ATP-binding cassette-like transporters are involved in
the transport of lignin precursors across plasma and
Yu-Chen Miao and Chang-Jun Liu1
Biology Department, Brookhaven National Laboratory, Upton, NY 11973
Edited by Ronald R. Sederoff, North Carolina State University, Raleigh, NC, and approved November 11, 2010 (received for review June 1, 2010)
Lignin is a complex biopolymer derived primarily from the conden-
sation of three monomeric precursors, the monolignols. The syn-
thesis of monolignols occurs in the cytoplasm. To reach the cell wall
where they are oxidized and polymerized, they must be trans-
ported across the cell membrane. However, the molecular mecha-
nisms underlying the transport process are unclear. There are
conflicting views about whether the transport of these precursors
we know little about what chemical forms are required. Using
isolated plasma and vacuolar membrane vesicles prepared from
Arabidopsis, together with applying different transporter inhibi-
derivatives by these native membrane vesicles. We demonstrate
that the transport of lignin precursors across plasmalemma and
port processes, involving ATP-binding cassette-like transporters.
Moreover, we show that both plasma and vacuolar membrane
vesicles selectively transport different forms of lignin precursors.
In the presence of ATP, the inverted plasma membrane vesicles
preferentially take up monolignol aglycones, whereas the vacuolar
vesicles are more specific for glucoconjugates, suggesting that the
chemical forms in conveying them to distinct sites, and that gluco-
sylation of monolignols is necessary for their vacuolar storage but
not required for direct transport into the cell wall in Arabidopsis.
derived from the condensation of three monomeric pre-
cursors, p-hydroxyphenyl, coniferyl, and sinapyl alcohols (termed
monolignols). Although lignin affords vital structural support to
terrestrial plants and provides hydrophobicity to their vascular
elements, its presence in cell walls constitutes a formidable ob-
stacle for digesting forage crops, pulping, and producing re-
newable biofuels from cellulose and hemicelluloses (1, 2).
Monolignols are synthesized in the cytosol. Thereafter, these
monomeric precursors are exported into the cell wall, where
they are polymerized and integrated into the wall to form p-
hydroxyphenyl, guaiacyl, and syringyl subunits (3). Accordingly,
monolignol transport across plasma membranes is a critical step
affecting the deposition of lignin and the thickening of the sec-
precursors, the molecular mechanisms underlying their sub-
cellularsequestration andextracellulartransportation aresketchy
(3). Earlier investigations in gymnosperms and angiosperms pro-
vided conflicting interpretations. Using [3H]Phe to label the de-
veloping xylem, several studies found that the radiolabel was
associated with the rough endoplasmic reticulum (ER) and the
Golgi body, and also with some vesicles fused with the plasma
membrane (4–6). The potential vesicular trafficking between the
cytosol and plasmalemma in differentiating tracheids of xylem
tissues was also reported (4). These autoradiographic and ultra-
structural analyses engendered the assumption that the lignin
exocytosis. However, Kaneda et al. (7) recently adopted a new
approach to preparing labeled xylem cells of lodgepole pine for
autoradiographic studies. Using cryofixation and freezing sub-
ignin is a complex and irregular biopolymer that is primarily
stitution techniques, they substantially minimized the damage in
sectioned cells, thus preventing the misinterpretation of autora-
diography. Then, feeding dissected xylem tissue with a [3H]Phe
radiotracer and selectively inhibiting phenylpropanoid and pro-
tein biosynthesis by different inhibitors, they discovered that the
radiolabel in the ER-Golgi was primarily incorporated into pro-
and Golgi-vesicle clusters abundant in the developing xylem cells
ER-Golgi-vesicle-mediated exocytosis does not play a major role
in transport of the monolignols (7).
Genetic and chemical analyses demonstrated that lignin bio-
synthesis displays considerable plasticity. Besides the three clas-
sical monolignols, some nontraditional phenolic monomers are
incorporated into lignin under certain circumstances (8, 9). For
example, in a natural cinnamyl-alcohol dehydrogenase (CAD)-
deficient mutant of pine and transgenic tobacco knocked down in
CAD, hydroxycinnamaldehydes were incorporated into lignin
(10). Similarly, a lack of the caffeic acid O-methyltransferase in
a maize bm3 mutant caused the accumulation of 5-hydrox-
yconiferyl alcohol and the buildup of this unusual precursor in
lignin (11). In addition, lignins are frequently acylated with ace-
tate or p-coumarate (12, 13); such acylation implicates the in-
corporation of acylated lignin monomers. The accommodation
of alternative monomers in lignification led to the suggestion of
nonspecific passive diffusion of lignin precursors across the
plasma membrane (14). This notion was supported by the ob-
servation of the in vitro partitioning of lignin monomers or ana-
logs by immobilized liposomes and/or lipid-bilayer discs (15, 16).
Although lignin biosynthesis displays considerable flexibility in
incorporating different monomeric precursors, many studies
note that these monomers are deposited differentially in discrete
regions of particular tissues or cells. For example, lignin in the
cell walls of vessels in birch wood is derived mainly from con-
iferyl alcohol, whereas its fiber wall incorporates both sinapyl
and coniferyl alcohols (17). Similarly, in Arabidopsis stems, the
lignin of the vascular bundle in vessels primarily contains
guaiacyl lignin (from coniferyl alcohol), whereas the inter-
fascicular fibers are enriched in syringyl units (from sinapyl al-
cohol) (18). Moreover, when feeding the labeled monolignols
into the developing xylem, the radiolabeled p-coumaryl alcohol is
preferentially laid down in the middle lamella/cell corners,
whereas coniferyl alcohol is mainly located within the secondary
wall (19). These data suggest that the biosynthesis and deposition
of lignin monomers into cell wall is a highly organized, regulated
process, and that active transportation mechanisms might se-
lectively permit the deposition of the particular monolignols.
Author contributions: C.-J.L. designed research; Y.-C.M. performed research; Y.-C.M. and
C.-J.L. analyzed data; and Y.-C.M. and C.-J.L. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
1To whom correspondence should be addressed. E-mail: firstname.lastname@example.org.
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.
| December 28, 2010
| vol. 107
| no. 52www.pnas.org/cgi/doi/10.1073/pnas.1007747108
mixture. The suspension was layered over a discontinuous sucrose gradient
[10%, 15%, 20%, 25%, 30%, 40%, and 50% (wt/vol) in 20 mM Tris·HCl
buffer (pH 7.6), 1 mM DTT, and 1 mM EDTA] in a 40-mL tube. The tubes were
centrifuged at 100,000 × g for 3 h. Successive 2-mL fractions were collected
from the top of the centrifuge tube, diluted with buffer B, and again
centrifuged at 100,000 × g for another 2 h. The pellets were resuspended in
0.2 mL of buffer B for subsequent assays.
Measurement of Vacuolar ATPase and PPase Activity. Vacuolar ATPase activity
was measured by the method of Ames (43). PPi-dependent H+translocation
by vacuolar membrane vesicles was assayed fluorimetrically at 25 °C using
quinacrine as ΔpH indicator. For details, see SI Materials and Methods.
Transport Activity Assay. For the uptake assay, we modified a method de-
scribed by Zhao and Dixon (31) and Sugiyama et al. (27). The 500-μL assay
mixtures contained 25 mM Tris·Mes (pH 8.0), 0.4 M sorbitol, 50 mM KCl, 5
mM MgATP, 0.1% (wt/vol) BSA, and the indicated concentration of phenolic
substrate. ATP was omitted from the nonenergized controls. Assays were
started by adding the membrane vesicles (50–100 μg of protein) while briefly
agitating the mixture at 25 °C. Batches of the reaction mixture (100 μL) were
removed at various times, and their reactions were terminated with 1.0 mL
of ice-cold washing solution (25 mM Tris·Mes, pH 8.0, 0.4 M sorbitol). The
mixtures underwent vacuum filtration through prewetted nitrocellulose
membrane filters (0.22-μm pore diameter; Millipore). The dried filters,
transferred to 20-mL glass vials containing 0.5 mL of 50% (vol/vol) methanol,
were extracted for 1 h at room temperature in an orbital shaker. The eluate
was analyzed by HPLC. The sample was resolved on a Gemini C18 reverse-
phase column (Phenomenex) in 0.2% acetic acid (A) with an increasing
concentration gradient of acetonitrile containing 0.2% acetic acid (B):
0–20 min, 30% B; 20–25 min, 100% B at a constant rate of 0.8 mL/min.
UV absorption was monitored at 254, 280, and 310 nm using a multiple-
wavelength photodiode array detector (Agilent).
For our tests of transport inhibitors, we preincubated them with the
membrane vesicles for 2 min at the following final concentrations: 1 mM
vanadate (in water), 5 μM verapamil (in DMSO), 150 μM glybenclamide (in
DMSO), 5 μM gramicidin D (in DMSO), 2 μM nigericin (in DMSO), 50 μM ni-
fedipine (in DMSO), 1 mM potassium cyanide (in water), and 1 mM NH4Cl (in
water). Sodium vanadate was depolymerized before use according to Good-
effects, each inhibitor was used at the concentration given for its specific in-
hibitory effect on transporters and pumps, or for disrupting membrane po-
tential as well as ΔpH based on values in the literature. To the 250-μL assay
mixture, 1–2.5 μL of each stock solution was added, where the concentration
of organic solvent was less than 1% (vol/vol). After incubation at 25 °C for 30
min, the transported phenolics were measured as described above. The sta-
tistical analyses on the obtained datasets were carried out with a Student’s
Kinetics of Transport of Monolignol or Its Glucoside. We assessed the kinetics
of transport of coniferyl alcohol by plasma membrane vesicles or of coniferin
by vacuolar vesicles. For details, see SI Materials and Methods.
ACKNOWLEDGMENTS. This work was supported by the Division of Chemical
Sciences, Geosciences, and Biosciences, Office of Basic Energy Sciences of the
US Department of Energy (DOE) through Grant DEAC0298CH10886 to C.-J.L.
Initially, this work was also partially supported by the Office of Biological
and Environmental Research of DOE through the pilot project of biofuel
Scientific Focus Area program (BO148).
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Miao and LiuPNAS
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