Development of a clickable designer monolignol for interrogation of lignification in plant cell walls

Article abstract of DOI:10.1021/bc500411u

Lignin is an abundant and essential polymer in land plants. It is a prime factor in the recalcitrance of lignocellulosic biomass to agricultural and industrial end-uses such as forage, pulp and papermaking, and biofuels. To better understand lignifi catio

Full text of DOI:10.1021/bc500411u

Article
pubs.acs.org/bc
Development of a Clickable Designer Monolignol for Interrogation of
Lignification in Plant Cell Walls
Natalie Bukowski,† Jyotsna L. Pandey,‡,§ Lucas Doyle,† Tom L. Richard,‡,§ Charles T. Anderson,‡,#
and Yimin Zhu*,†
†Department of Chemistry, Altoona College, The Pennsylvania State University, Altoona, Pennsylvania 16601, United States
§
#
‡Center for Lignocellulose Structure and Formation, Department of Agricultural and Biological Engineering, and Department of
Biology, The Pennsylvania State University, University Park, Pennsylvania 16802, United States
S
* Supporting Information
ABSTRACT: Lignin is an abundant and essential polymer in
land plants. It is a prime factor in the recalcitrance of
lignocellulosic biomass to agricultural and industrial end-uses
such as forage, pulp and papermaking, and biofuels. To better
understand lignification at the molecular level, we are
developing a lignin spectroscopic and imaging toolbox on
one “negligible” auxiliary. Toward that end, we describe the
design, synthesis, and characterization of a new designer monolignol, 3-O-propargylcaffeyl alcohol, which contains a
bioorthogonal alkynyl functional group at the 3-O-position. Importantly, our data indicate that this monolignol does not alter the
fidelity of lignification. We demonstrate that the designer monolignol provides a platform for multiple spectroscopic and imaging
approaches to reveal temporal and spatial details of lignification, the knowledge of which is critical to reap the potential of energy-
rich renewable plant biomass for sustainable liquid fuels and other diverse economic applications.
presence of other transport routes.9,14,19 Extensive evidence
INTRODUCTION
■
demonstrates that lignification is tightly regulated in vivo,14,20,21
Lignins are complex and irregular phenolic biopolymers,
derived primarily from monolignols.1−3 Essential to vascular
but how this process is spatially and temporally controlled is
largely unknown. New chemical and biochemical tools are
plants, lignins interweave with polysaccharides to form a strong
sorely needed to address these and other important research
and hydrophobic cell wall matrix, which confers mechanical
questions.
strength and facilitates water transport to support the upright
growth of land plants in a gravitropic environment, and also
helps protect plants against pathogens.3,4 Lignin is a major
factor in the recalcitrance of plant cell walls to degradation,
making processing of plant-derived biomass difficult for a
variety of natural and industrial applications, such as ruminant
forage, pulp and papermaking, and lignocellulosic biofuels. As a
result, lignin has attracted significant research interest as a
target for reduction, enhanced degradability, and/or repurpos-
ing as a valuable coproduct.5−11
Lignification, the biological process of lignin polymerization
and deposition, involves complex metabolic, transport, and
chemical processes. Monolignols are first synthesized in the
cytosol via a multienzyme pathway that starts with phenyl-
alanine.3,12,13 They are then transported across the plasma
membrane to the apoplast, or extracellular space, where they
undergo radical-mediated combinatorial polymerization and
deposition to form insoluble lignin.14,15 Despite much progress
through genetic and structural studies, many molecular details
of lignification remain elusive;3,16 for example, how mono-
lignols are transported across cell membranes is still under
debate.14,15 While recent studies suggest that some monolignol
transport is mediated by ATP-binding cassette-like transport
proteins,17,18 accommodation of a wide range of natural and
nonnatural lignin monomers during lignification indicates the
Spectroscopic and imaging approaches have become
increasingly powerful tools in the quest to understand
biological processes at the subcellular scale.22−26 Such
approaches have been developed to study events involving a
variety of biological polymers, including but not limited to
proteins, DNA, and carbohydrates.23,27−30 Recently, designer
monolignols for lignin research have begun to be developed.
Ralph and co-workers have developed fluorophore-tagged
monolignols that have been used to reveal the molecular
details of lignification.31,32 Theoretically, the same concept can
be extended to develop designer monolignols equipped with
other spectroscopic and imaging tags, which will allow for the
use of complementary spectroscopic and imaging methods to
elucidate the complexities of the lignification process. However,
many tags are larger than the monolignols themselves (Figure
1) and may therefore dramatically alter the biological activity
and subcellular localization of the monolignols to which they
are attached. Evidence that fluorophores can change the
distribution of tagged monolignols has been reported recently
Received: September 1, 2014
Revised: November 14, 2014
© XXXX American Chemical Society
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Figure 1. Chemical structures of several spectroscopic probes and monolignols. (A) Alexa 594, 1, for fluorescence imaging; 1,4,7,10-
tetraazacyclododecane-1,4,7,10-tetraacetic acid, 2, for magnetic resonance imaging; biotin, 3; and a terminal alkyne, 4, which is the tag used in this
paper; (B) coniferyl alcohol (CA), 5; and 3-O-propargylcaffeyl alcohol, 6, the designer monolignol developed in this paper.
in a study in which the authors used fluorescent monolignols to
monitor lignification in vitro and in plant tissue.33
Scheme 1. Synthesis of 3-O-Propargylcaffeyl Alcohol, 6
To develop a lignin spectroscopic and imaging toolbox on
one “negligible” auxiliary, we report the design, synthesis, and
initial application of 3-O-propargylcaffeyl alcohol as a designer
monolignol and versatile analog of coniferyl alcohol (CA) for
analysis of lignification (Figure 1B). Replacing the 3-O-methyl
group with a propargyl group introduces a “negligible”
perturbation, and this small bioorthogonal group serves as a
general platform for different spectroscopic and imaging
approaches.
RESULTS AND DISCUSSION
num hydride to generate the desired 3-O-propargylcaffeyl
alcohol in 92% yield.
■
Design and Synthesis of 3-O-Propargylcaffeyl Alco-
hol. In designing a general platform for experimental
interrogation of lignification, we sought to attach a small
bioorthogonal tag to a monolignol: once incorporated into
lignins, this tag will allow for the use of versatile bioorthogonal
chemistry to introduce various spectroscopic and imaging
probes for further studies.34 Specifically, a terminal alkyne was
selected as a tag of choice because of its small size, its
compatibility with robust click chemistry that has been used to
image glycans in plants and other organisms,30,34−37 and its
potential to generate a significant Raman scattering signal in a
silent region of the cellular Raman spectrum.38,39 To link a
terminal alkyne moiety to coniferyl alcohol, we chose to avoid
making any direct modification of aryl/alkenyl carbons, which
can alter the redox potential of the phenol and thus might have
a significant impact on lignification.40 The γ-O-positions of
monolignols have previously been used as modification sites in
designing new lignin monomers.9,11,41,42 However, due to the
loss of the γ-OH, such modifications can produce tetrahy-
drofuran-type β−β-interunit linkages instead of the typical
β−β-linkages, as previously reported for lignification involving
γ-acylated monolignols.43 Although no monolignol with
modification (other than isotopic ones) at the 3-O-methyl
has previously been investigated as a lignin precursor, we
hypothesized that replacing the 3-O-methyl group with a small
alkyl group would neither significantly alter the redox potential
of the phenol nor introduce new interunit linkages. Following
these considerations, we designed 3-O-propargylcaffeyl alcohol
as a coniferyl alcohol analog for this study (Figure 1B).
Horseradish Peroxidase-Catalyzed Dehydrogenative
Polymerization. As a potential probe for lignification, it is
important to determine whether the designer monolignol is
compatible with lignification, i.e., whether it can undergo
enzyme-initiated oxidation and participate in subsequent radical
coupling, particularly cross-coupling with naturally occurring
monomers into lignins. Indeed, we found that 3-O-
propargylcaffeyl alcohol readily undergoes oxidative coupling
in the conventional horseradish peroxidase (HRP)−H2O2
system. To test the fidelity of 3-O-propargylcaffeyl alcohol in
lignification, we produced synthetic lignins (DHPs) using 3:1
coniferyl alcohol/3-O-propargylcaffeyl alcohol with HRP and
H2O2 in the presence of a catalytic amount (2 mol %) of
methyl p-coumarate. The structural compositions of the DHPs
were determined using 2D heteronuclear single quantum
coherence NMR spectroscopy (HSQC-NMR, Figure 2). The
NMR spectrum of the DHP copolymers with 3-O-propargyl-
caffeyl alcohol is similar to that of DHP polymers generated
with only coniferyl alcohol, with typical β-O-4-, β-5-, and β−β-
substructures as the major interunit linkages. Only two new
peaks, which correspond to the two C−H signals of the
propargyl side chains, were observed in the NMR spectrum of
the DHP copolymers. These results suggest that 3-O-
propargylcaffeyl alcohol is incorporated into synthetic lignins
with the propargyl group intact and does not alter the fidelity of
in vitro lignification.
With the terminal alkyne groups incorporated into lignins,
we next performed Raman spectroscopic analysis on the DHP
copolymers. Although alkynes have been well-characterized as
spectroscopic and imaging tools by coupling with other
reporters through bioorthogonal reactions (see below), we
were also interested in identifying unique spectroscopic
properties of the DHP copolymers that could be useful for
future studies. As expected, the Raman spectrum of the DHP
copolymers with 3-O-propargylcaffeyl alcohol is very similar to
The synthesis of 3-O-propargylcaffeyl alcohol started from
the regioselective alkylation of methyl caffeate (Scheme 1). The
hydroxyl groups of methyl caffeate were deprotonated by
adding excess sodium hydride. The less acidic 3-hydroxyl group
generated the more nucleophilic phenoxide, which was
selectively alkylated to produce methyl 3-O-propargylcaffeate
in 56% yield. The ester was then reduced by diisobutylalumi-
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Figure 2. Aliphatic regions of the HSQC 2D-NMR spectra of (A) DHP copolymers of 3:1 CA/6 and (B) DHP polymers of CA.
that of the DHP polymers containing only coniferyl alcohol,
with the exception of an additional significant scattering peak at
2100 cm−1, which corresponds to the alkynyl stretching
vibration (Figure 3). Interestingly, this peak falls in a cellular
silent region, where most endogenous molecules show no
Raman scattering signals.38,39 We envision that the incorpo-
ration of 3-O-propargylcaffeyl alcohol into lignins should enable
direct detection of lignification in plant cell walls through
Raman imaging, without introducing additional spectroscopic
reporters.24,39
Incorporation in Arabidopsis Tissues. As an initial
example of using our designer monolignol to perform
molecular analysis of lignification in plants, we performed
feeding experiments using live Arabidopsis Col-0 seedlings.
Four-day-old seedlings were incubated in liquid Murashige and
Skoog (MS) medium containing 3-O-propargylcaffeyl alcohol
and/or coniferyl alcohol for 4 h. To demonstrate the flexibility
of 3-O-propargylcaffeyl alcohol as a platform for imaging, we
chose to exploit click-chemistry-enabled fluorescence imaging
for detection of incorporation of the monolignol. After
incorporation, seedlings were treated with Alexa 594-azide in
the presence of copper(II) sulfate and ascorbic acid to
fluorescently tag any incorporated alkynyl groups, followed by
imaging using spinning disk confocal fluorescence microscopy.
Seedlings treated with the designer monolignol displayed
significant fluorescence throughout the root tissues (Figure 4B,
C, E, F), whereas no visible fluorescence was observed in
control seedlings treated with only coniferyl alcohol (Figure 4A
and D). In seedlings treated with 3-O-propargylcaffeyl alcohol,
fluorescence intensity increased progressing from the root tip
to the differentiation zone, with maximum intensity observed in
the late differentiation zone (Figure 4E and F). Seedlings
treated with 3:1 coniferyl alcohol/3-O-propargylcaffeyl alcohol
(Figure 4E) exhibited lower fluorescence intensity than
seedlings treated only with 3-O-propargylcaffeyl alcohol (Figure
4F), the difference being most apparent in the late differ-
entiation zone. Incorporation in these tissues was observed in
similar studies that applied coniferyl alcohol and fluorophore-
tagged coniferyl alcohols in feeding experiments.20,32 These
Figure 3. Raman spectra of synthetic lignins, DHP polymers with CA
(blue trace), and DHP copolymers with 3:1 CA/6 (red trace). The
copolymers (red) show a characteristic alkyne peak at 2100 cm−1. The
two spectra are stacked with the red spectrum shifted by separation of
10 units.
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or the vasculature, which are typically lignifying tissues in
seedling roots.
In principle, the incorporation of 3-O-propargylcaffeyl
alcohol could also be detected by confocal Raman imaging at
2100 cm−1, at which wavelength the terminal alkyne in our
probe shows a significant peak but where biological systems
typically have no scattering signals.24,39 We envision that the
incorporation of the designer monolignol along with click
chemistry will enable the use of other imaging approaches, such
as super-resolution microscopy,44 to study the patterns and
dynamics of lignification in plant cell walls.
To further demonstrate the utility of the designer
monolignol, we performed incorporation experiments in stem
sections of Arabidopsis Col-0 plants to test whether the designer
monolignol could be assimilated into lignified cell walls. Forty-
micron-thick stem cryosections of eight-week-old plants were
incubated with 20 μM 3-O-propargylcaffeyl alcohol with 0.1
mg/mL HRP and labeled with Alexa 594-azide by the copper-
catalyzed click reaction before imaging using confocal
microscopy. The autofluorescence of both preexisting and
newly formed lignins was detected using a 405 nm excitation
laser, whereas incorporation of 3-O-propargylcaffeyl alcohol
was detected by imaging Alexa 594-azide using 561 nm
excitation. All stem sections showed 405-nm-excited autofluor-
escence specifically in lignifying tissues (Figure 5A−C), with
higher autofluorescence intensities in vascular bundles
compared to interfascicular fibers. Only stem sections treated
with 3-O-propargylcaffeyl alcohol showed significant fluores-
cence in the 561 nm channel, with higher fluorescence
appearing in interfascicular fibers than in vascular bundles.
No visible fluorescence was observed in control stem sections
that were treated with only coniferyl alcohol. These data are
consistent with the result observed in the feeding experiment
with live Arabidopsis seedlings, suggesting that lignin auto-
fluorescence does not contribute to fluorescence imaging at 561
nm (also see Figure S2 in the Supporting Information). As seen
in Figure 5D and E, fluorescence imaging at 405 and 561 nm
allows for the differentiation of total (previously existing and
newly formed) and newly formed lignins. Autofluorescence
intensity increased insignificantly upon the addition of
monolignols to the tissue (see Figure S3 in the Supporting
Information), suggesting that it was mainly contributed by pre-
existing lignins in the xylem cells of vascular bundles (Figure
5D) and at the cell corners and middle lamellae in
interfascicular fibers (Figure 5E). Regions of high fluorescence
intensity at 561 nm, which correspond to the locations of newly
formed lignin, were localized in inner wall layers and displayed
barely detectable autofluorescence (Figure 5D,E). Together,
these results indicate that the designer monolignol was
successfully incorporated into lignified cell walls and is a
substantially more sensitive method than autofluorescence for
detecting new lignins. We envision that it will be very useful in
revealing temporal and spatial details of lignification, which is
crucial in understanding the molecular details of this intriguing
process.
Figure 4. Images of 3-O-propargylcaffeyl alcohol’s incorporation into
roots of 4 day old Arabidopsis seedlings. (A,D) control seedlings
treated with 20 μM CA. (B,E) seedlings treated with 5 μM 3-O-
propargylcaffeyl alcohol, 6, and 15 μM CA. (C,F) seedlings treated
with 20 μM 6. Images were collected with a spinning disk fluorescence
confocal microscope using a 561 nm laser at 10% power and 500 gain
with an exposure time of 400 ms. (A−C) Contrast-enhanced mosaics
of contiguous images, starting at the root tip (left) and going through
the late differentiation zone (right), recorded using a 20× objective
(Scale bar, 100 μm). (D−F) Contrast-enhanced maximum intensity
projections of z series recorded at the indicated root zones with a
100× oil-immersion objective (Scale bar, 10 μm).
results demonstrate that our designer monolignol was
successfully incorporated into at least the surface tissues of
roots of four-day-old Arabidopsis seedlings and then fluo-
rescently labeled via a copper-catalyzed click reaction. We note
that copper-catalyzed click labeling is also affected by the
potential of the corresponding dyes to penetrate tissues. While
the similarity to coniferyl alcohol makes it likely that 3-O-
propargylcaffeyl alcohol is able to penetrate deep into the root
tissue, we observed that the penetration of the charged Alexa
594-azide into the interior of Arabidopsis roots was low and no
fluorescence was observed in the endodermis, Casparian strips,
CONCLUSION
■
We have designed, synthesized, and tested 3-O-propargylcaffeyl
alcohol as a designer monolignol and a coniferyl alcohol
surrogate. Our data suggest that this compound is compatible
with lignification and does not alter the fidelity of lignification.
The incorporation of 3-O-propargylcaffeyl alcohol into lignins
gives a unique Raman scattering signal in a cellularly silent
D
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EXPERIMENTAL SECTION
■
General. Coniferyl alcohol was synthesized following
published methods.45 Horeseradish peroxidase (type II, 150−
250 units/mg), Alexa 594-azide, Murashige and Skoog salts,
and Shandon Cryomatrix resin were purchased from Sigma-
Aldrich, Life Technologies, Caisson Laboratories, and Thermo
Scientific, respectively. Other commercial chemicals, including
solvents, were of reagent grade or better, purchased from
Sigma-Aldrich or Alfa Aesar, and used without further
purification. Flash chromatography was performed using silica
gel (230−450 mesh).
Nuclear Magnetic Resonance Spectroscopy. NMR
spectra of small molecules (in acetone-d6) and lignin DHPs
[in dimethyl sulfoxide-d6 (d6-DMSO)] were obtained using
standard Bruker pulse programs on a Bruker DPX-300 (300
MHz) spectrometer and a Bruker AV-III 500 (500 MHz)
spectrometer with a cryogenically cooled gradient probe and
inverse probe geometry (i.e., proton coils closest to sample),
respectively. Spectral processing was performed using Bruker
Topspin 3.1 software. Chemical shifts are reported in parts per
million (ppm) using the central residual solvent peaks as
internal references (δH/δC: acetone-d6, 2.04/29.8 ppm; d6-
DMSO, 2.49/39.5 ppm). Fully authenticated assignments were
made by the usual complement of 1D and 2D methods.46,47
Methyl 3-O-Propargylcaffeate. Methyl caffeate was
prepared by adding caffeic acid (5.01 g, 27.8 mmol) and acetyl
chloride (5.0 mL, 70 mmol) to 100 mL of methanol and
stirring at room temperature overnight. The crude product
(5.38 g) was obtained by removing the solvent under vacuum
and was used in the following reaction without further
purification. To a solution of methyl caffeate (1.91 g, 9.84
mmol) in anhydrous DMSO (300 mL) was added sodium
hydride (60% in mineral oil, 0.806 g, 20.2 mmol) at 0 °C. The
mixture was stirred at ambient temperature until no hydrogen
gas was evolved. The mixture was then cooled to 0 °C before
propargyl bromide (0.87 mL, 9.8 mmol) was added. The
mixture was slowly warmed to room temperature overnight.
The reaction was quenched by adding methanol (10 mL) at 0
°C. The mixture was extracted with ethyl acetate (200 mL × 4).
The combined organic phase was washed consecutively with 1
M hydrochloric acid (200 mL), water (200 mL), and brine
(200 mL); dried over magnesium sulfate; and concentrated
under vacuum. The crude product was then carefully purified
by column chromatography (25% to 40% ethyl acetate in
hexanes), followed by recrystallization from ethyl acetate to
give the desired methyl 3-O-propargylcaffeate (1.28 g, 56%
Figure 5. Autofluorescence (405 nm excitation) and click labeling
(561 nm excitation) in 40-μm-thick sections of lignifying 8-week-old
Arabidopsis stems. (A) Control section treated with 20 μM CA. (B)
Section treated with 20 μM 3-O-propargylcaffeyl alcohol, 6. (C)
Section treated with 20 μM 6 and 20 μM CA. (D) Xylem and (E)
interfascicular fibers (IFFs) of section treated with 20 μM 6 and 20
μM CA. Arrowheads in (A) indicate vascular bundles, with
interfascicular fibers lying between bundles. Images are contrast-
enhanced maximum intensity projections of z series recorded with a
spinning disk fluorescence confocal microscope. (A−C) were recorded
using a 20× objective with a 561 nm laser at 5% power, 150 gain, and
400 ms exposure time and a 405 nm laser at 100% power, 150 gain,
and 400 ms exposure time (scale bar, 100 μm). (D,E) were recorded
using a 63× objective with a 561 nm laser at 5% power, 10 gain, and
400 ms exposure time and a 405 nm laser at 100% power, 10 gain, and
400 ms exposure time (scale bar, 20 μm).
1
yield) as a light yellow powder. H NMR (acetone-d6, 300
MHz) δ: 8.43 (1H, b, OH), 7.60 (1H, d, J = 15.9 Hz, H7), 7.44
(1H, d, J = 1.9 Hz, H2), 7.22 (1H, dd, J = 1.9 and 13.7 Hz,
H6), 6.91 (1H, d, J = 13.7 Hz, H5), 6.41 (1H, d, J = 15.9 Hz,
H8), 4.91 (2H, d, J = 2.4 Hz, propargyl CH2), 3.72 (3H, s,
CH3), 3.12 (1H, t, J = 2.4 Hz, propargyl CH); 13C NMR
(acetone-d6, 75 MHz) δ: 167.7 (C9), 150.4 (C4), 146.5 (C3)
145.4 (C7), 127.1 (C1), 124.7 (C6), 116.7 (C5), 115.7 (C8),
113.6 (C2), 79.4 (propargyl C), 77.3 (propargyl CH), 57.2
(propargyl CH2), 52.0 (CH3).
3-O-Propargylcaffeyl Alcohol. To a solution of methyl 3-
O-propargylcaffeate (1.00 g, 4.56 mmol) in anhydrous
tetrahydrofuran (30 mL) at 0 °C was added diisobutylalumi-
num hydride (DIBAL-H, 1.5 M in toluene, 12.2 mL, 18.2
mmol) over 1 h. The solution was stirred at 0 °C for an
additional 30 min before quenching with ethyl acetate (30 mL).
region. We further demonstrated that, when coupled with Alexa
594-azide using click chemistry, it generates strong fluorescence
at 561 nm that is distinct from lignin autofluorescence and can
be easily imaged. Our design sets up a general platform for
using different spectroscopic and imaging approaches to
interrogate plant cell wall lignification. We anticipate that a
lignin spectroscopic toolbox on one “negligible” auxiliary for
both in vitro and in vivo studies of lignification will shed new
light on this complex and enigmatic polymer.
E
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To the mixture at 0 °C, half-saturated aqueous citric acid
solution (60 mL) was added over a period of 30 min. The
mixture was then stirred at ambient temperature for 3 h. The
aqueous layer was extracted with ethyl acetate (100 mL × 3).
The combined organic phase was washed with brine (100 mL),
dried over magnesium sulfate, and concentrated under vacuum.
The crude product was purified by column chromatography
(50% ethyl acetate in hexanes) to give the desired product
(0.85 g, 92% yield) as a pale-yellow solid. 1H NMR (acetone-d6,
300 MHz) δ 7.99 (1H, b, OH), 7.17 (1H, d, J = 1.8 Hz, H2),
6.93 (1H, dd, J = 1.8 and 8.0 Hz, H6), 6.83 (1H, d, J = 8.0 Hz,
H5), 6.52 (1H, d, J = 15.9 Hz, Hα), 6.24 (1H, dt, J = 5.5 and
15.9 Hz, Hβ), 4.83 (2H, d, J = 2.4 Hz, propargyl CH2), 4.24
(2H, d, J = 5.5 Hz, Hγ), 4.07 (1H, b, OH), 3.08 (1H, t, J = 2.4
Hz, propargyl CH); 13C NMR (acetone-d6, 75 MHz) δ 147.4
(C3), 146.2 (C4), 130.2 (Cα), 130.0 (C1), 128.0 (Cβ), 121.5
(C6), 116.4 (C5), 112.4 (C2), 79.7 (propargyl C), 77.0
(propargyl CH), 63.3 (Cγ), 57.2 (propargyl CH2). HRMS
(ESI) m/z calcd for C12H12O3 [M-H]− 203.0708, found
203.0700.
Procedure for Producing Dehydrogenative Polymers
(DHPs). A solution of 3-O-propargylcaffeyl alcohol (74 mg,
0.36 mmol), coniferyl alcohol (199 mg, 1.10 mmol), and
methyl p-coumarate (5.2 mg, 0.029 mmol) in 5 mL of acetone
was added dropwise to a solution of 10.8 M hydrogen peroxide
(0.134 mL, 1.45 mmol) in 200 mL of sodium phosphate buffer
(0.1 M, pH 6.5). Using a peristaltic pump, the resulting
solution was added over 20 h to 20 mL of sodium phosphate
buffer (0.1 M, pH 6.0) containing 3 mg of HRP. The reaction
mixture was stirred for an additional 4 h. The precipitate was
collected by centrifugation (12 000 g, 30 min), washed with
ultrapure water (40 mL × 2), and dried under vacuum to give
DHPs (0.162 g, 68%) as a yellow powder.
Raman Spectroscopy of DHPs. DHP polymers or
copolymers were freeze-dried before spectrum acquisition.
Raman spectra were collected using a Nicolet 8700 FT-Raman
spectrometer. The dried DHP powder was excited using a
diode-pumped 1064 nm Nd:YAG laser at 100 mW power and
the signal was collected with a liquid nitrogen-cooled
germanium detector. Fourier transform spectra were collected
in the range 250−3500 cm−1 with data spacing of 1.928 cm−1
and were averaged from 500 scans. Raman spectra were
baseline corrected and smoothed using the OMINIC spectra
software (Thermo Scientific).
Incorporation, Labeling, and Imaging of Arabidopsis
Seedlings. Arabidopsis thaliana seedlings of the Col-0 ecotype
grown at 22 °C under 24 h light for 4 days were transferred
from plates containing solid MS medium [2.2 g/L Murashige
and Skoog salts, 0.6 g/L 2-(N-morpholino)ethanesulfonic acid,
8 g/L agar−agar, and 10 g/L sucrose, pH 5.6] to 1 mL liquid
MS medium (MS medium lacking agar−agar) containing either
20 μM 3-O-propargylcaffeyl alcohol or 20 μM 3:1 coniferyl
alcohol:3-O-propargylcaffeyl alcohol. Control seedlings were
added to 1 mL liquid MS containing 20 μM coniferyl alcohol.
Seedlings were incubated in constant light at 22 °C for 4 h.
After incorporation, the seedlings were washed with liquid MS
medium (1 mL × 4) and transferred to 1 mL of click-labeling
solution containing 1 mM ascorbic acid, 1 mM CuSO4, and 0.1
μM Alexa 594-azide in liquid MS. Labeling was performed at 25
°C in the dark with rocking for 1 h. Seedlings were washed with
liquid MS (1 mL × 4) before imaging on a Zeiss Cell Observer
SD spinning disk fluorescence confocal microscope using a 561
nm excitation laser and a 617/73 emission filter with a 20× 0.5
NA air immersion objective or a 100× 1.4 NA oil immersion
objective. Maximum projections of collected z series were
generated using ImageJ, adjusting image brightness equally for
all images to maintain constant relative fluorescence intensities.
Images were collected from a total of three replicate
experiments with five seedlings imaged for each treatment.
Incorporation, Labeling, and Imaging of Arabidopsis
Stem Sections. Middle portions of eight-week-old Arabidopsis
Col-0 ecotype stems with secondary growth were cut into 8
mm pieces, embedded and frozen in Shandon Cryomatrix resin,
cryosectioned into 40-μm-thick sections using a Leica CM1950
cryostat, and placed in water. Sections were then transferred to
1 mL aqueous solutions of 0.1 mg/mL HRP containing either
20 μM 3-O-propargylcaffeyl alcohol or 20 μM 3-O-
propargylcaffeyl alcohol and 20 μM coniferyl alcohol. Control
sections were added to 1 mL of aqueous solutions of 0.1 mg/
mL HRP containing 20 μM coniferyl alcohol. These sections
were incubated at 25 °C for 3 h. After incorporation, the
sections were washed with water (1 mL × 4), transferred to 1
mL of click-labeling solution containing 1 mM ascorbic acid, 1
mM CuSO4, and 1 μM Alexa 594-azide in liquid MS medium
and rocked at 25 °C in the dark for 2 h. Sections were then
washed with water (1 mL × 2), transferred to 1 mL of 96%
ethanol, and rocked for 1 h to remove any unbound monomers
or dyes before washing with water (1 mL × 4). Images were
collected as above, with the addition of a 405 nm excitation
laser and a 450/50 emission filter to image autofluorescence
associated with lignin and a 63× 1.4 NA oil immersion
objective. Maximum projections of z series were generated
using ImageJ, adjusting image brightness equally for all images
to maintain constant relative fluorescence intensities. Fluo-
rescence intensities were quantified as raw integrated intensities
per unit area using ImageJ after using a common threshold for
the images to be compared to select lignified regions. Separate
threshold regions were set for images acquired under the 405
and 561 nm channels. Images were collected from a total of
three replicate experiments with three sections imaged for each
treatment.
ASSOCIATED CONTENT
* Supporting Information
■
S
NMR spectra of all compounds, autofluorescence and click
labeling of untreated Arabidopsis stem sections, quantification
of fluorescence intensities of Arabidopsis stem sections. This
material is available free of charge via the Internet at http://
pubs.acs.org.
AUTHOR INFORMATION
Corresponding Author
*E-mail: yuz30@psu.edu.
■
Author Contributions
Natalie Bukowski and Jyotsna L. Pandey contributed equally to
this work.
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
■
We thank Sarah Kiemle for advice on cryosectioning, which was
performed at the Penn State Microscopy and Cytometry
Facility - University Park, PA. This work is financially supported
by Penn State Altoona Research Development Grant (to Y.Z.)
and the Center for Lignocellulose Structure and Formation, an
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Energy Frontier Research Center funded by the U.S.
Department of Energy, Office of Science, Basic Energy Sciences
under Award # DE-SC0001090 (to T.R. and C.T.A.).
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NOTE ADDED IN PROOF
■
After we submitted this manuscript, Ralph and co-workers
published a similar click chemistry strategy for interrogation of
plant cell wall lignification, in which the designed monolignols
were modified at the γ-O-site instead of the 3-O-position
reported in this work. See: Tobimatsu, Y., Van de Wouwer, D.;
Allen, E.; Kumpf, R., Vanholme, B., Boerjan, W., and Ralph, J.
(2014) A click chemistry strategy for visualization of plant cell
wall lignification, Chem. Commun. doi: 10.1039/C4CC04692G.
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