Characterization of nonderivatized plant cell walls using high-resolution solution-state NMR spectroscopy

Article abstract of DOI:10.1002/mrc.2201

A recently described plant cell wall dissolution system has been modified to use perdeuterated solvents to allow direct in-NMR-tube dissolution and high-resolution solution-state NMR of the whole cell wall without derivatization. Finely ground cell wall material dissolves in a solvent system containing dimethylsulfoxide-d₆ and 1-methylimidazole-d₆ in a ratio of 4:1 (v/v), keeping wood component structures mainly intact in their near-native state. Two-dimensional NMR experiments, using gradient-HSQC (heteronuclear single quantum coherence) 1 -bond ¹³C-¹H correlation spectroscopy, on nonderivatized cell wall material from a representative gymnosperm Pinus taeda (loblolly pine), an angiosperm Populus tremuloides (quaking aspen), and a herbaceous plant Hibiscus cannabinus (kenaf) demonstrate the efficacy of the system. We describe a method to synthesize 1-methylimidazole-d₆ with a high degree of perdeuteration, thus allowing cell wall dissolution and NMR characterization of nonderivatized plant cell wall structures. Copyright

Full text of DOI:10.1002/mrc.2201

Research Article
Received: 13 December 2007
Revised: 10 January 2008
Accepted: 14 January 2008
Published online in Wiley Interscience: 28 March 2008
(www.interscience.com) DOI 10.1002/mrc.2201
Characterization of nonderivatized plant cell
walls using high-resolution solution-state NMR
spectroscopy†
Daniel J. Yelle,a,b∗ John Ralphc,d and Charles R. Friharta
A recently described plant cell wall dissolution system has been modified to use perdeuterated solvents to allow direct
in-NMR-tube dissolution and high-resolution solution-state NMR of the whole cell wall without derivatization. Finely ground
cell wall material dissolves in a solvent system containing dimethylsulfoxide-d6 and 1-methylimidazole-d6 in a ratio of 4 : 1
(v/v), keeping wood component structures mainly intact in their near-native state. Two-dimensional NMR experiments, using
gradient-HSQC (heteronuclear single quantum coherence) 1-bond 13C–1H correlation spectroscopy, on nonderivatized cell wall
material from a representative gymnosperm Pinus taeda (loblolly pine), an angiosperm Populus tremuloides (quaking aspen),
and a herbaceous plant Hibiscus cannabinus (kenaf) demonstrate the efficacy of the system. We describe a method to synthesize
1-methylimidazole-d6 with a high degree of perdeuteration, thus allowing cell wall dissolution and NMR characterization of
c
nonderivatized plant cell wall structures. Copyright ꢀ 2008 John Wiley & Sons, Ltd.
Supplementary electronic material for this paper is available in Wiley InterScience at http://www.interscience.wiley.com/
jpages/0749-1581/suppmat/
Keywords: NMR; HSQC; 1-methylimidazole-d6; dimethylsulfoxide-d6; Pinus taeda; Populus tremuloides; Hibiscus cannabinus; whole cell
wall dissolution; lignin; polysaccharides
Introduction
polymeric matrix (lignin), in which the polysaccharides are
embedded. Thus, it is the lignin that acts as the mortar to support
thepolysaccharides.Theplantcellwall,therefore,mayberegarded
as a highly complex nanocomposite.
Characterizing plant cell wall composition and structure requires a
varietyofapproaches.Nondestructivespectroscopicmethodsthat
can be applied to the cell wall (or whole plant material), including
solid-state NMR, IR, NIR, are often limited because of either poor
resolution and/or poor structural determination. Degradative
methods may be applied to the whole cell wall material, but
study of individual components typically requires their separation.
Improved solid-state NMR has been useful for examining the
intact plant cell walls[1–4]; however, the decreased relaxation
times and low peak dispersion with solid-state NMR limit its use
for thorough characterization of intact polymers. As solution-state
NMR instrumentation and experiments become more developed,
higher sensitivity and resolution permit the detailed structural
analysis of cell wall polymeric components more effectively.
Recently, two solvent systems were developed for cell wall
dissolution to allow quite detailed structural analysis of the entire
cell wall (following ball-milling), without component separation
or degradation via high-resolution solution-state NMR.[5] These
methods inspired the logical modification of one of these solvent
systems to characterize the cell wall following dissolution, i.e.
without derivatization or work-up steps.
Component isolation remains a valuable method to obtain
relatively pure fractions of individual wall polymers or polymer
classes. Isolating lignin, cellulose, and hemicelluloses from wood
has been long known to be a difficult task, since none of these
componentscanberemovedwithouteithersignificantchemicalor
mechanicalmodificationofthecellwall. Manyattemptshavebeen
made to mill the cell wall to extract lignin and polysaccharides.
Bjo¨rkman[6] in 1956 was one of the first to isolate up to 50%
of a ‘native’ lignin from ball-milled spruce via extraction with
aqueous dioxane to give what is called milled-wood lignin. The
carbohydrate-free yield in practice is usually less than 30% of the
total lignin, and is highly dependent on milling efficiency and the
type of plant. Treating ball-milled plant cell walls with a cellulolytic
enzyme that served to hydrolyze the polysaccharides allowed
the isolation of a cellulolytic enzyme lignin,[7] which retained
∗
Correspondenceto:Daniel J. Yelle,USDAForestService,ForestProductsLabora-
tory, One Gifford Pinchot Dr., Madison, WI 53726, USA. E-mail: dyelle@fs.fed.us
Secondaryxylemistheanatomicaltissuedistinguishingawoody
plant from a nonwoody plant. It is the xylem cells, tracheids
(gymnosperms)andlibriformfibersandvessels(angiosperms)that
housethegeneticcapabilitytobiosynthesizelignifiedwoodycells.
On a microscale, the cell wall consists of a middle lamella, primary
wall, and three secondary wall layers (S1, S2, and S3), where S2 is
the most substantial. After the cell wall polysaccharides (cellulose
and various hemicelluloses) are synthesized and assembled,
p-hydroxycinnamyl alcohols are exported to the wall and are
polymerized through radical coupling mechanisms to form a
a
b
c
USDA Forest Service, Forest Products Laboratory, Madison, Wisconsin, USA
Department of Forestry, University of Wisconsin-Madison, USA
US Dairy Forage Research Center, Madison, Wisconsin, USA
d
†
Department of Biological Systems Engineering, University of Wisconsin, USA
This article was published online on 28 March 2008. An error was subsequently
identified and corrected by an erratum notice that was published online only
on 23 April 2008; DOI: 10.1002/mrc.2251. This printed version incorporates the
amendment identified in the erratum notice.
c
Magn. Reson. Chem. 2008; 46: 508–517
Copyright ꢀ 2008 John Wiley & Sons, Ltd.

Nonderivatized plant cell wall NMR
essentially all the lignin, but also a substantial fraction of the
polysaccharides.
that NMI has the capability of disrupting the strong hydrogen
bonding interactions between cellulose and other components
to allow DMSO (a good swelling solvent for woody plants) fur-
ther access to the cell wall structure. In situ acetylation may be
performed to a high degree of substitution so that the resulting
acetylated cell wall material dissolves in CDCl3 for NMR experi-
ments. The cellulose, albeit after the degree of polymerization is
dramatically decreased by ball-milling, from the DMSO/NMI sys-
tem appears to remain intact. This nondegradative dissolution
(followed by acetylation) allows characterization of the ball-milled
whole cell wall to a much higher resolution than is possible with
solid-state NMR.
To learn more about the reactivity or degradation of various cell
wall components, we needed to find a way to dissolve the whole
cell wall, without derivatization, in suitable NMR solvents. Again,
if the whole cell wall is dissolved in a less modified state, chemical
interactions with various components may be characterized via
solution-state NMR. The DMSO/NMI system will fully dissolve the
wholecellwall. Logically, then, utilizingbothperdeuteratedDMSO
and NMI should allow direct NMR analysis of nonderivatized walls,
without the influence of strong solvent resonances. Characterizing
the chemistry of the nonderivatized plant cell wall via gradient
heteronuclear single quantum coherence (HSQC) 1-bond 13C–1H
correlations depicts the plant cell wall polymers in a fairly native
state and, although it is well recognized that ball-milling already
causes significant depolymerization of cell wall polymers; an
advantage is that the native polymeric structures that are often
obscured by traditional derivatization or isolation practices can
be studied. For example, natural acetates, found on syringyl units
of lignin, especially in kenaf bast fibers, are known to acylate a
substantial fraction of the side chains of lignin units (primarily
at the γ position)[23–25] and are masked in NMR spectra by the
current practice of acetylating the cell wall. Also, hemicelluloses
like galactoglucomannan and glucuronoxylan contain acetyl side
groups off their main chain, which should be fully distinguishable
by NMR.
Ourapproachistoleavetheplantcellwallmaterialinaminimally
altered state so as to characterize all components without
requiring extensive component extraction or isolation techniques.
By doing so, we leave the cell wall component polymers essentially
unchanged, thus enabling the characterization of specific cell wall
structures not seen before through NMR.
In this paper, we describe the following: (i) synthesis of
perdeuterated NMI, (ii) dissolution of the plant cell wall in the most
minimally altered state currently possible, and (iii) determination
of chemical shifts of various polymeric plant cell wall structures
via solution-state NMR techniques. The use of these techniques
enables the characterization of specific cell wall components
that may be partially reacted during derivatization or biological
degradation, and allows the approximate quantification of natural
acetates found in lignin and hemicelluloses.
Isolation of polysaccharide components from plant cell walls
typically involves the removal of lignin to obtain a holocellulose.
Maple holocellulose has been isolated with chlorination and
extraction of the lignin with 15% pyridine in ethanol.[8] Sodium
chlorite in acid solution on cell wall material retains hemicelluloses
almost quantitatively.[9–11] Complete delignification, without
significant loss of polysaccharides, is not obtainable by these
methods. Isolation of specific hemicelluloses in the cell wall must
be gently extracted and purified so as to maintain as natural a
state as possible. A neutral solvent, such as dimethylsulfoxide
(DMSO), is useful for extracting holocellulose to obtain a
xylan fraction with little structural change. Alkaline (KOH or
NaOH) solutions containing sodium borate allow extraction
of galactoglucomannans and glucomannans. However, alkali
facilitates deacetylaton of xylans and mannans. Pretreatments,
such as steam exposure (with and without explosion), on aspen
or birch for 10 min and subsequent water extraction, yields
mainly acetyl substituted xylans.[12] To extract the naturally
acetylated mannans, thermomechanical pulp of spruce (2%
consistency) may be suspended in distilled water with stirring at
60 ◦C,[13] while spruce holocellulose may be extracted with boiling
water to obtain the same.[14] Further isolation of holocellulose
components involves alkaline extraction, but the remaining α-
cellulose steadfastly retains some mannan and xylan.
Methodsthatinvolveball-millingarethebasisofseveralcellwall
characterization procedures, but ball-milling itself breaks bonds
and therefore causes structural changes. Knowing the effect that
ball-milling has on plant cell walls is important because milling the
cell wall destroys anatomical information that once existed. It has
been found that the intensity (ball-mill frequency) of milling has a
larger influence on the particle size than prolonging of the milling
time and that the cell corners and middle lamella are the most
resistant layers to milling, whereas the S1 and S2 layers are clearly
fibrillated early in the milling process.[15,16] These studies suggest
that most, if not all, of the extractable cell wall components are
obtained from these secondary cell wall layers, and not from the
whole cell wall.
Nondestructively dissolving plant cell walls to better under-
stand its native chemical structure has been a challenge. Many
investigators examined solvent systems that dissolve cellulose
(e.g. dimethylacetamide/LiCl), because this highly crystalline
component was anticipated to be the most difficult to dis-
solve. With the introduction of newer ionic liquids, compounds
such as 1-N-butyl-3-methylimidazolium chloride[17] and 1-allyl-3-
methylimidazoliumchloride[18] havebeenintroducedforcellulose
dissolution at temperatures >80 ◦C. The underlying benefits of
these solvents are the ease of recycling them because of their
low vapor pressure. However, for the purposes of dissolving the
plant cell wall, dissolution temperatures beyond 50 ◦C begin to
approach the thermal transition points of lignin, thus increas-
ing the probability of degradation. Specific ionic liquids do have
the capability of dissolving plant cell wall material,[19–21] but the
lack of degradative assessments of these systems currently limit
their application in structural studies. Two systems that were
shown to be effective and nondegradative in dissolving ball-
milled plant cell walls have been recently described.[5] DMSO
with tetrabutylammonium fluoride (TBAF),[22] or DMSO with 1-
methylimidazole (or N-methylimidazole, NMI) fully dissolves plant
cell walls at room temperature. It is currently not well understood
how these solvents act to facilitate dissolution. We hypothesize
Experimental
Chemicals
Ruthenium (III) chloride hydrate (35–40% Ru), tributyl phosphite
(94%), and methanol-d4 (99.8% D) were supplied by Acros
Organics (Geel, Belgium). Imidazole (recrystallized), 1,4-dioxane
(98%), palladium on activated carbon (10% Pd), D2O (99.9% D),
chloroform-d (99.8% D), and dimethylsulfoxide-d6 (99.5% D) were
supplied by Aldrich Chemical Company (Milwaukee, WI).
c
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Copyright ꢀ 2008 John Wiley & Sons, Ltd.
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D. J. Yelle, J. Ralph and C. R. Frihart
Synthesis of 1-methyl-d3-imidazole
Plant cell wall preparation for NMR
This synthesis is derived from prior published syntheses.[26,27]
Briefly, imidazole (10.89 g, 160 mmol), tributylphosphite (2.30 g,
9.2 mmol), ruthenium chloride hydrate (0.64 g, 3.1 mmol), and
methanol-d4 (20 g, 555 mmol) were sequentially added to dioxane
(300 ml) in a 2-l stainless steel Parr reactor vessel. The vessel
was sequentially sealed, purged with nitrogen, stirred, heated to
200 ◦C, and pressurized with nitrogen to 44 bar. These conditions
were maintained for 19 hours, then the vessel was cooled to
25 ◦C and depressurized before disassembly. The reaction mixture
was decanted from the spent catalyst and rotary evaporated to
recover the methanol-d4 and to give a crude yellow liquid. The
product was vacuum distilled (with assistance of a microcapillary)
to recover 1-methyl-d3-imidazole as a colorless liquid (9.01 g,
66.2%). GC/MS: m/z (EI) 88 (28.2), 87 (96.0), 86 (148.8), 85 (M+,
100.0). NMR (CDCl3) : δH 6.82 (H-4, s), 6.98 (H-5, s), 7.36 (H-2, s);
δC 119.83 (C-5), 129.11, 128.96 (C-4, d), 137.56 (C-2), 32.39 (CD3,
Ball-milledwoodwaspreparedfrom Pinustaeda sapwood, Populus
tremuloides sapwood, and Hibiscus cannabinus bast fiber (1-year-
old Tainung2 kenaf). Sequentially, 5 g of plant material was
chopped into approximately 2 × 10 mm pieces, placed into a
Retsch (Newtown, PA) MM 301 mixer mill (equipped with two
50-ml stainless steel jars, each with one 25-mm stainless steel
ball bearing at −196 ◦C) for 2 min at 30 Hz. After the mixer
mill, Hibiscus cannabinus was the only species Soxhlet extracted
sequentially with water, 80% ethanol, and acetone to isolate the
cell wall (essentially free of chlorophyll and other extractives). The
wood species were not extracted for reasons we describe in the
results and discussion. Then, 2 g of pulverized plant material was
loaded into a Retsch PM 100 planetary ball-mill (equipped with a
50-ml ZrO2 jar and ten 10-mm ZrO2 ball bearings) and milled for
7 h (600 rpm, 10 min. pause every 20 min.); the jar temperature
remained at <50 ◦C.
septet, 1J13
= 21 Hz).
2
C–
H
Plant cell wall dissolution
Deuterium exchange of 1-methyl-d3-imidazole to give 1-
methylimidazole-d6
To a 5-ml flask, ball-milled plant cell wall material (60 mg) was
added, followed by DMSO-d6 (400 µl) and 1-methylimidazole-
d6 (100 µl)andstirredatroomtemperaturetogiveaconcentration
of 120 mg/ml. A clear solution formed in approximately 3 h,
depending on the sample, but was usually stirred overnight. Using
a 178-mm NMR tube pipette, the viscous solution was transferred
directly to a 5-mm NMR tube.
This exchange is derived from a combination of prior published
methods.[27,28] Briefly, dry palladium catalyst (1 g Pd/C) was well
stirred for 1 hr in a double-necked flask purged with nitrogen
followed by capping one neck of the flask with a 230-mm round
balloon full of hydrogen. A hose, connected to the second neck of
the flask, allowed hydrogen to flow through the flask and through
a water bath at a rate of ∼1 bubble per second via a needle
valve. 1-methyl-d3-imidazole (5.03 g, 57.4 mmol) was dissolved in
25 g of D2O, added to the reduced catalyst, degassed with three
freeze-thaw cycles under slight vacuum, heated with stirring to
100 ◦C and held for 2 h. The product was filtered through a bed
of celite and vacuum distilled to give 1-methyl-d3-imidazole-d
(4.25 g, 82%). In a sealed pressure tube, 1-methyl-d3-imidazole-d
(584 mg, 6.8 mmol) was dissolved in D2O (20 g) and 63 mg of Pd/C
was added. The tube was purged with hydrogen (as previously
described), sealed, heated with stirring to 160 ◦C and held for
24 h. After cooling, the reaction mixture was diluted with H2O
(20 ml), filtered through a 0.2-µm nylon membrane, washed with
H2O (3×10 ml), and vacuum distilled to give 1-methylimidazole-d6
(456 mg, 76%). GC/MS:m/z (EI)89(6.65)88(M+, 100.0), 87(29.9), 86
(17.6). NMR (CDCl3) : δC/ppm 119.03, 119.50, 119.96 (C-5, triplet),
128.42, 128.88, 128.34 (C-4, triplet), 136.82, 137.32, 137.82 (C-2,
NMR, general
For imidazoles in CDCl3, NMR spectra were acquired at 27 ◦C
on a Bruker DRX-360 instrument (1H 360.13 MHz, 13C 90.56 MHz)
fitted with a 5 mm 1H/broadband gradient probe with inverse
geometry (proton coils closest to the sample). Proton and carbon
spectra were referenced to internal tetramethylsilane (TMS) at
0 p◦pm. For plant cell wall samples, NMR spectra were acquired at
27 C on a Bruker DMX-500 (1H 500.13 MHz, 13C 125.76 MHz)
instrument equipped with a sensitive cryogenically cooled
5-mm TXI 1H/13C/15N gradient probe with inverse geometry.
The central solvent peak was used as an internal reference for
plant cell walls, DMSO (δC 39.5, δH 2.49 ppm). All processing and
numerical integration calculations, utilizing single specimen data,
were conducted using Bruker Biospin’s TopSpin v. 2.0 software.
Performing such integrations on NMR data from polymeric
materials is sensitive to crosspeak overlap, thus all integrations
were measured only on intense well-resolved crosspeaks.
triplet), 32.39 (CD3, septet, 1J13
= 21 Hz).
2
C–
H
Routine 1D 1H and 13C NMR spectra
GC/MS of imidazoles
1H and 13C spectra were recorded with 32K data points for a
spectral width of 3094 Hz (8.6 ppm) and 64K data points for a
spectral width of 20 000 Hz (220 ppm), respectively. Exponential
multiplication (LB = 0.3 for 1H, 2.0 for 13C) and one level of
zero-filling was performed prior to Fourier transformation.
All measurements were carried out with a Varian 4000 ion trap
massspectrometer(WalnutCreek,CA)coupledtoaVarian3800gas
chromatographthatwasequippedwithaCTCAnalyticsCombi-Pal
autosampler (Zwingen, Switzerland). A Siltek-passivated single-
gooseneck liner (Restek, Bellefonte, PA) and a RXi-1ms (30 mm
long × 0.25 mm inner diameter, 0.25 µm film thickness, Restek,
Bellefonte, PA) were used. Split (200 : 1) injections (1 µl) were
performed. Operating conditions: injector temperature, 275 ◦C;
carrier gas, helium at 1.0 ml/min; oven temperature program,
40–125 ◦C at 15 ◦C/min. The mass spectrometer was operated
in positive electron ionization mode (70 eV) over a range of m/z
40–125 at the following temperatures: source, 170 ◦C; manifold,
40 ◦C; transfer line, 275 ◦C.
2D HSQC spectra
The standard Bruker pulse implementation (invietgssi) of
the gradient-selected sensitivity-improved inverse (1H-detected)
HSQC experiment[29] was used for acquiring the 2D spectra. The
phase-sensitive HSQC spectra were determined with an acquisi-
tion time of 183 ms using an F2 spectral width of 3501 Hz (7 ppm)
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Copyright ꢀ 2008 John Wiley & Sons, Ltd.
Magn. Reson. Chem. 2008; 46: 508–517

Nonderivatized plant cell wall NMR
in 1280 data points using 88 transients (16 dummy scans) for
each of 400 t1 increments of the F1 spectral width of 16 349 Hz
(130 ppm) (F1 ‘acquisition time’ of 12.2 ms). Processing used a
Gaussian function for F2 (LB = −0.18, GB = 0.005) and a cosine
squared function for F1 prior to 2D Fourier transformation. The
resulting data matrix was 1024 × 1024 points. 13C-decoupling
during acquisition was performed by GARP composite pulses from
the high-power output decoupling channel.
complete (–CD3) N-methylation with a distribution of products
with the ring protons exchanged, as well. For example, the ex-
pected 1-methyl-d3-imidazole (27%) was found in addition to
1-methyl-d3-imidazole-d (40%), 1-methyl-d3-imidazole-d2 (26%),
and 1-methylimidazole-d6 (7%).
After N-methylation, further exchange of the protons on 1-
methyl-d3-imidazole was accomplished. Imidazoles are much less
prone to isotope exchange than their corresponding imidazolium
salts.[35,36] Previous researchers used D2O with the heterogeneous
catalyst palladium (10%) on activated carbon to exchange all
ring protons to give an yield of 91%.[27] In our several attempts
with this method, the only position that exchanged to a high
degree was H(2) with very little H(4/5) exchange. Since palladium
catalysts seem to be quite variable in their activity with this
method, our result prompted the search for a more consistent ring
proton exchange method for H(4/5). Other researchers described
a method of deuterium exchange of heterocyclic compounds
with D2O and heterogeneous Pd/C under 2.5 atm of hydrogen at
160 ◦C.[28] Although they did not study 1-methylimidazole, most
heterocycles studied displayed almost quantitative deuterium
incorporation of aromatic ring protons. Utilizing a somewhat
modified procedure, we achieved quantitative exchange of the
H(4/5) protons, but little H(2) exchange. Therefore, to obtain
high deuterium exchange of 1-methylimidazole ring protons,
we incorporated modifications of both procedures, to give an
overall yield of ∼62% of the fully deuterated material. Figure 2
NMR assignments
All polysaccharide 1H and 13C chemical shift assignments for
the plant species described here were assigned using previous
literature on Populustremula,[30] Linumusitatissimum,[31] and Picea
abies.[14,32,33] Lignin assignments were confirmed with the NMR
database of lignin and cell wall model compounds.[34] Assignment
of several polysaccharide crosspeaks were not attempted here,
but research is underway toward further assignments.
Results and Discussion
Preparing the plant cell walls
The steps in preparing the plant material for dissolution are
crucial for acquiring quality NMR spectra. The existing dissolution
method[5] provided the basis for this program that involves
decreasing the number of steps toward dissolution. The new
method is designed to decrease material losses, eliminate
the acetylation and subsequent isolation steps, decrease the
sample preparation time, and be able to analyze milligram scale
quantities. In particular, if an ethylenediamime tetraacetic acid
(EDTA, chelating) wash step is deemed necessary to remove
the paramagnetic iron from the material (to increase relaxation
behavior for high-resolution spectra), ∼10% wt loss of cell wall
material can result. Also, we wanted to eliminate the extraction
steps, which remove low molecular weight compounds from
the cell wall, but are also capable of disrupting the hydrogen
bond network of polymeric components. The major reason for
analyzing the cell wall in a nonextracted state is for the purposes
of better understanding the reactivity of all plant components
(including low molecular compounds in the cell lumina) toward
derivatization, which is a project currently under investigation.
Figure 1 illustrates the process we used for dissolving the plant
cell walls, beginning with chopped plant material and ending with
ball-milled micron-sized agglomerates for NMR analysis. Through
environmental scanning electron microscope (ESEM) imaging of
several samples, the mixer-milled material gave an estimated
particle size of 100 µm and the ball-milled material was estimated
to be <5 µm prior to dissolution.
1
shows the H spectra of the various stages in perdeuteration of
1-methylimidazole.
Plant cell wall dissolution
Inthesystempreviouslydescribed,[5] theDMSO : NMIratiowas2 : 1
(v/v) with plant cell wall material at a concentration of 40 mg/ml.
The solution becomes completely clear in a matter of 3 h or less,
giving strong evidence of dissolution. For dissolution of plant cell
walls to be directly placed into a 5-mm NMR tube, this would mean
that only 20 mg of plant cell wall material would be dissolved in
0.5 ml. Particularly for 1D 13C NMR, analyzing less than 60 mg of
plant cell walls in 0.5 ml of solvent requires a very high number
of scans (i.e. days rather than hours), especially if the cell wall
component of interest is lignin, typically only ∼20% of the cell
wall. WefoundthattheuseofDMSO-d6 andNMI-d6 inaratioof4 : 1
(v/v), to give a cell wall concentration of 120 mg/ml, was sufficient
for dissolution. Clarity of the solution at this concentration is not
as high, and the solution is also quite viscous; however, such
solutions are still quite suitable for 2D NMR.[37] Spectra reacquired
on identical samples after several months were identical and
showed no degradation.
13C/1H NMR of the plant cell wall
Synthesis of NMI-d6
The spectra described in the following figures contain color-
coded contours, which correspond to their respective colored
structure shown in Fig. 3. In Fig. 4, we show HSQC 1-bond
13C–1H correlation spectra for all the three species investigated:
the gymnosperm Pinus taeda (loblolly pine), the angiosperm
Populus trembuloides (quaking aspen), and the herbaceous plant
Hibiscus cannabinus (kenaf). All the major cell wall components
are depicted, including polysaccharides (aliphatic and anomeric
regions) and lignin (side-chain and aromatic) regions. The
enhanced peak dispersion and sensitivity using a cryogenically
cooled sample probe allows for the potential to fully assign
The dissolution method requires synthesis of the perdeuterated
1-methylimidazole by N-methylation with deuterated methanol
and exchanging the hydrogens at the 2, 4, and 5 positions on
the ring with deuterium. The N-alkylation route we pursued was
based on a ruthenium-trialkylphosphite complex to alkylate acidic
NH groups of azoles.[26] A modification of this procedure uses
methanol-d4, which has been shown to give N-alkylation of imi-
dazole in 49% yield.[27] Alkylation by this method in our lab gave
an isolated yield of 66% (after vacuum distillation with a micro-
capillary nitrogen bleed). GC/MS analysis of the product showed
c
Magn. Reson. Chem. 2008; 46: 508–517
Copyright ꢀ 2008 John Wiley & Sons, Ltd.
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D. J. Yelle, J. Ralph and C. R. Frihart
CD3
D
O
S
N
:
D
CD3
D3C
N
D
4 :1 (v/v)
10 µm
100 µm
10 µm
a
b
c
d
e
Figure 1. A diagram illustrating the nonderivatized dissolution of plant cell walls. Pinus taeda is shown here as an example: a, shavings produced via
conventional planer; b, cryogenically mixer-milled; c, planetary ball-milled; d, dissolution method; e, NMR tube of cell walls in solution.
guaiacyl units, while gymnosperms only have guaiacyl units.
Angiosperm lignins are known to accommodate a higher β-
aryl ether (A) content, a lower phenylcoumaran (B) content, and
have an increased proportion of resinol (C) units compared to
the gymnosperms. It is also quite noticeable that the kenaf
has the lowest lignin content of the three, and consequently
contains a proportionately larger quantity of polysaccharides, as
seenbytheintenseanomericcorrelations. Itisevidentthatmostof
the polysaccharide anomeric correlations are fairly well resolved,
allowing for partial characterization of the H-1/C-1 crosspeaks
of glucan (cellulose), xylan, mannan, galactan, and arabinan.
Here, we focus on assignments of lignin side-chain units, lignin
guaiacyl and syringyl aromatics, naturally acetylated structures,
and polysaccharide anomerics.
Figure 2. Proton spectra of various stages in NMI deuteration: a, 1-
(i) Lignin side-chain units
methylimidazole; b, 1-methyl-d3-imidazole; c, ring deuteration as per
Hardacre et al.[27]; d, ring deuteration as per Esaki et al.[28]; e, ring
deuteration with combination of methods.
Figure 5 shows the remarkably well-resolved lignin side-
chain correlations for all the three species. Depicted are
the major β-aryl ether units (A), phenylcoumaran units
(B), resinol units (C), dibenzodioxocin units (D), cinnamyl
alcohol endgroups (X1), and the lignin methoxyls (–OMe).
The β-aryl ether units are well dispersed. For example, if a
the polymeric structures in the whole cell wall. In the aromatic
region, one of the major differences between angiosperms and
gymnosperms is evident; angiosperms have both syringyl and
OMe
OH
α
O
HO
γ
5
HO
O
γ
β
O
β
γ
β
HO
γ
O
β
β
4
α
O
OMe
α
α
γ
α
O
OMe
A
B
C
D
OH
γ
CH3
β
6
5
6
2
2
α
N
G
S
5
4
2
OMe
MeO
OMe
N
OR
OR
X1
NMI
Xylan acetylated
Not assigned, unresolved
Methoxyl
Mannan acetylated
Polysaccharide anomerics
Figure 3. Key to the chemical structures found in the spectra in Figs 4–7. The structures are: (A) β-aryl ether in cyan, (B) phenylcoumaran in green,
(C) resinol in purple, (D) dibenzodioxocin in red, (X1) cinnamyl alcohol endgroups in magenta, (G) guaiacyl in dark blue, (S) syringyl in fuchsia, and (NMI)
1-methylimidazole in gray. Other structures include: lignin methoxyl in brown, H-2/C-2 & H-3/C-3 correlations for the acetylated structure of β-D-ManpI
in maroon, H-2/C-2 & H-3/C-3 correlations for the acetylated structure of β-D-XylpI in chartreuse, polysaccharide anomerics in orange, and structures
currently not assigned or unresolved in black.
c
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Copyright ꢀ 2008 John Wiley & Sons, Ltd.
Magn. Reson. Chem. 2008; 46: 508–517

Nonderivatized plant cell wall NMR
(c) Kenaf
(b) Aspen
(a) Pine
60
60
70
60
70
70
80
80
80
90
90
90
100
110
120
100
110
120
100
110
120
3
3
3
ppm
7
6
5
4
ppm
7
6
5
4
ppm
7
6
5
4
Figure 4. HSQC spectra from nonderivatized plant cell walls: a, pine; b, aspen; and c, kenaf at 500 MHz. Note that the spectra cover the range for all the
cell wall components – cellulose, hemicelluloses, and lignin. All contour colors can be matched to their respective structures in Fig. 3. Expanded regions
are shown in Figs 5–7.
(a) Pine
(b) Aspen
(c) Kenaf
60
60
70
80
60
70
80
70
80
ppm
ppm
3.0
5.5
5.0
4.5
4.0
3.5
3.0 ppm
5.5
5.0
4.5
4.0
3.5
3.0
5.5
5.0
4.5
4.0
3.5
Figure 5. Aliphatic region of the whole plant cell wall HSQC spectra: a, pine; b, aspen; and c, kenaf. The black contours are currently unresolved, mostly
because of heavy overlapping in these regions. All contour colors and labels can be matched to their respective structure in Fig. 3. As noted in the
text, AβG is any syringyl or guaiacyl coupled to a guaiacyl β-aryl ether, AβS is any syringyl or guaiacyl coupled to a syringyl β-aryl ether, and Aβγ Ac
is a tentative assignment for a β-aryl ether linkage with a γ -acetate. 2-O-Ac-β-D-Manp, H-2/C-2 correlation for the acetylated structure of β-D-ManpI;
3-O-Ac-β-D-Manp, H-3/C-3 correlation for the acetylated structure of β-D-ManpI[32]; 2-O-Ac-β-D-Xylp, H-2/C-2 correlation for the acetylated structure of
β-D-XylpI; 3-O-Ac-β-D-Xylp, H-3/C-3 correlation for the acetylated structure of β-D-XylpI[30,31]
.
guaiacyl or syringyl unit couples with a guaiacyl at the β-
position of a β-aryl ether linkage (AβG), the Aβ correlation is
shown at 4.37/84.0 ppm.[34] This correlation clearly separates
from the correlations where a guaiacyl or syringyl unit
couples with a syringyl at the β-position (AβS) shown
at 4.19/86.3 ppm and 4.02/87.7 ppm.[34] The pine, having
exclusively guaiacyl units, displays strong phenylcoumaran
correlations(Bα,5.48/87.4 ppm;Bβ,3.46/53.5 ppm),[34] which
are only visible in the aspen and kenaf at lower contour levels
(notshown).Syringylstructures,withtheadditionalmethoxyl
group at the 5 position on the ring, prevent the formation
of linkages such as the phenylcoumaran and the 5–5-
linked portion of dibenzodioxocins. Resinol correlations (Cα,
4.64/85.3 ppm; Cβ, 3.05/53.9 ppm; Cγ1, 4.20/71.5 ppm)[34]
are shown to be more significant in the aspen and kenaf than
in pine. Generally, Cγ correlations are only visible in isolated
milled-wood lignins; the Cγ2 correlation remains masked
by polysaccharides. The dibenzodioxocin correlations Dα
and Dβ in pine are shown here to reside at 4.86/83.6 ppm
and 3.90/85.8 ppm.[34] The cinnamyl alcohol endgroup
c
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Copyright ꢀ 2008 John Wiley & Sons, Ltd.
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D. J. Yelle, J. Ralph and C. R. Frihart
correlation X1γ is shown at 4.14/61.7 ppm[34] and is visible
in all the three species. Spirodienone/β − 1 structures were
detected in the aspen spectra only; however, because of the
low quantity of such linkages in lignin (about 1.8% in aspen
lignin),[38] the spectrum showed these only at low contour
intensity. Spirodienone correlations Sα, Sβ, and Sβꢁ (not
shown) were found at 5.14/81.4 ppm, 2.81/60.3 ppm, and
4.18/79.8 ppm.[38,39] Lignin methoxyls, typically a strong and
broad contour, show some separation between the guaiacyl
and syringyl methoxyls in the aspen and kenaf spectra.[34]
(ii) Lignin guaiacyl and syringyl aromatics
to the acetate methyls (see supplementary material). We
integrated these acetate contour regions in all the spectra
and compared them to those of the methoxyl correlations.
The values obtained show that for pine, aspen, and kenaf,
the ratio of methoxyl to acetate was 1 : 0.1, 1 : 0.3, and
1 : 0.6, respectively. Knowing that the syringyl (β-aryl ether
type) units of kenaf are ∼60% acetylated,[43] and that
syringyl units predominate the lignin portion, an 1 : 0.6
ratio of methoxyl : acetate is a reasonable estimate and
an indication that a large portion of the acetates are on
lignins. Furthermore, a distinctive difference between the
kenaf acetate contour compared to those in aspen and
pine is an extended contour region at 1.75/20.3 ppm. Aspen
syringyl units are only ∼4% acetylated[43,44]; acetylated
xylans may contribute largely to the acetate content. The
major component of angiosperm hemicelluloses is an
O-acetyl-4-O-methylglucurono-β-D-xylan (∼16% of aspen
and ∼13% of kenaf cell wall),[45] where about 7 out of
10 xylose residues contain an O-acetyl group at C-2 or
C-3.[46] The 2-acetylated xylan (2-O-Ac-β-D-Xylp) and the
3-acetylated xylan (3-O-Ac-β-D-Xylp) H-2/C-2 and H-3/C-3
correlations (shown in Fig. 5) are depicted at 4.56/73.8 ppm
and 4.85/75.2 ppm, respectively, in aspen and kenaf.[30,31]
The relative ratio of 3-O-Ac-β-D-Xylp: 2-O-Ac-β-D-Xylp
was estimated from contour integration to be 1 : 1.12
in aspen and 1 : 1.29 in kenaf. Previous literature on an
isolated 4-O-methylglucuronoxylan in aspen shows that the
2-O-acetylated xylose units are approximately one-third less
abundant than the 3-O-acetylated xylose units.[30] However,
O-acetyl migration during isolation methodologies makes
this estimation difficult to ascertain.[47] For the pine, no data
exists to support acetates on guaiacyl lignin. Thus, the ac-
etates are expected to be on the mannans (∼10% of loblolly
pine hemicelluloses),[45] in particular galactoglucomannan,
the principal hemicellulose component in gymnosperms.
For the angiosperms, glucomannans are a secondary hemi-
cellulose component, comprising ∼2% of aspen and kenaf
hemicelluloses.[45] The C-2 and C-3 positions of the mannan
chain are partially substituted with O-acetyl groups in a
frequency of about 1 out of 3–4 hexose units.[46] The acety-
lated mannan (2-O-Ac-β-D-Manp and 3-O-Ac-β-D-Manp)
H-2/C-2 and H-3/C-3 correlations (shown in Fig. 5) are evi-
denced at 5.33/71.0 ppm and 4.85/73.8 ppm.[32,33] Previous
researchers have shown that the ratio of 2-O-Ac-β-D-Manp:
3-O-Ac-β-D-Manp in Norway spruce is not equal, with the
2-O-Ac-β-D-Manp slightly predominating (e.g. 2.2 : 1, 1.7 : 1,
and 1.1 : 1).[14,32,48] Here, through volume integration of the
above contours, we report a ratio of 1.5 : 1 for loblolly pine.
(iv) Polysaccharide anomerics
In Fig. 6, we show the lignin aromatic regions for the
three species. Some interesting differences between the
aspen and the kenaf spectra can be seen. In the aspen
spectra, it is evident that the syringyl : guaiacyl ratio is
much lower than with the kenaf; the pine thus showing
exclusively guaiacyl units. With this dissolution method,
since we are viewing the whole cell wall, we have the
capability of estimating the syringyl : guaiacyl (S : G) ratio
in the aromatic region of the lignin. Figure 6 shows the
syringyl (S) and guaiacyl (G) correlations of the aromatic
C–H groups in four basic conglomerates: S-2/6, G-2, G-
5/6, and G-5. Previous literature suggested that the S : G
ratio in kenaf and aspen lignin to be between 4.3–7.8 and
1.6–2.2, respectively.[40–42] Through volume integration of
the S (S-2/6) and G (G-2, G-5/6, G-6) contours of the kenaf
and aspen cell wall HSQC spectra, we determined the ratios
to be 4.6 and 2.0, respectively, which falls into the expected
ranges. The syringyl correlations shown are from β-aryl ether
(A) and resinol (C) lignin units, while the guaiacyl correlations
are from β-aryl ether (A), phenylcoumaran (B) and resinol
(C) lignin units. The correlation at 7.07/120.5 ppm is from the
residually protonated H-5/C-5 of 1-methylimidazole solvent,
which, even though greatly suppressed, is still visible in the
aromatic region.
(iii) Naturally acetylated structures
An interesting feature of kenaf lignin is its extensive
acetylation of syringyl units with all the acetate on the
γ -position of the β-aryl ether side chain. Thus, it appears
that approximately two-thirds of the β-aryl ether units
are acetylated.[23,43] The Aγ -acetate contours, undoubtedly
present in the kenaf spectrum, are unfortunately found in
a region highly overlapping with polysaccharide contours
of the spectrum (4.15–4.45, 63.2 ppm), shown in Fig. 5. We
cannot, using these spectra, assign any clear correlation
to these acetylated Aγ ’s; however, the extended contour
in this region suggests its existence. Logically, if the γ -
OH is acetylated, an Aβ correlation around 4.62/84.6 ppm
should be present (compound 3071, Ref. [34]). In the
syringyl Aβ region for aspen and kenaf, there are two
contours, 4.19/86.3 ppm and 4.02/87.7 ppm, as discussed
previously. Aspen shows some guaiacyl Aβ correlation
(4.37/84.0 ppm); however, in the kenaf spectrum a strong
correlation exists in a similar region where a guaiacyl Aβ
typically resides (4.41/83.4 ppm). As noted in Fig. 6, kenaf
contains few guaiacyl units. Hence, this strong correlation is
consistent with the Aβ on γ -acetylated β-aryl ether units,
e.g. likely evidence of the Aγ acetate. Thus, the kenaf Aβ at
4.41/83.4 ppm has been tentatively labeled Aβγ Ac in Fig. 5.
Another contour of interest for acetate quantification is the
acetate methyl group. All the three species studied show
a large contour centered at 1.99/20.9 ppm corresponding
In Fig. 7, we show the anomeric region for all the three
species tentatively assigning several anomerics, including
D-glucan, D-xylan, D-mannan, and D-galactan as the major
contours. All the three spectra show internal anomerics
of the (1→4)-linked β-D-glucopyranoside (β-D-GlcpI)
at 4.38/103.1 ppm,[31–33] as well as the (1→4)-linked
β-D-xylopyranoside (β-D-XylpI) at 4.32/102.1 ppm[30]
and (1→4)-linked β-D-mannopyranoside (β-D-ManpI) at
4.57/100.7 ppm.[32,33] The pine depicts anomeric correlations
of the terminal α-D-galactopyranoside (α-D-GalpT), (1→6)-
linked to β-D-Manp, at 4.84/99.0 ppm.[32,33] The pine and
kenafspectraalsoshowanomericcorrelationsofaterminalβ-
D-galactopyranoside (β-D-GalpT), (1→6)-linked to β-D-Manp,
c
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Copyright ꢀ 2008 John Wiley & Sons, Ltd.
Magn. Reson. Chem. 2008; 46: 508–517

Nonderivatized plant cell wall NMR
(a) Pine
(b) Aspen
(c) Kenaf
105
110
115
120
105
110
115
120
105
110
115
120
S-2/6
S-2/6
G-2
G-2
G-2
G-5/6
G-5/6
G-5/6
G-5
G-5
G-5
NMI
NMI
NMI
7.4
7.2
7.0
6.8
6.6
ppm
7.4
7.2
7.0
6.8
ppm
7.4
7.2
7.0
6.8
6.6
ppm
6.6
Figure 6. Aromatic region of the whole plant cell wall HSQC spectra: a, pine; b, aspen; and c, kenaf. The guaiacyl and syringyl contours are a combination
of free phenolic and etherified aromatic units. Contour colors and labels can be matched to their respective structure in Fig. 3.
(b) Aspen
(a) Pine
(c) Kenaf
95
95
95
4-O-MeGlcA
4-O-MeGlcA
4-O-MeGlcA
100
105
100
105
100
105
ppm
ppm
ppm
4.2
5.4 5.2 5.0 4.8 4.6 4.4 4.2
5.4 5.2 5.0 4.8 4.6 4.4 4.2
5.4 5.2 5.0 4.8 4.6 4.4
Figure 7. Anomeric region of the whole plant cell wall HSQC spectra: a, pine; b, aspen; and c, kenaf. Assignments are as follows: β-D-GlcpI, internal β-D-
glucopyranoside units (cellulose)[31–33]; β-D-ManpI, internal β-D-mannopyranoside units[32,33]; β-D-XylpI, internal β-D-xylopyranoside units[30]; α-D-GalpT,
terminal α-D-galactopyranoside units[32,33]; β-D-GalpT, terminal β-D-galactopyranoside units[33]; α-L-ArafT, terminal α-L-arabinofuranoside units[31,32,50]
;
2-O-Ac-β-D-Manp, anomeric correlation for the acetylated H-2/C-2 structure of β-D-ManpI; 3-O-Ac-β-D-Manp, anomeric correlation for the acetylated
H-3/C-3 structure of β-D-ManpI[32]; 2-O-Ac-β-D-Xylp, anomeric correlation for the acetylated H-2/C-2 structure of β-D-XylpI; 3-O-Ac-β-D-Xylp, anomeric
correlation for the acetylated H-3/C-3 structure of β-D-XylpI[30,31]; 4-O-MeGlcA, 4-O-methyl-α-D-glucuronic acid[30]; α and β-D-GlcpR, reducing ends of
glucopyranoside[32,33]; β-D-ManpR, reducing end of mannopyranoside[32]; α and β-D-GalpR, reducing ends of galactopyranoside[33]; α and β-D-XylpR,
reducing ends of xylopyranoside.[30]
.
at 4.32/105.7 ppm,[32,33] which is absent in aspen. Acety-
lated mannan structures show their respective anomeric
correlations at 4.78/99.1 ppm (2-O-Ac-β-D-Manp) and
4.69/100.0 ppm (3-O-Ac-β-D-Manp),[32] while the acetylated
xylan structures show their respective anomeric correlations
at4.53/99.8 ppm(2-O-Ac-β-D-Xylp)and4.44/102.0 ppm(3-O-
Ac-β-D-Xylp).[30,31] Arabinoglucuronoxylan, 5–10% of the cell
wall weight in gymnosperms, is a framework of (1→4)-linked
β-D-XylpI units that are substituted at C-2 by 4-O-methyl-
α-D-glucuronic acid (4-O-MeGlcA) groups in a frequency of
two residues per ten β-D-XylpI units.[46] The 4-O-MeGlcA
anomeric correlations for pine are depicted at 5.15/97.8 ppm.
However, the 4-O-MeGlcA in the aspen and kenaf, (1→2)-
linked in glucuronoxylan, depict their anomeric correlations
c
Magn. Reson. Chem. 2008; 46: 508–517
Copyright ꢀ 2008 John Wiley & Sons, Ltd.
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D. J. Yelle, J. Ralph and C. R. Frihart
at 5.20/97.7 ppm and 5.25/97.7 ppm.[30] This variation in
δH may be caused by the presence of the carboxylic acid;
chemical shift changes in the DMSO-d6/NMI-d6 solvents
would be expected. The arabinoglucuronoxylan is also sub-
stituted with terminal (1→3)-linked α-L-arabinofuranoside
(α-L-ArafT) units in a frequency of 1.3 residues per ten
β-D-XylpI units.[46] The galactans of angiosperms are known
to contain rhamnose and arabinose units, e.g. giving a
molar ratio of 1.7 : 1 : 0.2 (Gal : Ara : Rha) in sugar maple.[49]
Thus, the α-L-Araf anomeric correlations for pine, aspen,
and kenaf are depicted at 4.83/108.4 ppm.[31,32,50] Reducing
ends of the polysaccharides can also be seen in the spectra
and are tentatively assigned here. The reducing α and β
ends of D-Glcp residues are shown at 4.94/92.7 ppm and
4.40/96.9 ppm.[32,33] For pine, the reducing α end of D-Manp
and the reducing α and β ends of D-Galp residues are shown
at 4.97/94.0 ppm,[32,33] 4.98/92.8 ppm and 4.31/97.7 ppm.[33]
For aspen, the reducing α and β ends of D-Xylp residues are
shown at 4.98/94.2 ppm and 4.33/97.8 ppm.[30] In the kenaf
spectrum, only the reducing β end of D-Xylp is observed.
Acknowledgements
We would like to thank especially Dr. Fachuang Lu, Dr. Hoon Kim,
Dr.TakuyaAkiyama,andDr.PaulSchatz(U.S.DairyForageResearch
Center, Madison, WI) for assistance in establishing methodology
and for very beneficial discussions on lignin chemistry and plant
cell wall dissolution. Also, we would like to thank Kolby Hirth
(USDA Forest Products Laboratory, Madison, WI) for GC/MS data
analyses of the imidazoles. Partial funding to J.R. through the
Office of Science (BER), U.S. Department of Energy, Interagency
agreement No. DE-AI02-06ER64299 is gratefully acknowledged.
NMR experiments on the Bruker DMX-500 cryoprobe system were
carriedoutattheNationalMagneticResonanceFacilityatMadison,
WI, which is supported by National Institutes of Health grants
P41RR02301 (Biomedical Research Technology Program, National
Center for Research Resources) and P41GM66326 (National
Institute of General Medical Sciences). Additional equipment
was purchased with funds from the University of Wisconsin, the
National Institutes of Health (RR02781, RR08438), the National
Science Foundation (DMB-8415048, OIA-9977486, BIR-9214394),
and the U.S. Department of Agriculture.
Conclusions
References
The methodologies developed here help in characterizing
the structures of several native-state plant cell wall compo-
nents. Through the effective synthesis of perdeuterated 1-
methylimidazole to a high degree of deuterium incorporation,
we described an approach to dissolve ball-milled wood and
characterize the majority of the plant cell wall components via
high-resolutionsolution-stateNMR. Todirectlydissolveball-milled
cell wall material in nondegradative solvents and run NMR of these
samples allows characterization without derivatization. The HSQC
1-bond 13C–1H correlation spectra of pine, aspen, and kenaf de-
pict all major plant cell wall components with excellent resolution,
dispersion, and in their near-native state. Through these spectra,
weassignedcorrelationstonaturallyacetylatedmannanandxylan
structures found in galactoglucomannan, glucomannan, and glu-
curonoxylan. The existence of a β-aryl ether γ -acetate, believed
to exist in kenaf lignin, has been explored further through the
tentative assignment of Aβγ Ac. The well-dispersed contours in the
polysaccharide anomeric region of the HSQC spectra allowed us to
tentatively assign several correlations in hemicelluloses. Through
contour integration techniques, we estimated the ratios of cell
wall structures. For example, literature values for the S : G ratio
and acetate content match our data fairly closely. We assume
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possibly applying determined response factors, will allow more
accurate quantification in the future. More research needs to be
done to define polysaccharide chemical shifts; most chemical shift
data obtained for assignments were derived from oligosaccha-
rides dissolved in D2O, and not in DMSO-d6/NMI-d6. However, this
research broadens the utility of the DMSO/NMI method to better
understand plant cell wall chemistry, which may be applied to
various fields. We are currently employing these methods to inves-
tigate chemical modification, microbiological decay mechanisms,
and genetically engineered plants and trees.
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