Title
Author(s)
Citation
Issue Date
An
ideal lignin
facilitates full biomass utilization
Li, Yanding; Shuai, Li; Kim, Hoon; Motagamwala, Ali
Hussain; Mobley, Justin K.; Yue, Fengxia; Tobimatsu, Yuki;
Havkin-Frenkel, Daphna; Chen, Fang; Dixon, Richard A.;
Luterbacher, Jeremy S.; Dumesic, James A.; Ralph, John
Science advances (2018), 4(9)
2018-9-28
URL
http://hdl.handle.net/2433/234963
Right
© 2018 The Authors, some rights reserved; exclusive licensee
American Association for the Advancement of Science. No
claim to original U.S. Government Works. Distributed under a
Creative Commons Attribution NonCommercial License 4.0
(CC BY-NC).
Type
Journal Article
Textversion
publisher
Kyoto University
SCIENCE ADVANCES | RESEARCH ARTICLE
ORGANIC CHEMISTRY
An “ideal lignin” facilitates full biomass utilization
Yanding Li1,2, Li Shuai1,3, Hoon Kim1,4, Ali Hussain Motagamwala1,5, Justin K. Mobley1,
Fengxia Yue1,4, Yuki Tobimatsu1,4,6, Daphna Havkin-Frenkel7,8, Fang Chen9,10,
Richard A. Dixon9,10, Jeremy S. Luterbacher11, James A. Dumesic1,5, John Ralph1,2,4*
Lignin, a major component of lignocellulosic biomass, is crucial to plant growth and development but is a major impediment to efficient biomass utilization in various processes. Valorizing lignin is increasingly realized as being essential. However, rapid condensation of lignin during acidic extraction leads to the formation of recalcitrant condensed
units that, along with similar units and structural heterogeneity in native lignin, drastically limits product yield and
selectivity. Catechyl lignin (C-lignin), which is essentially a benzodioxane homopolymer without condensed units,
might represent an ideal lignin for valorization, as it circumvents these issues. We discovered that C-lignin is highly
acid-resistant. Hydrogenolysis of C-lignin resulted in the cleavage of all benzodioxane structures to produce catechyltype monomers in near-quantitative yield with a selectivity of 90% to a single monomer.
Lignin is a polymeric material composed of phenylpropanoid subunits
and is one of the largest sources of naturally produced aromatics on the
planet. Because of its aromatic nature, lignin has a higher energy density
than polysaccharide polymers, as well as a higher potential commercial
value (1). However, because of lignin’s complexity, its efficient utilization, either as a polymer or from its derivable small-molecule products,
is currently problematic (1–3).
Although mild depolymerization methods, such as oxidative (4, 5)
and hydrogenolytic (6–8) procedures, have produced encouraging
results in laboratory-scale experiments, their applicability in industrial
processes has been limited. Direct hydrogenolysis, that is, the hydrogenation of unprocessed solid biomass by a heterogeneous metal catalyst,
remains one of the most promising methods for cleaving lignin’s ether
bonds and producing aromatic monomers in high yields (8–10). However, hydrogenolysis still suffers from product complexity issues. In
most wild-type biomass, the lignin polymer is composed of three phenylpropanoid subunits—p-hydroxyphenyl (H), guaiacyl (G), and syringyl
(S)—derived by combinatorial radical coupling from the three main
monolignols (p-coumaryl, coniferyl, and sinapyl alcohols). Although
H units are typically at low-levels, this results in at least three different
types of monomers (H, G, and S), each with a selection of side chains, as
the primary hydrogenolysis products, which makes monomer separation and utilization difficult. Lignin’s principal alkyl-aryl-ether units
with their b–O–4 interunit bonds (45 to 85%) can be selectively cleaved,
but other linkages including b–5 (1 to 12%), b–b (5 to 12%), 5–5 (1 to
9%), 4–O–5 (~2%), and b–1 (1 to 2%), which are also present in lignins,
1
U.S. Department of Energy Great Lakes Bioenergy Research Center, and Wisconsin
Energy Institute, University of Wisconsin–Madison, Madison, WI 53726, USA. 2Department of Biological Systems Engineering, University of Wisconsin–Madison, Madison, WI
53706, USA. 3Department of Sustainable Biomaterials, Virginia Tech, Blacksburg, VA
24061, USA. 4Department of Biochemistry, University of Wisconsin–Madison, Madison,
WI 53706, USA. 5Department of Chemical and Biological Engineering, University of
Wisconsin–Madison, Madison, WI 53706, USA. 6Research Institute for Sustainable
Humanosphere, Kyoto University, Gokasho, Uji, Kyoto 611-0011, Japan. 7Department of Plant Biology and Pathology, Rutgers, State University of New Jersey,
New Brunswick, NJ 08901, USA. 8Bakto Flavors LLC, 772 Cranbury Crossroad, North
Brunswick, NJ 08092, USA. 9BioDiscovery Institute and Department of Biological
Sciences, University of North Texas, Denton, TX 76203, USA. 10Center of Bioenergy
Innovation, Oak Ridge National Laboratory, Oak Ridge, TN 37831, USA. 11Laboratory
of Sustainable and Catalytic Processing, Institute of Chemical Sciences and Engineering, École Polytechnique Fédérale de Lausanne, CH-1015 Lausanne, Switzerland.
*Corresponding author. Email: jralph@wisc.edu
Li et al., Sci. Adv. 2018; 4 : eaau2968
28 September 2018
remain largely uncleaved (8); carbon-carbon (C–C) and diaryl ether
(4–O–5) units typically result from dimeric or higher oligomeric products.
The use of extracted lignins rather than whole biomass has the
advantage that the material can be fully dissolved in organic solvents,
facilitating catalyst recovery and continuous processing. However,
acidic industrial lignin fractionation is known to cause some b-ether
cleavage and condensation between units via the electrophilic substitution of acid-generated benzylic carbocation intermediates on the electronrich aromatic rings (7, 11), limiting depolymerization yields (Scheme 1A)
(12–14). There are some elegant solutions focusing on suppressing the
condensation reaction, either using a capping agent (7, 15) or using twostep strategies (4, 5, 11). However, extra chemicals or catalysts are needed
to achieve this goal.
Bioengineered biomass could be used to achieve higher hydrogenolysis yields and simpler product mixtures. For example, the recent use of
formaldehyde protection during lignin extraction from a high-S poplar
lignin (7, 16) that has up to 98% syringyl S units and ~90% b–O–4 linkages [from nuclear magnetic resonance (NMR) estimates] prevented
condensation reactions and allowed an unprecedentedly high monomer
yield (78%) under hydrogenolytic conditions (7). However, even in this
high-S lignin, some 10% of the linkages are C–C bonds that do not
cleave. The use of formaldehyde to protect the lignin from condensation
reactions also resulted in some formaldehyde addition to the ring, complicating the hydrogenolysis products with methyl-substituted aromatics. Without formaldehyde, the lignin extracted under acidic
conditions had significant condensation, thwarting the production of
monomers and resulting in a hydrogenolysis monomer yield of only
26% (7). Although new methods for displacing formaldehyde for the
protection from acid-catalyzed condensation reactions, retaining much
of the yield (70%) and producing a simpler monomer mix, have recently
been revealed (17), extra protection chemicals remain necessary during
the lignin extraction.
RESULTS
An “ideal lignin” archetype
On the basis of the plethora of information stemming from the lignin
biosynthetic research community over the last decade, and with the revelations regarding lignins’ structural malleability from studies on lignin
pathway mutants and transgenics as well as on various “natural” plants
discovered to have unusual lignins, researchers have been able to
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INTRODUCTION
Copyright © 2018
The Authors, some
rights reserved;
exclusive licensee
American Association
for the Advancement
of Science. No claim to
original U.S. Government
Works. Distributed
under a Creative
Commons Attribution
NonCommercial
License 4.0 (CC BY-NC).
SCIENCE ADVANCES | RESEARCH ARTICLE
contemplate designing lignins for improved utilization (18). It is now
a realistic juncture to posit the characteristics for an ideal lignin archetype for biomass processing. For the depolymerization of the
polymer to monomers, lignin should have at least the following three
characteristics. First, if acidic pretreatment is used, then it should be
stable under acidic conditions to prevent condensation and the generation of undesired new C–C bonds. Second, it should contain only
ether (C–O) interunit linkages in its backbone so that it can be fully
depolymerized. Finally, it should be generated in planta from a single
phenylpropanoid monomer to allow the production of the simplest
array of compounds.
We have reported the discovery of an unusual catechyl lignin
(C-lignin) present in the seed coats of vanilla (Vanilla planifolia)
(19) and various members of the Cactaceae of the genus Melocactus
(20). In this special case, the lack of O-methyltransferase (OMT) activity
for conversion from catechyl C to guaiacyl G and, subsequently, on to
syringyl S, aromatic-level precursors, results in 100% C units in the cell
wall (CW). This C-lignin was, somewhat surprisingly, found to be
essentially a homopolymer synthesized almost purely by b–O–4 coupling of caffeyl alcohol with the growing polymer chain, producing
benzodioxanes as the dominant unit in the polymer (Fig. 1A). If it has
particular stability toward biomass pretreatment conditions, then this
A
B
S
C
M3
M1
Scheme 1. Mechanisms for lignin condensation, C-lignin structure, and monomer M3 formation. (A) Mechanism of lignin acidolysis and condensation routes. (B) The
benzodioxane structure acts as a “shield” that can protect C-lignin from unwanted acidolysis and condensation reactions. (C) Proposed mechanism for the cyclization
reaction of M1 to M3.
C6
A EL
60
4
O
O
C3
OH
Benzodioxane
internal units
4
O
O
C LBL
B KL
C-NR6
C2
C5
C4
80
n
1
HO
6
5
2
3
HO
4
Benzodioxane
end-units
100
OH
C1
Tyr
120
Phe
8
7
Pyridine
Protein
Cellulose
Unresolved
6
5
4
3
140
13
8
7
6
5
4
3
8
7
6
5
4
1
H
C
ppm
Fig. 1. NMR spectra. Partial 2D HSQC NMR spectra of (A) EL, (B) KL, and (C) LBL from vanilla (V. planifolia) seed coat. There are no obvious lignin structural changes after the acidic
lignin extraction processes. Cellulose was labeled following the conventional monosaccharide nomenclature; NR is the nonreducing end of the cellulose. Protein residuals were
labeled by the aromatic amino acid. Tyr, L-tyrosine; Phe, L-phenylalanine; ppm, parts per million.
Li et al., Sci. Adv. 2018; 4 : eaau2968
28 September 2018
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G
SCIENCE ADVANCES | RESEARCH ARTICLE
C-lignin might therefore represent an example of such an ideal lignin
that can, in principle, be depolymerized to a single product by hydrogenolysis. Furthermore, this substrate has the potential to produce valuable catechol monomers, whereas the large majority of monomers
produced from lignin have been S or G derivatives (1, 2). Expanding
the arsenal of lignin-derived platform molecules could play an important role in the successful use of this fraction within future biorefineries.
Here, we describe the ideal nature of this lignin via a revised compositional characterization of the vanilla seed coat fiber, new features of the
C-lignin’s reactivity and stability, and our successful attempts at
converting it to monomers in near-quantitative yields.
Normalized intensity (a.u.)
Sample Mn (kDa) MW (kDa) PDI
106
Mp (kDa)
LBL
2.0
13.5
6.7
15.8
EL
0.8
13.2
16.3
12.8
LBL
EL
105
10000
1000
100
Relative molecular weight (Da)
10
1
Fig. 2. Molecular weight profiles. Molecular weight profiles of EL (cyan) and LBL
(magenta) from V. planifolia seed coat measured by gel-permeation chromatography
(GPC). The x axis indicates the apparent molecular weight of individual lignin polymers
and is shown as a log scale. The y axis shows the response of a UV-light detector (at
280 nm) normalized to the most abundant signal in each chromatogram. The most
abundant signal in the each of the two samples corresponds to a molecular weight
of ~13,000 Da (determined via polystyrene standards); comparison shows that there
was no obvious lignin polymer degradation during the acid pretreatment. PDI is the
polydispersity index. a.u., arbitrary units. MW, molecular weight.
Li et al., Sci. Adv. 2018; 4 : eaau2968
28 September 2018
Response of C-lignin to traditional degradative methods
To investigate the potential for C-lignin depolymerization, we applied
two traditional lignin degradative analytical methods, alkaline nitrobenzene oxidation (NBO) and thioacidolysis, to a C-lignin model compound, the caffeyl alcohol dimer D1 (C-dimer), and to the vanilla bean
seed coat CW (Fig. 3 and fig. S3). Although relatively low yields of the
corresponding monomeric products (30 to 60%) were obtained from
the dimeric compound, the use of the CW gave monomeric products
in extremely low yields (<1%). As discussed widely in the past, both
thioacidolysis and alkaline oxidation need the involvement of a free benzylic hydroxyl group on the lignin side chain (25, 26). It was therefore
concluded that, because of the stability of the 1,4-benzodioxane structure, especially under the tested acidic and alkaline oxidative conditions,
traditional lignin chemical degradation methods are ineffective for the
depolymerization of C-lignin. A computational approach to evaluate
the bond dissociation energy (BDE) of C-lignin using density functional
theory models suggested that depolymerization of C-lignin is theoretically possible (27). Although the benzodioxane b–O–4 bond calculates
to have a slightly higher BDE value than a conventional b–O–4 bond, it
is still much lower than the BDEs of lignin’s C–C bonds (28).
Catalytic hydrogenolysis of C-lignin
We reasoned that hydrogenolysis had the potential to more efficiently
depolymerize C-lignin. We first sought efficient methods for cleaving
dimeric model D1, rationalizing that, although the corollary is not necessarily true, any reaction conditions that did not produce high yields
from D1 would have little chance of being effective on the polymer.
When hydrogenolysis was applied to the C-dimer D1 and vanilla seed
coat CW, analysis by gas chromatography with flame-ionization detection (GC-FID) showed that the products were rather simple with dominant products M1 (catechylpropanol), M2 (catechylpropane), and M3
(chroman-6,7-diol), together with some minor products (Fig. 3). The
major products, M1 and M2, were identified by comparison with
authentic synthetic standards. The initially puzzling minor product
M3, which is a cyclization product from M1, was separated from the
product mixture by silica-gel chromatography, characterized, and structurally identified by NMR and high-resolution mass spectrometry
(MS). Because it was not obvious whether the chromane ring oxygen
originated from the lignin g-OH or from water, the hydrogenolysis
reaction was run in 18O-labeled H218O. No 18O was detected in the
product M3, so the cyclization mechanism was concluded to involve
the g-OH via a radical disproportionation reaction (Scheme 1C) (29).
This is the first report of this lignin hydrogenolysis product. The minor
impurity peaks displayed in the chromatograms from the CW materials
(fig. S3) were derived from the solvent, polysaccharide, and fatty acid
products, which were identified via GC-MS.
Monomer production data under different conditions are shown in
Fig. 4. Yields are normalized to the total molar concentration of caffeyl
alcohol in C-lignin determined from quantitative 13C NMR (table S2).
Not surprisingly, the monomer distributions were heavily affected by
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Acid stability of C-lignin
Because of the lack of an accessible and eliminable benzylic hydroxyl
group in C-lignin units (Scheme 1B), condensation reactions due
to the formation of benzyl cations might be mitigated under acidic
conditions. We therefore examined the acid stability of the polymer
to determine whether acidolytic methods could be used to purify the
lignin. Comparison of the two-dimensional (2D) heteronuclear
single-quantum coherence (HSQC) NMR spectra from the enzyme
lignin (EL) (derived by removing polysaccharides via crude cellulases
treatment) (21) and Klason lignin (KL) showed no significant differences in the lignin structure (Fig. 1, A and B) (22). The C-lignin survives
even the harshest of acidic pretreatment methods—the KL isolation
procedure includes a 1-hour treatment in 72% (w/w) sulfuric acid,
followed by dilution to 4% (w/w) sulfuric acid and autoclaving at
121°C for 1 hour—while retaining its original lignin structure. An
efficient acidic lithium bromide (LiBr) pretreatment method was also
used to purify the lignin. This treatment method is known for its quick
and near-quantitative removal of the polysaccharides to give an LiBr
lignin (LBL) (Fig. 1C) (23). The molecular weight of the LBL was shown
to be similar to that of the EL (Fig. 2). The C-lignin polymer appeared to
survive this pretreatment based on the retention of its key lignin structural features in its NMR spectra and little change in its molecular
weight distribution. On the basis of these results, we can conclude that,
unlike normal S-G lignins, polysaccharides can be removed via acid
pretreatment from C-lignin without its suffering from unwanted condensation reactions. After removing the polysaccharides, the resulting
lignins (EL, KL, and LBL) were completely soluble in various organic
solvents [for example, acetone, dioxane, or tetrahydrofuran (THF)]
mixed with water to match lignin solubility parameters (24). Efficient
lignin solubilization should greatly facilitate continuous processing in
an industrial setting.
SCIENCE ADVANCES | RESEARCH ARTICLE
HO
Products (TMS derivatives):
HO
Products from polysaccharides
HO
OH
M1
M2
HO
OH
Unidentified products and impurities
M4
M5
M6
M7
M3
HO
HO
HO
HO
HO
HO
HO
HO
HO
HO
M8
O
HO
HO
O
OH
HO
O
D1
OH
OH
CW
12
13
14
15
16
17
18
19
20
21
D1
min
Fig. 3. GC-FID spectra of hydrogenolysis products from dimeric compound D1 and from CW. Hydrogenolysis condition: Pt/C, 200°C, 40-bar H2, 15 hours. Coloring
of peaks matches that of the structures for monomers M1 to M8. Products from polysaccharide in the CW are colored light green, and unidentified products from other
non-lignin compounds are left in black. TMS, trimethylsilyl. Note that the upper D1 product chromatogram is offset by ~0.3 min.
C-dimer
100
CW
LBL
CW
CW
Pd/C
LBL
Ru/C
Monomers (mol%)
80
60
40
20
0
H
H
eO
M
eO
ne
H
xa
io
M
D
ne
H
eO
M
eO
M
F
TH
xa
H
eO
io
D
M
ne
H
eO
M
H
xa
io
D
eO
M
473 K, 15 hours, 4-MPa H2
HO
M4
HO
HO
M5
HO
HO
M1
M7
OH
HO
M8
HO
HO
HO
HO
HO
M6
HO
HO
M2
OH
HO
O
M3
HO
HO
Fig. 4. Hydrogenolysis monomer yields from different catalyst and solvent
combinations. Yields are on a C-lignin molar basis (see also table S3, from left to
right: entries 1, 2, 3, 5, 7, 9, 10, 12, 14, 19, and 21).
the choice of catalyst and solvent (17, 30, 31). Here, we illustrate that
Pt/C showed a slightly higher reactivity, whereas Pd/C and Ru/C
showed a much better product selectivity. More side chain truncation
products were obtained from C-LBL compared to that from vanilla seed
coat CW, suggesting that a significant degree of side chain truncation
occurred during the acid pretreatment stage or that the isolated lignin
was more accessible to the catalyst. In terms of solvent effects, methanol
produced a slightly higher monomer yield compared to dioxane,
whereas THF gave a substantially lower yield. Both monomer yield
and reaction selectivity were maximized using Pd/C or Ru/C as catalyst
and methanol as the solvent. Retaining or losing the hydroxy group on
the side chain can be controlled by simply changing the catalyst to satisfy the different intended purposes for using the catechyl monomers.
Li et al., Sci. Adv. 2018; 4 : eaau2968
28 September 2018
Thus, Pd/C produced the catechylpropanol monomer M1 with 89%
selectivity, whereas Ru/C produced the catechylpropane monomer
M2 with 74% selectivity. Increasing the hydrogenolysis reaction time
from 3 to 15 hours (table S3, entries 16 to 19) led to an ~10% increase
in lignin conversion and monomer yield. The resulting product oil
mixture after vacuum drying was completely soluble in methanol,
ethanol (EtOH), dioxane, pyridine, and other solvents but only partially soluble in acetone, ethyl acetate, and dichloromethane (DCM)
due to the presence of products from degraded polysaccharides and
other non-lignin components. The mass balance and total organic
carbon (TOC) analyses (Table 1) indicated that volatile products
were minimal or insignificant.
A 2D HSQC NMR spectrum of the total hydrogenolysis product,
which was completely soluble in dimethyl sulfoxide (DMSO)/pyridine
(4:1, v/v), demonstrated that the C-lignin had been completely depolymerized, that is, no detectable residual benzodioxane structures remained (fig. S4A). The major products were fully authenticated by
comparison with synthetic compounds M1 and M2 and with authenticated isolated M3. No detectable products from side reactions or recondensation were detectable. The GPC molecular weight profile of the
hydrogenolysis products mixture from C-LBL before and after the hydrogenolysis reactions showed a dominant monomer peak (fig. S4B).
The high–molecular weight fractions were separated from monomer
fractions, and the fractions were characterized by HSQC NMR (fig. S7).
The data revealed that only traces of the original benzodioxane structures
from the C-lignin remained in the product and that the high–molecular
weight fractions contained only nonaromatic components present in the
original sample and were therefore not from the lignin proper. It can
therefore be safely concluded that essentially all of the C-lignin in the
samples was depolymerized to monomeric compounds during hydrogenolysis. The non-lignin components in the lignin stream were nonextractable oils, waxes, or the other (difficult to remove) components in the
sample that are not necessarily associated directly with the phenylpropanoid polymer.
DISCUSSION
Prospects for C-lignin and its derived
catechylpropanoid monomers
Catechols in nature are remarkably biochemically active; because of the
interaction of the vicinal phenolic hydroxyl groups, catechols play a vital
role in both biomedical and biomimetic functional materials (32). Their
synthesis is challenging because of the difficulty of transforming phenols
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Pt/C
LBL
SCIENCE ADVANCES | RESEARCH ARTICLE
Table 1. Mass balance and TOC on hydrogenolysis of C-LBL and its
resulting product oil.
Feed
CW
Dissolved C-LBL
55–74%
~100%
23–35%
50–60%
TOC of C-LBL
—
62.66 ± 0.23%
TOC of product oil
—
61.44 ± 0.34%
Solid recovery*
Oil recovery
†
*Solid includes recovered CW material and catalyst.
CW and C-LBL mass basis.
†Oil yield on a
Li et al., Sci. Adv. 2018; 4 : eaau2968
28 September 2018
MATERIALS AND METHODS
C-lignin sample pretreatment
Processing of seed coat material
Vanilla seed and pod were received as a mixture from a natural vanilla
processing plant (Bakto Flavors LLC). The mixture was sifted, and the
lower-density remaining pod powder was blown away using a heat gun
(set on cold). Preparation of vanilla seed coat NMR samples was via
methods described previously (22). Briefly, isolated vanilla seed coats
(4 × 300 mg) were ball-milled (30 × 10 min, 5-min cooling cycle) using
a Retsch PM100 ball mill vibrating at 600 rpm with ZrO2 vessels
containing ZrO2 ball bearings. Preground seed coat was extracted using
a modified Bligh and Dyer extraction (41) to remove oils and extractives.
Modified Bligh and Dyer extraction
Vanilla seed material (100 g in total) was shaker-milled (MM400,
Retsch) at 3600 rpm for 5 min using a 50-ml stainless steel jar and
a single 20-mm ball bearing. The milled sample was transferred to a
1-liter volumetric flask, and a magnetic stir bar was added. Deionized
(DI) water (80 ml), chloroform (100 ml), and methanol (200 ml) were
added, and the mixture was stirred at 50°C for 30 min. To the mixture
was then added 100 ml more of chloroform, and then, after another
30 min, 100 ml of DI water was added. The stirring was continued at
50°C for 24 hours, and the insoluble material was removed by centrifugation (3800 rpm for 15 min), retaining the solids by decanting
off the solvent and keeping the filtrate as well. The residue was extracted
again by the same method. The filtrates were combined, and the solvents were removed by rotary evaporation to produce the extractives
fraction for analysis.
EL from vanilla seed coat
The ball-milled extract-free vanilla seed coat material (1 g) was
placed in centrifuge tubes and digested at 35°C with crude cellulases
[CELLULYSIN cellulases, Trichoderma viride; sample (50 mg/g) in
acetate buffer (pH 5.0); two times over 3 days; fresh buffer and enzyme were added each time; catalog no. D00074989, Calbiochem],
leaving all of the phenolic polymers and residual polysaccharides totaling 859 mg (85.9%) (table S1).
Acidic LiBr pretreatment of C-lignin from vanilla seed coat
C-LBL was prepared using the acidic LiBr trihydrate method described
previously (23). Briefly, ball-milled extract-free vanilla seed coat material
(1 g) was added into a 40-ml glass vial with a polytetrafluoroethylene
(PTFE) lined cap, together with 4.50 ml of acidic 60 weight % (wt %) LiBr
solution containing 0.04 M HCl. The vial was immersed into an oil bath
preheated at 110°C under magnetic stirring. The mixture was filtered
under vacuum and washed with water. The residues were dried at 40°C
under reduced pressure (yield, 72.4%; table S1).
Compositional analysis
KL analysis was performed by the two-stage sulfuric acid hydrolysis
following the National Renewable Energy Laboratory’s standard protocol
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to catechols; although researchers have recently developed several
catechol synthetic methods (33), applying those methods at scale remains complicated. There is little reference to high-yielding biomass
conversion to catechols, although catechols were reported as hydrogenolysis products from organosolv lignin of candlenut shells (using
Cu-doped porous metal oxides) in which the cleaving of the aromatic
methoxyl groups during the reaction was claimed (34). Catechols act
as important intermediates for the conversion of lignin-derived monomers to value-added platform chemicals via the bacterial b-ketoadipate
pathway (35). We therefore contend that it would be beneficial, and
more energy-efficient for aromatic metabolism/catabolism, if high
yields of catechols could be obtained directly from lignin.
Our study provides a new perspective for the production of catechols
from a renewable biomass source rather than petroleum. Compounds
M1 and M2 are currently not available in bulk, so their commercial
value is not obvious. However, an enriched diversity of the raw materials
from the catechol family would likely provide significant value. A LiBr
pretreatment method is able to convert a fraction of b–O–4 units in
S-G–type lignins into benzodioxanes (36). Large amounts of catechols
are potentially producible if we could produce benzodioxane-type lignins
in energy crops. We do not yet know if genetically engineered plants,
including plantation trees such as pines and poplars, in which the production of lignin is on a large scale, will tolerate C-lignins in stem tissues;
C-polymers have been evidenced in a gymnosperm tracheary element
system, in which OMT activity was down-regulated (37), but have not
yet been found in OMT–down-regulated dicots, and we suspect that
additional activities will need to be suppressed for the synthesis and
deposition of the C-lignin polymer. Given the unique acid-resistant
property of C-lignin, the potential value of the monomeric products,
the homogeneous nature of C-lignins that is already known to aid lignin
fiber production (38), and the high conversion to catechol monomers
by hydrogenolysis reported here, we suggest that continuing to pursue
the means to produce C-lignins in planta is decidedly worthwhile.
C-lignin therefore has numerous compelling features for a biorefinery
operation aimed at delivering value from its lignin component. It maintains its native structure after treatment under even strongly acidic
conditions; acid pretreatment can therefore be applied to vanilla seed
coats to recover the polysaccharide while retaining the native C-lignin
structure. After sufficient pulverization followed by acid pretreatment,
C-lignin could be dissolved in organic solvents, enabling both detailed
NMR analysis and continuous processing schemes. C-lignin can be completely depolymerized by a hydrogenolytic method to produce simple
monomeric catechols near-quantitatively and, by selecting the catalyst,
with a single monomer accounting for 90% of the monomer product.
The yield and selectivity for a single monomer are higher than for any
other lignin or biomass to date (fig. S6). There is therefore considerable
potential for economic hydrogenolysis of C-lignin–rich waste biomass
resources only now being structurally characterized, such as Jatropha
(Jatropha curcas) seed coats (39) and candlenut (Aleurites moluccanus)
shells (40), and via genetic engineering if high levels of C-lignin could
be expressed in traditional biomass sources. Such an approach toward
significantly valorizing lignins and biomass in biorefining processes
would aid process economics.
SCIENCE ADVANCES | RESEARCH ARTICLE
C-lignin characterization and quantification
Lignin characterization by 2D NMR spectroscopy
NMR spectra were acquired on a Bruker Biospin AVANCE III
700 MHz spectrometer fitted with a cryogenically cooled 5-mm QCI
1
H/31P/13C/15N gradient probe with inverse geometry (proton coils
closest to the sample), and spectral processing used Bruker’s TopSpin
3.5pl6 (Mac) software. For NMR experiments, ball-milled whole vanilla
seed coat material was swelled in DMSO-d6/pyridine-d5, isolated lignins
and C–DHP (dehydrogenation polymer) were dissolved in 4:1 v/v
DMSO-d6/pyridine-d5, and model compounds were dissolved in
acetone-d6. The central solvent peaks were used as the internal references
(dC/dH: DMSO, 39.5/2.49; acetone, 29.84/2.05 ppm). Standard Bruker
implementations of the traditional suite of 1D and 2D [gradient-selected
and 1H-detected; for example, correlation spectroscopy (COSY), 1H–13C
HSQC (Fig. 1), and heteronuclear multiple-bond correlation (HMBC)]
NMR experiments were used for structural elucidation and assignment
authentication for monomers and dimers. Adiabatic 2D HSQC
(“hsqcetgpsisp2.2”) experiments for ball-milled seed coat material in a
gel state were carried out using the parameters described previously
(22). Processing used typical matched Gaussian apodization in F2
(LB = −0.5; GB = 0.001) and squared cosine-bell apodization in F1.
The characterization of the vanilla seed coat C-lignin was initially
consistent with the previous report (19) but belied some issues. For both
the CW and its EL (derived by removing polysaccharides via crude cellulases treatment) (21), characterization revealed each lignin to be an
almost 100% benzodioxane polymer with only a trace level of the resinol
(b–b) structure. Although not as high as we previously reported (~80%)
(19), the seed coat sample had a very high KL value (~65%). However,
the 2D HSQC NMR of the so-purified lignins contained many peaks in
the aliphatic region that were not from the lignin itself (fig. S1). An
alternative method (below) was therefore required for lignin quantification in these materials.
C-lignin quantification by 13C NMR
Samples for quantitative 13C NMR analysis were prepared by accurately
weighing predried C-LBL samples (100 mg) dissolved in 1-ml internal
standard [1,3,5-trioxane, DMSO-d6 (3.12 mg/ml)] solution. The C-LBL
concentration was also 100.0 mg/ml. Relaxation reagent chromium(III)
Li et al., Sci. Adv. 2018; 4 : eaau2968
28 September 2018
acetylacetonate [Cr(acac)3; ~2 mg] was added to the samples to facilitate
the relaxation of the magnetization. Quantitative 13C NMR spectroscopy was performed as previously described (43). The NMR spectra
were acquired on the 700-MHz spectrometer described above. Relaxation delays were set to be ~5 times the longest T1 values of carbon signals
(for inverse-gated proton decoupled 13C NMR spectra); in our case, d1 =
12.5 s was used to fully relax of all of the carbons with the aid of the relaxation reagent. For the inverse-gated proton-decoupled 13C spectrum,
at least 38 hours (10K scans) were required. Spectral processing used both
Bruker’s TopSpin 3.5pl6 (Mac) and MestreNova 11.0 (Mac) software.
The acquired FIDs were processed typically with a 5-Hz line broadening.
The central solvent peaks were used as the internal references (dC/dH:
DMSO, 39.5/2.49 ppm). Baseline was corrected manually over the
50- to 100-ppm region using TopSpin.
13
C NMR is mostly used to quantify low–molecular weight technical
lignins (such as kraft lignin and organosolv lignin) or milled wood
lignins (43, 44). It is difficult to quantify native lignin with 13C NMR for
two reasons. One is the poor solubility of lignin, and the other is the
overlapping peaks from the lignin side chain with polysaccharide peaks.
However, C-LBL is a perfect sample for 13C NMR analysis. First, the
lignin structure is simple; there is only one type of structure in the lignin
backbone—the benzodioxane derived from b–O–4-coupling. The
chemical shifts of the benzodioxane carbons are unique (75 to 80 ppm),
so that there is little chance of signal overlap with other components.
Second, C-lignin is acid-resistant. Unlike the S-G–type lignins, harsh
acid pretreatment can be applied to C-lignin without destroying the
benzodioxane structure. Thus, we can easily remove the polysaccharides
by acid pretreatment, further minimizing the signal overlap problem.
According to the 2D HSQC spectrum of C-lignin (fig. S1), Ca and
Cb have the potential to allow 13C NMR quantification of the phenylpropanoid unit derived from caffeyl alcohol in the C-lignin (fig. S2). Cg
cannot be used for the quantification because of the signal overlap with
the unknown peaks (dH, 4.00 to 4.35 ppm; dC, 60.0 to 62.5 ppm). The
aromatic region of C-lignin cannot be used for the quantification because of the overlap with signals from protein residues (tyrosine and
phenylalanine) (45). Ca and Cb may seem equally good for the C-lignin
quantification; however, when looking at the HSQC spectrum at a lower
contour level, peaks from polysaccharide residues cannot be completely
ignored even after the acidic LiBr pretreatment; the residual C3 and C5
of the cellulose overlap with the Cb of the C-lignin. Because the relaxation reagent Cr(acac)3 was added to reduce the experiment time, the line
broadening caused by the relaxation reagent made the overlap between
Cb and the cellulose residues even worse. As a result, Ca was chosen for
the quantification as it had minimal peak overlap issues. Assuming that
C-lignin is derived from pure caffeyl alcohol, the detailed calculation
was as shown below (table S2)
cIS 3
ACb
cCb ¼
AIS
cCb
Y CA ¼
rLBL
W ligninðLBLÞ ¼ Y CA MwCA 100%
W ligninðLBLÞ
W ligninðCWÞ ¼
LBL%
In the equations, cIS (mmol/ml) is the molar concentration of internal
standard (IS; 1,3,5-trioxane), AIS is the peak integral of internal standard
in the quantitative 13C NMR spectrum, cCb (mmol/ml) is the molar concentration of caffeyl alcohol unit in the C-lignin polymer, ACb is the peak
integral of Cb in the quantitative 13C NMR spectrum, rLBL (mg/ml) is the
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(42). Briefly, 0.3 g of biomass (weighed to the nearest 0.1 mg) was
treated in 72% (w/w) H2SO4 at room temperature for 60 min. The
slurry was diluted to 4% (w/w) H2SO4 and autoclaved at 121°C for 60 min.
After filtration, the acid-insoluble lignin (AIL = KL) and the acid-soluble
lignin were quantitated gravimetrically and spectrophotometrically,
respectively (table S1). Monosaccharides in the KL filtrates (hydrolysates)
were quantitated using high-performance ion-chromatography on a
Dionex ICS-3000 system equipped with an integrated amperometric
detector and a CarboPac PA1 column (4 × 250 mm) at 30°C. DI water
was used as an eluent at a flow rate of 0.7 ml/min according to the following gradient: 0 to 25 min, 100% water; 25.1 to 35 min, 30% water and
70% 0.1 M NaOH; and 35.1 to 42 min, 100% water. The post-run eluent
of 0.5 M NaOH at a flow rate of 0.3 ml/min was used to purge remaining
materials from the column to ensure baseline stability and detector sensitivity (23). Crude protein content was determined from the nitrogen
(N) content using a 6.25 N-to-protein factor (table S1). The total N
was determined using an elemental combustion system (model 4010,
Costech Analytical Technologies). Samples (approximately 10 mg)
were accurately weighed into tin combustion cups using a microbalance. After complete combustion, total N was measured as N2 gas.
The compositional analysis results are shown in table S1.
SCIENCE ADVANCES | RESEARCH ARTICLE
mass concentration of C-LBL sample, YCA (mmol/mg) is the mole
amount of caffeyl alcohol (CA) per milligram of C-LBL, MwCA
(mg/mmol) is the molecular weight of caffeyl alcohol, Wlignin(LBL) is
the weight percentage of C-lignin in C-LBL, LBL% is the weight percentage of C-LBL obtained from whole CW, and Wlignin(CW) is the
weight percentage of C-lignin in whole CW.
Li et al., Sci. Adv. 2018; 4 : eaau2968
28 September 2018
Analytical methods
GC-MS qualitative analysis of low–molecular weight products
Samples were dissolved in pyridine, and BSTFA was added for TMS
derivatization. The mixture was heated to 50°C for 30 min. An aliquot
of the sample (1 ml) was injected by an autosampler into a GC-MS
(GC2010/PARVUM2, IC-1 column, Shimadzu Co.) equipped with a
fused silica capillary column (30-m × 0.25-mm film; SHR5XLB capillary
column, Shimadzu Co.) operating in split mode (split ratio of 20:1) to
identify the products. The products were identified by comparison with
the peak retention times and mass spectra of the authentic compounds
and (or) by comparing with entries in the National Institute of Standards
and Technology mass spectral library (fig. S5).
GC-FID quantitative analysis of low–molecular
weight products
The identified major products were quantified by GC-FID (GC-2014,
Shimadzu Co.) using calibration curves derived from authentic synthetic
compounds (table S3). The yields of major hydrogenolysis products
catechylpropanol M1 and catechylpropane M2 were quantified by
using the calibration curves generated from their authentic synthetic standards. The yields of minor products without a primary hydroxy group
[chroman-6,7-diol M3, catechol M4, 4-methylcatechol M5, 4-ethylcatechol
M6, and 4-(1-propenyl)catechol M7] were calculated by the effective carbon number (ECN) method based on the yield of catechylpropane M2,
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Lignin depolymerization methods
Alkaline NBO
NBO was performed as previously described (46). Dimeric model compound D1 (5 mg) or extracted vanilla seed coat (40 mg) was mixed with
nitrobenzene (0.4 ml) and 2 M NaOH (7 ml) in a 10-ml stainless steel
reactor vessel (Taiatsu Techno Co.) and heated at 170°C for 2 hours in
an oil bath. The reactor was then cooled in ice water, and 1 ml of freshly
prepared ethyl vanillin (3-ethoxy-4-hydroxybenzaldehyde; 5 mg/ml) in
0.1 M NaOH solution was added to the reaction mixture as an internal
standard. The mixture was transferred to a 100-ml separatory funnel
and washed three times with 15 ml of DCM. The remaining aqueous
layer was acidified with 2 M HCl until the pH was below 3.0 and extracted
with 2 × 20 ml of DCM and 20 ml of diethyl ether. The combined organic
layers were washed with DI water (20 ml) and dried over MgSO4. After
filtration, the filtrate was collected in a 100-ml pear-shaped flask and dried
under reduced pressure. For the TMS derivatization step, NBO products
were transferred with pyridine (3 × 200 ml) into a GC vial, and N,Obis(trimethylsilyl)trifluoroacetamide [BSTFA; 100 ml] was added. The
mixture was heated to 50°C for 30 min. The silylated NBO products
were analyzed by GC-MS and quantified by GC-FID using calibration
curves (fig. S3, A and B).
Thioacidolysis followed by Raney nickel desulfurization
Thioacidolysis was performed as previously described (47). The thioacidolysis reagent was prepared freshly by adding 2.5 ml of EtSH and
0.7 ml of BF3 etherate to a 25-ml volumetric flask containing 20 ml of
distilled 1,4-dioxane and then complemented with dioxane to exactly
25 ml. Freshly made thioacidolysis reagent (4.0 ml) was added to a 5-ml
screw-cap reaction vial containing extractive-free CW (40 mg) or model
compound (15 mg) and a magnetic stir bar. The vial cap was screwed on
tightly, and the vial was kept in an oil bath containing a heating block at
100°C for 4 hours with stirring. After the reaction, the vial was cooled in
an ice-water bath for 2 min. A solution of 4,4′-ethylidenebisphenol in
dioxane was prepared and used as an internal standard. The product
mixture was transferred to a separatory funnel and 10 ml of saturated
NaHCO3 solution, along with internal standard solution, was added.
Then, 5 ml of 1 M HCl solution was added to adjust the pH of the
solution to below 3. The aqueous layer was extracted three times with
20 ml of DCM, and the combined organic phase was washed with
saturated NH4Cl, dried over anhydrous MgSO4, and evaporated under
reduced pressure at 40°C. The resulting products were desulfurized via
Raney nickel. Briefly, the thioacidolysis products were dissolved in 3 ml
of distilled dioxane with 1 ml of Raney nickel 3202 (Sigma-Aldrich)
slurry. The mixture was heated at 80°C for 2 hours. After the reaction,
nickel powder was removed using a magnet, and the reaction mixture
was transferred quantitatively with DCM into a separatory funnel
charged with 10 ml of NH4Cl and 10 ml of DCM. Then, 5 ml of 1 M
HCl solution was added to adjust the pH of the solution to below 3. The
aqueous layer was extracted twice with 10 ml of DCM, and the combined
organic phase was washed with brine, dried over anhydrous MgSO4, and
evaporated under reduced pressure at 40°C. For the TMS derivatization
step, products were transferred with pyridine (3 × 200 ml) into a GC vial,
and BSTFA (100 ml) was added. The mixture was heated to 50°C for
30 min. The silylated thioacidolysis products were analyzed by GC-MS
and quantified by GC-FID using calibration curves (fig. S3, C and D).
Hydrogenolysis
Hydrogenolysis was performed as previously described (7). In cases in
which isolated C-LBL was used as a feedstock, 200 mg of C-LBL was
dissolved in 30 ml of methanol or dioxane/water (9:1, v/v) or THF/
water (96:4, v/v) in a 100-ml high-pressure Parr reactor along with
100 mg of catalyst (5 wt % Pt/C, Pd/C, or Ru/C). The reactor was stirred
with a mechanical propeller and heated via a high-temperature heating
jacket. Once closed, the reactor was purged three times and then pressurized with H2 (40 bar, 4 MPa). The reactor was heated to the desired
temperature and then held at that temperature for the specified residence
time. After the reaction was completed, the reactor was cooled in a water
bath to room temperature. The resulting liquid was filtered through a
nylon membrane filter (0.8 mm, 47 mm; Whatman) and washed with
EtOH. The solvent was removed under reduced pressure at 40°C with a
rotary evaporator. The crude products were dissolved in EtOH and
made up to 10 ml in a volumetric flask. A 1 ml of aliquot was transferred
into three 5-ml vials and then dried under reduced pressure. The dried
samples were used for GC, GPC, and NMR analyses. For GC sample
preparation, the sample was dissolved in 0.9 ml of pyridine and
0.1 ml of BSTFA, incubated at 50°C for 30 min, and then subjected
to GC-FID and GC-MS. For NMR sample preparation (fig. S4A), the
sample was dissolved in 0.6 ml of DMSO-d6/pyridine-d5 (4:1, v/v)
and then transferred to a 5-mm NMR tube for NMR. For GPC sample
preparation (fig. S4B), the sample was dissolved in 1 ml of dimethylformamide (DMF) containing 0.1 M LiBr.
For the cases in which hydrogenolysis was performed directly on the
CW material, 200 mg of preextracted vanilla seed coat was mixed with
30 ml of methanol and 100 mg of the catalyst (5 wt % Pt/C, Pd/C, or
Ru/C). The remaining procedure was performed as described above.
For the cases in which the lignin model compound was used as the
feedstock, a solution of 50 mg of dimer D1 in 30 ml of methanol or
dioxane/water (9:1, v/v) was mixed with 50 mg of the catalyst (5 wt %
Pt/C). The remaining procedure was performed as described above.
SCIENCE ADVANCES | RESEARCH ARTICLE
whereas the minor product with a primary hydroxy group (caffeyl alcohol M8) was calculated on the basis of the yield of catechylpropanol
M1. The theoretical ECN of TMS-derivatized catechol M4 (10.0), 4methylcatechol M5 (11.0), 4-ethylcatechol M6 (12.0), catechylpropane
M2 (13.0), 4-(1-propenyl)catechol M7 (12.9), chroman-6,7-diol M3
(12.0), catechylpropanol M1 (15.5), and caffeyl alcohol M8 (15.4) was
used for the calculation. The ECN contribution of aliphatic carbon 1.0,
aromatic carbon 1.0, olefinic carbon 0.95, primary alcohol −0.5, and
TMS 3.0 was used as described (7, 17, 48). The detailed calculation
was as follows
nmonomer ¼
Amonomer
ECNM1 or M2
nM1 or M2
AM1 or M2
ECNmonomer
nCA ¼ Y CA mLBL
Y monomer ¼
nmonomer
100%
nCA
Li et al., Sci. Adv. 2018; 4 : eaau2968
28 September 2018
Synthetic model compounds and compound authentication
Synthetic methods are fully described in the Supplementary Materials.
SUPPLEMENTARY MATERIALS
Supplementary material for this article is available at http://advances.sciencemag.org/cgi/
content/full/4/9/eaau2968/DC1
Synthetic model compounds and compound authentication.
Calibration curves and NMR spectra.
Fig. S1. 2D HSQC NMR.
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In the equations, nmonomer (mmol) is the molar amount of monomer
in each analyzed sample, Amonomer is the peak area of monomer in the
GC-FID chromatogram, nM1 or M2 (mmol) is the molar amount of M1
or M2 in each analyzed sample based on its calibration curve, AM1 or M2
is the peak area of M1 or M2 in the GC-FID chromatogram, ECNmonomer
is the effective carbon number of monomer, ECNM1 or M2 is the effective
carbon number of M1 or M2, nCA (mmol) is the molar amount of caffeyl
alcohol in the feedstock, YCA (mmol/mg) is the mole amount of caffeyl
alcohol per milligram of C-LBL from the quantitative 13C NMR analysis
(table S2), mLBL (in milligrams) is the weight of C-LBL in the feedstock,
and Ymonomer is the yield of monomer based on the molar amount of
caffeyl alcohol in the feedstock.
Analytical GPC
Molecular weight distributions of lignins were determined by GPC
using a Shimadzu LC20-AD LC pump equipped with a Shimadzu
SPD-M20A UV-vis detector set at 280 nm and a Polymer Standard
Services GPC column and guard column [PSS PolarSil analytical Linear
S, 8-mm inner diameter (ID) × 5 cm and 5-mm particle size → PSS
PolarSil analytical Linear S, 8-mm ID × 30 cm and 5-mm particle size].
The samples and column compartment were held at 40°C during analysis. The mobile phase was DMF with 0.1 M LiBr, and the flow rate was
1 ml/min. Molecular weight distributions were determined using Wyatt
ASTRA 7 software (Wyatt Technology Corporation) via a conventional
calibration curve using a ReadyCal polystyrene kit from Sigma-Aldrich
[catalog no. 76552, M(p) 250-70000].
GPC fractionation of hydrogenolysis product mixtures
Using LBL as a hydrogenolysis feedstock and dioxane/water as the
solvent, the product mixture was dried in vacuo, redissolved in pure
dioxane with sonication, filtered through a PTFE membrane (0.2 mm),
and then subjected to GPC. The GPC conditions here were slightly different from those in the analytical GPC method. For the fractionation,
dioxane was used as the mobile phase instead of 0.1 M DMF/LiBr solution at a slower flow rate (0.3 ml/min) to achieve better fractionation.
Four fractions were separated and collected (fig. S7A). The ultraviolet
(UV) absorption contour map showed that different molecular weight
fractions had completely different UV absorption properties. Because of
peak overlap, each fraction was characterized by using its 2D HSQC
NMR spectra and subtracting the overlapped fractions’ spectra
(fig. S7, B to E; note that f2 was characterized by subtracting f1 and
f3 from f2, f3 was characterized by subtracting f2 and f4 from f3, and
f4 was characterized by subtracting f3 from f4]. As seen in the NMR
spectra, peaks from some nonaromatic components appear in all
fractions (f1 to f4). The molecular weight of these nonaromatic components cannot be measured accurately because of the low GPC resolution and peak tailing, and/or the possibility that these nonaromatic
components have a wide molecular weight distribution. Fractions f1
and f2 were almost identical to each other and contained only traces
of aromatic peaks. The highest molecular weight component(s) in
the product mixture was therefore not from lignin but from other
components in the seeds. Fraction f3 contained the major hydrogenolysis products M1 and M3. This fraction exhibited the strongest UV absorption in the UV contour map, which means that it was the dominant
aromatic-containing mixture in the product. Fraction f4 was the other
major hydrogenolysis product M2, which has a slightly lower molecular
weight compared with M1 and M3. It is inferred that there was a large
amount of high–molecular weight products (the products in f1 and f2),
which are distributed from f1 to f4 because of peak overlap. As these
products lack aromatic rings, they are not from the caffeyl alcohol–
derived phenylpropanoid polymer. Thus, they must be produced from
other components existing in the seed, such as waxes, fatty acids, etc.
These observations support our conclusion that the lignin content
of vanilla seed coats is not determined accurately by KL and other
traditional lignin analytical methods because of these nonextractable,
nonaromatic components.
TOC analysis
A TOC analyzer (TOC-VCPH, Shimadzu Co.) with a solid sample
module (SSM-5000A, Shimadzu Co.) was used to determine the total
carbon content of the vanilla seed coat material and its hydrogenolysis
products, its fractions, and the nonvolatile products. The hydrogenolysis products were dried at 50°C for 30 min to remove EtOH and then
dried at 50°C in a vacuum oven for 30 min to completely remove water
and EtOH. The dried solid samples (20.00 ± 1.00 mg) and hydrogenolysis products were measured as solids.
Using LBL as a hydrogenolysis feedstock and dioxane/water as the
solvent, there was no significant change in the carbon content before
(62.66 ± 0.23 wt %) and after (61.64 ± 0.34 wt %) the reaction (±SD,
n = 2). Solvent degradation products (for example, ethylene glycol,
diethylene glycol, etc.) were detected in the product mixture and identified by GC-MS when dioxane was used as solvent. It is still possible
that some components in the C-LBL can either become volatile or attach
to the catalyst. However, considering that the volatile products (for example, methane, ethane, and hexane) have much higher carbon contents
(~75 to 85 wt %) compared with the solvent degradation products (~35
to 45 wt %), the loss of volatile products while introducing solvent degradation products should cause a significant decrease of carbon content. In
our experiment, we did not observe any carbon content decrease nor did
we observe any weight increase of the catalyst. This result suggested that
the loss of volatile products during work-up and the effect of the solvent
degradation products were negligible and also implied that most of the
carbon-containing compounds were retained in the product mixture.
SCIENCE ADVANCES | RESEARCH ARTICLE
Fig. S2. Quantitative 13C NMR spectrum of C-LBL.
Fig. S3. NBO and thioacidolysis products.
Fig. S4. 2D HSQC NMR and molecular weight distributions.
Fig. S5. GC-MS total-ion chromatograms of hydrogenolysis monomer products.
Fig. S6. Yield and selectivity data.
Fig. S7. GPC fractionation of hydrogenolysis products from LBL.
Table S1. Compositional analysis of vanilla seed coat CWs.
Table S2. Quantitative 13C NMR analysis of C-lignin content in the C-LBL and CW.
Table S3. Monomer yields from hydrogenolysis.
REFERENCES AND NOTES
Li et al., Sci. Adv. 2018; 4 : eaau2968
28 September 2018
Acknowledgments
Funding: Funding was provided by the U.S. Department of Energy (DOE) Great Lakes
Bioenergy Research Center (DOE Biological and Environmental Research Office of Science
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SCIENCE ADVANCES | RESEARCH ARTICLE
DE-FC02-07ER64494 and DE-SC0018409), the DOE Center of Bioenergy Innovation (DE-AC05000R22725) and the Swiss Competence Center for Energy Research: Biomass for a Swiss
Energy Future, through the Swiss Commission for Technology and Innovation grant
KTI.2014.0116. We are also grateful to N. Li and X. Pan (University Wisconsin–Madison) for help
with LiBr solubilization methods and to Y. Mottiar and S. Mansfield (University of British
Columbia, Vancouver, Canada) and R. Vanholme and W. Boerjan (Vlaams Instituut voor
Biotechnologie, Gent, Belgium) for discussions on designing lignins that resulted in recent
papers on this topic. We are also grateful to valuable Science Advances’ reviewer comments.
Author contributions: J.R., J.S.L., H.K., Y.L., L.S., and J.A.D. were responsible for the
conception, planning, and organization of the experiments. D.H.-F. provided the vanilla seed
coat material. Y.T., F.C., and R.A.D. provided information and were involved in discussions
relating to it. Y.L. performed the synthesis of dimer D1 and the synthetic C-lignin and isolated
lignins, performed the hydrogenolysis experiments with L.S., and isolated, quantified, and
identified the products. Y.L. also performed NBO reactions and analysis and sugars analysis.
L.S. and A.H.M. aided in the hydrogenolysis experiments. F.Y. performed the thioacidolysis
analysis and helped with the model compound synthesis. H.K. helped Y.L. perform the
quantitative 13C NMR. J.K.M. helped with the catalyst considerations and carried out the GPC
and molecular weight analysis. Y.L. and J.R. were responsible for NMR and MS data and
interpretation. The manuscript was primarily written by Y.L. and J.R. with critical input from all
coauthors. Figures were prepared by Y.L. and J.R. with support from H.K. Competing
interests: The authors declare that they have no competing interests. Data and materials
availability: All data needed to evaluate the conclusions in the paper are present in the
paper and/or the Supplementary Materials. Additional data related to this paper may be
requested from the authors.
Submitted 26 May 2018
Accepted 22 August 2018
Published 28 September 2018
10.1126/sciadv.aau2968
Citation: Y. Li, L. Shuai, H. Kim, A. H. Motagamwala, J. K. Mobley, F. Yue, Y. Tobimatsu,
D. Havkin-Frenkel, F. Chen, R. A. Dixon, J. S. Luterbacher, J. A. Dumesic, J. Ralph, An “ideal
lignin” facilitates full biomass utilization. Sci. Adv. 4, eaau2968 (2018).
Downloaded from http://advances.sciencemag.org/ on November 6, 2018
Li et al., Sci. Adv. 2018; 4 : eaau2968
28 September 2018
10 of 10
An ''ideal lignin'' facilitates full biomass utilization
Yanding Li, Li Shuai, Hoon Kim, Ali Hussain Motagamwala, Justin K. Mobley, Fengxia Yue, Yuki Tobimatsu, Daphna
Havkin-Frenkel, Fang Chen, Richard A. Dixon, Jeremy S. Luterbacher, James A. Dumesic and John Ralph
Sci Adv 4 (9), eaau2968.
DOI: 10.1126/sciadv.aau2968
http://advances.sciencemag.org/content/4/9/eaau2968
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