL). Then, the combined organic layer was washed with deionized water (10 mL), dried over Na2SO4, and filtered. The filtrate was collected in a 100 mL pearshaped flask, and the solvent was evaporated to dryness under reduced pressure. For the TMS derivatization step, the dry product mixture was transferred to a GC vial using pyridine (200 μL × 3). BSTFA (150 μL) was then added, and the vial was heated at 50 °C for 30 min. The trimethylsilylated products were analyzed using GC−MS and identified by comparison to the peak retention times and mass spectra of the authentic compounds. The GC oven temperature program was 10 min at 150 °C, 5 °C/min to 280 °C, and 20 min at 280 °C. The total running time was 56 min. The identified products and their retention times were 3.61 min, p-hydroxybenzaldehyde (Hy); 6.43 min, vanillin (V); 8.16 min, EV (IS); 9.22 min, phydroxybenzoic acid (HA); 12.24 min, syringaldehyde (Sy); 14.40 min, vanillic acid (VA), and 18.25 min, syringic acid (SA). The following FID RF values (relative to EV) were determined experimentally: 0.81, Hy; 0.99, V; 0.79, HA; 1.20, Sy; 1.00, VA; and 1.23, SA. Quantitative measurements were performed in triplicate applying a FID. Derivatization Followed by Reductive Cleavage (DFRC). The DFRC analyses of WCW and EL samples were conducted according to published protocols.19−21 The extracted WCW or EL sample (20 mg) was mixed with acetyl bromide in acetic acid (1:4, v/v, 4.0 mL) in a stoppered flask and heated in a sand bath for 2.5 h at 50 °C. An internal standard (50 μL of 1 mg mL−1 EBP in ethyl acetate) was then added, after which the solvents/reagents were removed under reduced pressure on a rotary evaporator (10 min at 50 °C). The residue was mixed with 5 mL of 1,4-dioxane/acetic acid/water (5:4:1, v/v/v) and 50 mg of zinc nanopowder and stirred at room temperature for 20 h. The solution was then transferred with 4 mL of CH2Cl2 to a 50 mL separatory funnel that contained 15 mL of saturated aqueous ammonium chloride. After shaking, the organic phase was separated and the aqueous layer was extracted repeatedly with CH2Cl2 (4 × 10 mL). The combined organic extracts were dried with anhydrous B
DOI: 10.1021/acs.jafc.8b02014 J. Agric. Food Chem. XXXX, XXX, XXX−XXX
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Journal of Agricultural and Food Chemistry
Table 1. Yields of Lignin-Derived Thioacidolysis Monomers from WCW and EL Preparations of Bark (WB), Inner Bark (WIB), and Wood (WW) Materials, Measured by GC−FID with Two Different IS (EBP and C24) monomer yield (μmol/g of lignin)a lignocellulosic material
IS
WB-WCWc
EBP C24 EBP C24 EBP C24 EBP C24 EBP C24 EBP C24
WIB-WCWc WW-WCWc WB-ELd WIB-ELd WW-ELd
H 2.1 2.0 1.4 1.2 7.2 6.2 1.2 1.6 0.3 0.5 5.4 5.2
G
(0.1) (0.1) (0.1) (0.2) (1.0) (0.8) (0.3) (0.4) (0.04) (0.04) (0.3) (0.2)
98 94 169 147 669 582 40 57 39 58 400 391
monomer ratiob S
(10) (9) (8) (19) (30) (16) (10) (14) (3) (9) (18) (8)
90 87 197 172 1406 1231 34 49 40 59 694 683
(11) (10) (13) (26) (68) (38) (6) (9) (3) (10) (31) (13)
total yield
H/G
S/G
190 182 368 320 2082 1820 76 107 79 117 1100 1079
0.02 0.02 0.01 0.01 0.01 0.01 0.03 0.03 0.01 0.01 0.01 0.01
0.92 0.93 1.16 1.17 2.10 2.12 0.85 0.86 1.01 1.01 1.73 1.74
(21) (19) (21) (45) (99) (54) (16) (23) (6) (19) (48) (21)
a Total yield [standard deviation (SD) in parentheses] of the thioethylated monomers with H, G, and S structure. bThe molar monomer ratio was calculated from the monomer yields. cIn units of μmol/g of Klason lignin. dIn units of μmol/g of enzyme lignin.
Table 2. Yields of Lignin-Derived NBO Monomers from WCW and EL Preparations of Bark, Inner Bark, and Wood Materials, Measured by GC−FID monomer yield (μmol/g of lignin)a lignocellulosic material WB-WCWb WIB-WCWb WW-WCWb WB-ELc WIB-ELc WW-ELc
H 51 68 54 30 24 25
(5) (7) (11) (2) (1) (2)
monomer ratio
G 191 287 962 94 116 681
S
(7) (4) (32) (2) (5) (41)
142 307 1908 70 115 1151
(6) (6) (41) (4) (4) (52)
total yield
H/G
S/G
385 662 2925 194 254 1858
0.27 0.24 0.06 0.32 0.19 0.04
0.74 1.07 1.98 0.74 0.99 1.69
(10) (14) (20) (2) (10) (44)
Total yield (SD in parentheses) of the NBO monomers with H (Hy and HA), G (V and VA), and S (SA and Sy) structure. bIn units of μmol/g of Klason lignin. cIn units of μmol/g of enzyme Lignin. a
Table 3. Yields of Lignin-Derived DFRC Monomers from WCW and EL Preparations of Bark, Inner Bark, and Wood Materials, Measured by GC−FID monomer yield (μmol/g of lignin)a lignocellulosic material
H
WB-WCWb WIB-WCWb WW-WCWb WB-ELc WIB-ELc WW-ELc
0 0 4.5 (0.5) 0 0 4.0 (0.1)
monomer ratio
G 26 48 292 10 11 157
(1) (18) (3) (1) (1) (5)
5.5 11 410 2.6 7.6 255
S
total yield
H/G
S/G
(0.6) (2) (44) (0.1) (0.7) (11)
31 59 706 13 19 416
0.00 0.00 0.02 0.00 0.00 0.03
0.22 0.24 1.41 0.26 0.70 1.63
(2) (20) (43) (1) (0) (16)
Total yield (SD in parentheses) of the DFRC monomers with H, G, and S structure. bIn units of μmol/g of Klason lignin. cIn units of μmol/g of enzyme lignin.
a
sodium sulfate and evaporated to dryness under reduced pressure using the rotary evaporator. The residue was mixed with freshly prepared pyridine/acetic anhydride (1:1, v/v, 4 mL) and kept overnight in the dark for complete acetylation. After evaporation, the oily residue was loaded onto a solid-phase extraction (SPE) catridge (Supelco Supelclean LCSi SPE tube) using CH2Cl2 (3× 1 mL) to remove most of the polysaccharide-derived products and then eluted with hexane/ethyl acetate (1:1, v/v, 8 mL). The DFRC products were separated by GC using the SHR5XLB capillary column. The GC oven temperature was ramped at 3 °C/min from 140 to 240 °C, held for 1 min, raised at 30 °C/min to 300 °C, and held for 12 min, with a total running time of 48.33 min. Retention times of the peracetates were 15.35 min, H (phydroxycinnamyl alcohol); 17.83 and 20.52 min, G (cis- and transconiferyl alcohol, respectively); 22.70 and 25.82 min, S (cis- and transsinapyl alcohol, respectively); and 28.78 min, EBP (IS). Quantitative
measurements were performed in triplicate applying a FID. The following FID RF values (relative to EBP) were used: 1.43, H; 1.37, G; and 1.46, S.19
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RESULTS AND DISCUSSION Klason Lignin Determination. The Klason lignin content of bark (24.7 ± 0.1%) was higher than that of inner bark (18.0 ± 0.1%) and wood (22.4 ± 0.1%). Similar or higher bark lignin contents have been reported for Salix psammophila (31.7%), Salix sacchlinensis (23.8%), and Salix schwerinii (26.5%).22 For bark, Klason lignin may possibly include components other than lignin, even though the samples were pre-extracted with several solvents as described in the widely accepted method.13 Monomer (H, G, and S) Composition of Bark, Inner Bark, and Wood Lignin. WCW and EL preparations of bark, C
DOI: 10.1021/acs.jafc.8b02014 J. Agric. Food Chem. XXXX, XXX, XXX−XXX
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Journal of Agricultural and Food Chemistry
Figure 1. (Top) Aromatic (δC/δH, 96−150/6.0−8.2 ppm) and (bottom) side-chain (δC/δH, 48−92/2.0−6.0 ppm) regions of 2D HSQC NMR spectra of EL preparations from (left) bark, (middle) inner bark, and (right) wood.
hardwood (willow and poplar) lignins were less condensed than the softwood (pine) lignin.17,19 Slightly lower monomer yield values were obtained when C24 was used as the IS instead of EBP, which is consistent with earlier observations.17 Although the yields of thioacidolysis products with the H structure were low in general, the H/G ratio was higher in bark in comparison to wood (Table 1). Similarly, NBO produced H monomers in higher yields from bark in comparison to wood (Table 2). On the contrary, H monomers were not always detected by the DFRC approach, even though the total monomer yield from WCW of wood (706 μmol/g of lignin) was comparable to a previously reported level (732 μmol/g of lignin)19 (Table 3). It has been well-documented that thioacidolysis and DFRC monitor real, authentic H units in lignin, whereas NBO can yield H monomers also from phydroxybenzoates that acylate lignin side chains24 and even from tyrosine.25 Despite the possible sources of error, the real H/G ratio of bark lignin is obviously higher in comparison to that of wood. The systematically lower monomer yields from EL in comparison to WCW (Tables 1−3) may result from the fact that EL does not represent pure lignin, a common problem noted previously in certain samples.25−27 2D NMR Spectroscopy of WCW and EL. To obtain more detailed information on the structure of the lignins, WCW and
inner bark, and wood were characterized for their lignin monomer composition by thioacidolysis, NBO, and DFRC. These degradative methods estimate the relative proportions of monolignols in labile β-aryl ether structures. The methods revealed consistently major differences between the bark and wood lignins (Tables 1−3). The predominant unit in bark lignin was G over S and H; the S/G ratio was found to vary from 0.2 to 0.9, depending upon the method used. On the contrary, S was significantly more abundant in wood lignin, with the S/G ratio ranging from 1.4 to 2.1, similar to data reported earlier.9,10 The monomer yields and S/G ratio increased progressively from bark to inner bark to wood. This might explain why the bark lignin is more condensed than the lignin in other tissues of willow.23 Generally, DFRC gave lower monomer yields than thioacidolysis and NBO, but the trend in the S/G ratio, when moving from bark to wood, was the same, independent of the analytical method used. Table 1 summarizes the yields of the thioacidolysis products from WCW and EL of bark, inner bark, and wood. The total monomer yield from WB-WCW was roughly 50% smaller than that from inner bark and around 90% smaller than that from wood. The lignin monomer yield that released from WWWCW (2082 μmol/g of lignin) was comparable to the monomer yield from poplar (2064 μmol/g of lignin) and twice that from loblolly pine (1055 μmol/g of lignin). Obviously, the D
DOI: 10.1021/acs.jafc.8b02014 J. Agric. Food Chem. XXXX, XXX, XXX−XXX
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Journal of Agricultural and Food Chemistry
Table 4. Structural Characteristics of WCW Lignin and EL from Bark, Inner Bark, and Wood Obtained by Integration of 1 H−13C Correlation Contours in the Corresponding HSQC Spectra WB WCW lignin interunit linkages (%)a β−O−4′ aryl ethers (A′/A) phenylcoumaran (B) resinols (C) lignin aromatic unitsb G (%) S (%) S/G ratio
WIB EL
WCW
86 2 12 67 33 0.5
WW EL
WCW
80 3 17
61 39 0.6
43 57 1.3
51 49 0.9
EL 89 3 7
33 67 2.0
34 66 1.9
a Percentage on total volume of A, B, and C signals (calculated from the α-C/H correlations). bPercentage of total volume of G2, G′2, S2/6, and S′2/6 signals.
Figure 2. Anomeric region of glycosides (δC/δH of 90−110/3.5−6.0) in HSQC spectra of WCW and EL samples of bark, inner bark, and wood in DMSO-d6/pyridine-d5 (4:1), with color codes of (blue) cellulose, (green) xylan, and (red) pectin. Abbreviations: β-D-Glcp, β-D-glucopyranoside; α-D-Xylp, α-D-xylopyranoside; β-D-Xylp, β-D-xylopyranoside; 2-O-Ac-β-D-Xylp, acetylated β-D-xylopyranoside; 4-O-MeGlcA, 4-O-methyl-α-Dglucuronic acid; α-L-Rhap, α-L-rhamnopyranose; α-D-GalpA, α-D-galactopyranuronic acid; β-D-Galp, β-D-galactopyranoside; α-L-Araf, α-Larabinofuranoside; and β-L-Araf, β-L-arabinofuranoside.
samples (Figure 1). The C α −H α , C β−H β , and C γ −H γ correlations of A were observed at δC/δH of 71.9/4.97, 83.6/ 4.38, and 60.0−60.8/3.59−3.69 ppm, respectively. Additionally, Cα−Hα, Cβ−Hβ, and Cγ−Hγ correlations in the phenylcoumaran substructures B were seen at δC/δH of 86.9/5.55, 53.2/3.57, and 62.8/3.74 ppm, respectively, while the resinol (β−β) substructures C showed the corresponding correlations at δC/δH of 84.9/4.66, 53.4/3.03, and 70.9/4.15 ppm. WW-EL showed the unsaturated Cγ−Hγ correlation of cinnamyl alcohol end group X1 at 61.5/4.17 ppm and Cα−Hα and Cβ−Hβ correlations of spirodienone SD at 5.18/81.7 and 2.86/60.9 ppm, respectively. The X1 and SD signals were not seen in HSQC spectra of WB-EL and WIB-EL. Lignin Monomers. The main signals in the aromatic region of the 2D HSQC spectra of all EL samples originated from G and S units of lignin (Figure 1). The S units showed prominent resonances for C2,6−H2,6 correlations at δC/δH of 103.9/6.76 and 106.3/7.10 ppm. The G units showed C2−H2 correlation at δC/δH of 110.8/7.02 ppm and C5−H5 and C6−
EL of bark, inner bark, and wood were investigated by solution-state 2D NMR spectroscopy. 2D HSQC of the EL preparations provided information on typical structural characteristics of lignin, such as the S/G ratio, linkage types between the monomers, and purity of the samples. The impurities included some carbohydrates and proteins, which were also comparatively explored in the corresponding WCW samples. All of the spectral data were assigned by comparison to the published literature.11,25,28−32 The aromatic (δC, 95−152 ppm; δH, 5.5−8.0 ppm) and sidechain (δC, 48−91 ppm; δH, 2.0−6.0 ppm) regions of HSQC NMR spectra of EL from bark, inner bark, and wood are depicted in Figure 1. The majority of lignin signals were also observed in HSQC spectra of the WCW preparations although several polysaccharide signals dominated these spectra and partially overlapped with some lignin signals. Interunit Linkage Characterization. The correlation peaks from methoxyl groups and side chains in the β-aryl ether substructures A dominated the HSQC spectra of the EL E
DOI: 10.1021/acs.jafc.8b02014 J. Agric. Food Chem. XXXX, XXX, XXX−XXX
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Journal of Agricultural and Food Chemistry H6 correlations at δC/δH of 6.5−7.1/115−120 ppm. A small fraction of G′ signals were from α-carbonyl structures, which seemed to be somewhat more abundant in bark and inner bark in comparison to wood, which again was richer in α-carbonylcontaining S′ units. Cinnamyl alcohol end groups (X1) with Cα−Hα and Cβ−Hβ correlations at δC/δH of 128.4/6.49 and 128.4/6.29 ppm, respectively, were present in wood but not in bark and inner bark. Aromatic Amino Acid Moieties. Three strong and wellresolved HSQC signals observed at δC/δH of 126.01/7.13, 127.81/7.20, and 129.01/7.22 ppm revealed the presence of phenylalanine, especially in bark and inner bark but also in wood in a smaller amount.25 Another peak at δC/δH of 129.90/ 7.04 ppm showed the occurrence of tyrosine (C2,6−H2,6) likewise in bark and inner bark and in a lower content in wood. A large number of aliphatic C−H signals of amino acids were present in the 50−85/3−5 ppm region of the HSQC spectra (Figure 1). These protein signals were also present in the NMR spectra of the corresponding WCW preparations, confirming that phenylalanine and tyrosine were naturally present in the original biomass, especially in the bark, as reported earlier for not only willow33 but also corn cob and kenaf samples.25 However, a smaller fraction of the amino acids could possibly arise from the cellulolytic proteins used in EL separation. The parenchyma tissue of inner bark may act as a temporary storage of reduced nitrogen as a protein reserve in overwintering.34 Quantitation. Table 4 summarizes the relative abundances of the lignin monomers and the main linkage types between the monomers. The quantitation of these structures was based on the volume integration of the corresponding contours in the HSQC spectra.12,28 The S/G ratios of EL from bark, inner bark, and wood, determined by HSQC, were 0.6, 0.9, and 1.9, respectively. These values were similar to those obtained by the three wet chemical methods (thioacidolysis, NBO, and DFRC) on average (0.6, 0.9, and 1.7, respectively). As characteristic of most G/S lignins, WB-EL and WIB-EL were rich in β-aryl ether structures (86 and 80%, respectively), followed by resinols (12 and 17%) and phenylcoumarans (2 and 3%). In comparison to WIB-EL, both WB-EL and WW-EL contained more β-aryl ether structures (89%) and less resinols (7%). Polysaccharide Anomeric Correlations. The presence of carbohydrates in the WCW and EL samples was studied through the correlations in the δC/δH of 90−110/3.5−6.0 ppm region, which is characteristic for anomeric centers of glycosides (Figure 2). Full assignment of HSQC signals of this region is presented in Table S2 of the Supporting Information. Strong signals of cellulose (β-D-Glcp) and acetylated 4-O-methylglucuronoxylan35 (β-D-Xylp, 2-O-Ac-βD-Xylp, 2,3-di-O-Ac-β-D-Xylp, and 4-O-MeGlcA) dominated the HSQC spectrum of WW-WCW (Figure 2). Although the same signals were also present in the HSQC spectra of WCW preparations of bark and inner bark, the strong signals of pectic polysaccharides36 (α-D-GalpA, α-L-Rhap, α-L-Araf, β-L-Araf, and β-D-Galp) are characteristic of inner bark.5 Traces of the pectic polysaccharides were also present in EL preparations of bark and inner bark, while xylan-derived signals were more dominant in the HSQC spectrum of WW-EL. This may indicate that the bark lignin is covalently bonded to pectins, whereas wood lignin forms complexes with xylan. On the basis of the 2D NMR data, the carbohydrate content of the EL samples was estimated to be ca. 10%.
In conclusion, willow bark lignin has a significantly lower syringyl unit content in comparison to wood lignin. Although β-aryl ether interunit linkages are dominant in both bark and wood lignin, resinol structures are more frequent in the bark lignin. Willow bark is rich in proteins and pectins, and the latter seem to connect with the bark lignin. In contrast, wood lignin is rather linked with xylans. These differences in the chemical composition and structure may partly reflect the fact that the share of thin-walled parenchymous tissue is much larger in bark than in wood. However, further studies would be needed to experimentally differentiate between the structural polymers present in the parenchymous and sclerenchymous tissues of the bark. This information would be valuable in designing processes for the separation of willow bark sclerenchyma bundles and fibers as part of the proposed willow biorefinery scheme.5
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ASSOCIATED CONTENT
S Supporting Information *
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jafc.8b02014.
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Section of the assignments for the interunit linkage (Table S1), polysaccharide lignin 2D 1H−13C HSQC correlation peaks (Table S2), and WCW NMR spectra from the 2D HSQC spectra (Figure S1) (PDF)
AUTHOR INFORMATION
Corresponding Author
*Telephone: 00358-505160048. E-mail: tapani.vuorinen@ aalto.fi. ORCID
Jinze Dou: 0000-0001-8782-3381 Hoon Kim: 0000-0001-7425-7464 John Ralph: 0000-0002-6093-4521 Tapani Vuorinen: 0000-0002-5865-1776 Funding
This work was funded by the Foundation of Walter Ahlström and the Finnish Cultural Foundation. Hoon Kim, Yanding Li, Dharshana Padmakshan, Fengxia Yue, and John Ralph were funded by the DOE Great Lakes Bioenergy Research Center (DOE BER Office of Science DE-FC02-07ER64494 and DESC0018409). Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS The authors thank Sarah Liu for the skillful technical assistance in the isolated enzyme lignin (whole cell wall) preparation and the synthesis of the authentic DFRC compound by Matthew Robert Regner from the Department of Biochemistry and the DOE Great Lakes Bioenergy Research Center, University of WisconsinMadison. Appreciation is extended to Yibo Ma from Aalto University for the preparation of the willow wood.
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REFERENCES
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DOI: 10.1021/acs.jafc.8b02014 J. Agric. Food Chem. XXXX, XXX, XXX−XXX
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