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Plant Physiol. (1999) 119: 1137-1146
Glycerol Is a Suberin Monomer. New Experimental Evidence for an
Old Hypothesis1
Laurence Moire,
Alain Schmutz,
Antony Buchala,
Bin Yan,
Ruth E. Stark, and
Ulrich Ryser*
Institut für Botanische Biologie, Universität Freiburg,
A. Gockelstrasse 3, CH-1700 Freiburg, Switzerland (L.M., A.S.,
A.B., U.R.); and Department of Chemistry, City University of New York
College of Staten Island, 2800 Victory Boulevard, Staten Island,
New York 10314-6600 (B.Y., R.E.S.)
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ABSTRACT |
The monomer composition of the
esterified part of suberin can be determined using gas
chromatography-mass spectroscopy technology and is accordingly believed
to be well known. However, evidence was presented recently indicating
that the suberin of green cotton (Gossypium hirsutum cv
Green Lint) fibers contains substantial amounts of esterified glycerol.
This observation is confirmed in the present report by a sodium dodecyl
sulfate extraction of membrane lipids and by a developmental study,
demonstrating the correlated accumulation of glycerol and established
suberin monomers. Corresponding amounts of glycerol also occur in the
suberin of the periderm of cotton stems and potato (Solanum
tuberosum) tubers. A periderm preparation of wound-healing
potato tuber storage parenchyma was further purified by different
treatments. As the purification proceeded, the concentration of
glycerol increased at about the same rate as that of
 , -alkanedioic acids, the most diagnostic suberin monomers.
Therefore, it is proposed that glycerol is a monomer of suberins in
general and can cross-link aliphatic and aromatic suberin domains,
corresponding to the electron-translucent and electron-opaque suberin
lamellae, respectively. This proposal is consistent with the reported
dimensions of the electron-translucent suberin lamellae.
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INTRODUCTION |
Suberin and cutin are insoluble, lipophilic biopolymers. Together
with complex mixtures of soluble lipids, they form the protective layers of higher plants: the cuticle of the epidermis and the suberin
layers of the periderm and the exodermis. These cell layers are
diffusion barriers for water and other small, polar compounds (Schönherr, 1976 , 1982; Soliday et al., 1979; Vogt et al., 1983; Riederer and Schreiber, 1995; Kerstiens, 1996; Schreiber et al., 1996).
Cuticle, exodermis, and periderm also constitute the first constitutive
barrier against penetration of the plant body by pathogens (Kolattukudy
1977, 1985, 1987, 1996; Kolattukudy and Köller, 1983; Yang et
al., 1993). Wounding triggers suberization irrespective of the natural
protective covering of a plant organ (Kolattukudy and Soliday, 1985).
However, the complex mixtures of waxy components of a cuticle are a
dynamic system, and physical healing of regions depleted in their wax
may be possible (Grace and van Gardingen, 1996).
Suberins and cutins are considered to be closely related polyesters
composed of fatty acids, differing in their chain length and their
substitution patterns, along with smaller amounts of long-chain
alcohols and hydroxycinnamic acids (Kolattukudy and Agrawal, 1974;
Kolattukudy 1980a, 1980b, 1984; Holloway, 1983, 1984). In his tentative
suberin and cutin models, Kolattukudy (1980a) proposed that suberin, in
contrast to cutin, contains a lignin-like aromatic domain, covalently
linked to the aliphatic suberin domain by ester bonds. The presence of
an aromatic suberin domain was supported by the nondegradative
technique of solid-state C-NMR spectroscopy,
using partially purified suberin preparations from wound-healing potato
tuber slices (Garbow et al., 1989; Stark et al., 1989, 1994). However,
no proof was obtained for the proposed aromatic-aliphatic linkages in
suberin. Using solid-state 13C-NMR spectroscopic
analysis of isotopically enriched and enzymatically purified
wound-healing potato suberin, Bernards et al. (1995) presented evidence
that the polyaromatic domain of suberin is composed mainly of
covalently linked hydroxycinnamic acids rather than of the
corresponding alcohols found in lignin. The chemical-shift data for
isotopically labeled resonances was consistent with significant nonester covalent cross-linking between the phenolic units in suberin.
However, it is not known in which particular cell wall domain these
hydroxycinnamic acids are located (Bernards and Lewis, 1998). Smaller
amounts of syringyl and guaiacyl lignin monomers were identified in
potato wound-healing periderm, and their inter-unit bonding patterns
were determined after thioacidolysis, a procedure that cleaves
alkylaryl ether linkages (Borg-Olivier and Monties, 1993; Lapierre et
al., 1996). The lignin is thought to be located in the middle lamellae
and the primary walls of the periderm (Esau, 1977; Fahn, 1990; Lulai
and Morgan, 1992; Thomson et al., 1995).
As seen in transmission electron micrographs, the suberin layers are
lamellated. The surface parallel lamellation is apparently not affected
by extractions with hot organic solvents (for review, see Ryser and
Holloway, 1985). The chemical basis of the lamellation is not known
(Schmidt and Schönherr, 1982). In many species, perhaps in the
majority, the outer layer of the cuticle also has a lamellate
construction (Jeffree, 1996). In suberin the electron-translucent lamellae are more consistent in thickness than the electron-opaque ones
and measure about 3 nm (Falk and El Hadidi, 1961; Schmutz et al.,
1996). It is generally assumed that the waxes are confined to the
electron-translucent lamellae. On the basis of inhibitor experiments,
it was proposed that the constant thickness of these lamellae is
determined by perpendicularly oriented aliphatic suberin monomers
covalently linked to glycerol (Schmutz et al., 1996).
To date two model systems have been used to study the monomer
composition, structure, and biogenesis of suberin layers: (a) the
ABA-induced formation of a wound periderm at the surface of tissue
slices of potato (Solanum tuberosum L.) tuber
storage parenchyma (Kolattukudy and Dean, 1974), and (b) green cotton
(Gossypium hirsutum cv Green Lint) fibers harvested from
plant-grown seeds or from ovules cultured in vitro (Ryser et al., 1983;
Ryser and Holloway, 1985). The green cotton fiber phenotype is
conditioned by a single dominant gene locus Lg (Kohel,
1985).
A caffeoyl-fatty acid-glycerol ester was isolated from the wax of green
cotton fibers (Schmutz et al., 1993, 1994b), and evidence was presented
indicating that glycerol is also an important constituent of cotton
fiber suberin (Schmutz et al., 1993). The immediate significance of
this finding was obvious: The aliphatic monomers of cotton fiber
suberin can only form linear polymers on their own, whereas glycerol
allows the formation of a three-dimensional cross-linked network, as
suggested by the lamellar ultrastructure and insolubility of suberin.
In the present study we show that the glycerol in the suberin of green
cotton fibers is not a contaminant from the wax or from membrane lipids
and accumulates synchronously with the aliphatic suberin monomers.
Comparable amounts of glycerol were also determined in the stem
periderm of cotton plants. During the purification of the suberin of
the wound-healing periderm of potato tubers, glycerol accumulated at
the same rate as ,-dicarboxylic fatty acids, the most diagnostic
suberin monomers (Matzke and Riederer, 1991). These results suggest
that glycerol may be a monomer of suberins in general. The molar
concentrations of glycerol in the examined tissues are sufficient to
propose a structural role for glycerol in suberin.
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MATERIALS AND METHODS |
Plant Material
Cotton (Gossypium hirsutum L.) cv St 406 and a genotype
(Lg) with green fibers (obtained from GERDAT,
Bouaké, Ivory Coast) were grown in a greenhouse at 25°C to
30°C during the day and at 19°C to 22°C at night. Under these
conditions secondary wall formation in the fibers started at 20 to
25 d after anthesis and the fruit capsules opened at 60 to 70 d after anthesis.
Cotton fibers were detached with tweezers from ovules at different time
points after anthesis. The periderm was isolated from the lower part of
the main stem of the cotton cultivar with white fibers (St 406) by
scraping with a razor blade. Seeds of cv St 406 were evacuated in
distilled water to a pressure of 20 to 27 kPa to remove air, soaked
overnight at 4°C in distilled water, and dissected to separate the
chalazal region from the remaining seed coat and the nucellar cuticle
fraction, containing collapsed nucellar cells, the nucellar cuticle,
and the fringe layer, the inner epidermis of cotton seed coats (Ryser
et al., 1988). The isolated tissues were then immediately frozen in
liquid N2, ground to a fine powder with a mortar
and pestle, and finally lyophilized and weighed.
Extraction of the Purified Cell Walls of Cotton Fibers and
Tissues
The lyophilized cell walls were extracted three times with hexane
at about 40°C (2 × 5 h and 1 × 16 h), three
times with chloroform:methanol (2:1, v/v) at about 60°C (2 × 5 h and 1 × 16 h), and finally two times with methanol
at about 70°C (2 × 5 h) in closed Pyrex tubes. In some
experiments an additional extraction was performed with boiling SDS
(2% [w/v] in distilled water) for 4 h before the standardized solvent extractions. After the SDS extraction the fibers were washed on
glass fiber filters (GF/A, Whatman) with distilled water.
Purification of the Wound-Healing Periderm of Potato Tuber
Parenchyma
According to previously published procedures (Stark and Garbow,
1992; Pacchiano et al., 1993), fresh potatoes (Solanum
tuberosum L. cv Russet Burbank) were cleaned and peeled under
sterile conditions. The tubers were cut into 2- × 20- × 30-mm
sections and aerated in a dark incubator at 25°C for 7 d. The
brown layer of wound periderm on the surface of the potato discs was
collected by blade peeling to streamline the removal of unsuberized
storage parenchyma cells (Bernards et al., 1995). Unsuberized cell wall
materials were removed by standard cellulase (ICN) and pectinase
(Sigma) enzymatic treatments. Subsequently, soluble lipids and wax were removed using a 2:1 (v/v) mixture of methylene chloride:methanol and
exhaustive extraction with a Soxhlet apparatus (Fisher Scientific) for
48 h. Finally, the suberin was extracted with 1,4-dioxane:water (96:4) at room temperature overnight (Ralph et al., 1995) to remove soluble lignins and residual sugars. The resulting pieces of suberized periderm tissue were powdered at liquid N2
temperature in a freezer mill. Dry suberin samples were obtained by
thorough drying in a Speed Vac concentrator (Savant Instruments,
Farmingdale, NY) at 60°C for more than 2 h until there was
no further weight loss. Typically, 1 kg of potato yielded approximately
2 g of dry suberin.
NMR Spectroscopy
All solid-state, cross-polarization, magic-angle spinning NMR
spectra were acquired on a Unity plus spectrometer (Varian Instruments, Palo Alto, CA) operating at a 1H frequency of
300.001 MHz and a 13C frequency of 75.445 MHz.
The experiments were conducted with a 7-mm magic-angle spinning probe
(Varian Instruments) at a regulated temperature of 25°C. Typically,
150 mg of suberin (dry weight) was used. The 90° pulses for
1H and 13C were
both set to 5.5 µs. The rotor-spinning speed was
maintained at 3000 ± 2 Hz by a speed controlling device (Varian).
All 13C chemical shifts were referenced to
tetramethylsilane via hexamethylbenzene as a secondary substitution
reference.
Depolymerization Procedure and GC-MS
The chloroform:methanol extracts and the exhaustively extracted
cell wall residues were depolymerized by acid-catalyzed
transesterification with 5% (w/v) HCl in methanol for 16 h
(extracts) or for 40 h (cell wall residues) at 50°C (Holloway,
1984). The hydrolyzed cell wall residues were washed on glass-fiber
filters with methanol and chloroform:methanol (2:1, v/v), lyophilized,
and weighed. After addition of 5% (w/v) aqueous NaCl solution to the
filtrates and the hydrolyzed chloroform:methanol extracts, the aqueous
layers were removed for the determination of glycerol and the organic layers dried over Na2SO4.
Salt precipitates were removed by filtration. The solvent was
evaporated and the resulting components converted to their
corresponding trimethyl silyl ethers. The same treatment was performed
for potato powders. GC analysis was performed using N2 as the carrier gas at 1 mL
min 1 on a SE 52 Permabond column (25 m, i.d.
0.33 mm, film thickness 0.5 µm; Macherey and Nagel, Duren, Germany)
at a temperature of 140°C for 2 min and then elevated to 280°C at
4°C min1. The ionizing energy of the MS was
70 eV. Before transesterification 1-pentadecanol was added as the
internal standard to the samples. A mixture of ferulic acid, C16
,-alkanedioic acid, C16 -hydroxyalkanoic acid, C18:1 alkanoic
acid, and 1-pentadecanol was used as the external standard.
Determination of Glycerol
The aqueous layer obtained after transesterification was
concentrated under reduced pressure, as necessary, and the glycerol was
quantitated enzymatically (Eggstein and Kuhlmann, 1974) with a
commercial test combination (Boehringer Mannheim). The test couples
glycerokinase (EC 2.7.1.30) with pyruvate kinase (EC 2.7.1.40) and
lactate dehydrogenase (EC 1.1.1.27). Finally, the consumption of NADH
is measured at 340 nm. Glycerokinase phosphorylates glycerol (Km = 60 µmol; pH 9.8; 25°C) and
dihydroxyacetone. However, the conversion of dihydroxyacetone is slow.
After transesterification, dihydroxyacetone can no longer be measured
with the test combination and reduces the measurable concentrations of
glycerol, probably by forming stable homo- and heterodimers. Each test
was made with at least two different volumes of the transesterification
reaction mixture to check the accuracy of the method. In the absence of glycerokinase, no significant amounts of NADH were consumed with the
exception of one sample (the chalazal region of the seed coat), where a
corresponding correction was made. Glycerol was also identified by TLC
in alkaline hydrolysates of exhaustively extracted green cotton fibers
(Schmutz et al., 1993). Other polyols such as mannitol or erythritol
could not be identified in the hydrolysates (A. Schmutz and U. Ryser, unpublished results).
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RESULTS |
Association of Bound Glycerol with the Suberin of Green Cotton
Fibers
The wax fraction of green cotton fibers contains about 10 times
more bound glycerol than the suberin fraction (Schmutz et al., 1993),
yet the monomer composition and the chain-length distribution of the
fatty acids were reported to be surprisingly similar in the wax and in
suberin (Schmutz et al., 1996). An estimation of a possible
contamination of the suberin fraction with wax components was therefore
not possible. In a different approach, the possibility of a
contamination of cotton fiber suberin with glycerides of the wax was
tested by extracting the purified cell walls of the green fibers with
boiling SDS before the conventional wax extraction with organic
solvents. After acid-catalyzed transesterification of the extracts and
the cell wall residues, the liberated glycerol was determined
enzymatically, and the other wax and suberin monomers were
characterized by GC-MS. A single SDS treatment removed the major wax
monomers almost quantitatively (Fig. 1A),
whereas the same treatment had no significant effect on the amount of
suberin monomers, including glycerol (Fig. 1B). This observation
indicates clearly that glycerol is covalently bound to suberin and is
not a contaminant due to unextracted wax or membrane lipids.
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| Figure 1.
Effect of an extraction with boiling SDS (2%,
w/v) on the glycerol content of the wax and suberin fractions of green
cotton fibers. A, Wax fraction; B, suberin fraction. After the SDS
extraction, the fibers were dewaxed with organic solvents as usual. The
remaining wax (A) and suberin (B) were then depolymerized by
acid-catalyzed transesterification and the monomer concentration was
determined as described in ``Materials and Methods''. Gly, Glycerol;
HCA, hydroxycinnamic acids; AA, alkanoic acids, ADA,
,-alkanedioic acids; HAA, -hydroxyalkanoic acids.
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Accumulation of Bound Glycerol and Other Suberin Monomers during
the Development of Green Cotton Fibers
The accumulation of the major suberin and wax monomers, determined
after acid-catalyzed transesterification, was compared with the
increase of fiber dry weight, a good indicator of secondary wall
cellulose deposition. During the time period of primary wall formation,
up to 16 d after anthesis, the rate of dry weight accumulation remained low. It then increased by a factor of about 10 at the onset
of secondary wall formation at about 20 d after anthesis (Fig. 2A).
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| Figure 2.
Accumulation of glycerol and other wax and suberin
monomers during the development of green cotton fibers (mean values of
two independent experiments). A, Accumulation of cell wall dry weight.
Secondary wall deposition began at about 20 d after anthesis, as
indicated by the increased rate of dry weight deposition. B,
Accumulation of glycerol, hydroxycinnamic acids, and alkanoic acids in
the wax fraction. C, Accumulation of glycerol, hydroxycinnamic acids,
and alkanoic acids in the suberin fraction. dpa, Days postanthesis;
HCA, hydroxycinnamic acids; Gly, glycerol; HAA, -hydroxyalkanoic
acids; ADA, ,-alkanedioic acids.
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In the wax, secondary wall formation is accompanied by the accumulation
of glycerol, -hydroxyalkanoic acids, alkanoic acids, ,-alkandioic acids, and hydroxycinnamic acids (Fig. 2B), as well
as smaller amounts of alkanols and long-chain alkanoic acids (Fig.
3A). The accumulation of the wax monomers
is clearly delayed compared with dry weight accumulation. A reduction
of the amount of wax monomers by more than 50% was observed in the
oldest fibers, analyzed shortly before opening of the fruit
capsules. The observed reduction may be due to a partial degradation of
the wax during the phase of fiber maturation.
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| Figure 3.
Accumulation of short- ( C18) and long-chain
(>C18) alkanoic acids in the wax and suberin fractions of green cotton
fibers (mean values of two independent experiments). A, Wax; B,
suberin. dpa, Days postanthesis; AA, alkanoic acids.
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In suberin, glycerol, -hydroxyalkanoic acids, and ,-alkandioic
acids accumulate during secondary wall formation (Fig. 2C), as do
smaller amounts of long-chain alkanoic acids (Fig. 3B). Only traces of
alkanols and hydroxycinnamic acids were determined in the suberin
fraction. As already observed for the wax monomers, the
accumulation of suberin monomers was delayed compared with dry weight accumulation. This delay may be explained by the observation that the first secondary wall layer is always cellulosic, and is
thicker in some fibers than in others. In contrast to the situation in
the wax, no significant reduction in the amounts of monomers was
observed in the suberin fraction of the oldest fibers.
Short-chain (C16 and C18) alkanoic acids accumulated in the wax
and in the suberin fraction at the beginning of secondary wall
formation and then declined again, whereas the longer acids (C20-C26)
accumulated during secondary wall formation (Fig. 3), suggesting that
the short-chain alkanoic acids do not represent genuine suberin
monomers.
The Glycerol Content of the Stem Periderm and of Different Tissues
of Cotton Seeds
Stem periderm and three different parts of mature cotton seeds of
cv St 406 were isolated as described in ``Materials and Methods''.
The chalazal region and a nucellar cuticle fraction were separated from
the remaining seed coat. The nucellar cuticle fraction is composed of
collapsed nucellar cells, the nucellar cuticle, and the innermost epidermis of the cotton seed coat, the so-called fringe layer (Ryser et
al., 1988). In the chalazal region the inner integument is appreciably
thicker than in the rest of the seed coat. The thickened region,
forming a kind of plug, was isolated in mature, hydrated seeds by
separating it from the rest of the seed coat and the fringe layer. A
large portion of the cell walls of the chalazal region could be stained
with phloroglucinol-HCl, probably indicating that these walls are
lignified. Only the innermost, central cells of the chalazal region
were suberized in the studied cultivar, as determined by staining with
a mixture of the lipophilic Sudan III and IV dyes (results not shown).
The purified cell wall powders of the different tissues were
exhaustively extracted with hot organic solvents. The wax fractions and
the polyester part of the cell wall residues were then depolymerized by
acid-catalyzed transesterification. Glycerol was determined enzymatically, and the suberin and cutin monomers were characterized by
GC-MS.
The glycerol content of the wax and suberin fractions, together with
the GC-MS analyses of the suberin fractions, are summarized in Table
I. It is important to remember that
complex tissues were analyzed and that not all of the compounds
detected were necessarily derived from cutin or suberin, but may be
derived from other cell wall layers or from incompletely removed
cellular contents. Smaller amounts of hydroxybenzoic acids were
determined in the seed coats and in the chalaza, probably derived from
the lignified cell walls observed in these tissues by
phloroglucinol-HCl staining.
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Table I.
Glycerol and aromatic and aliphatic suberin and
cutin monomers released after acid-catalyzed transesterification of
extractive free stem periderm and of three different tissues of seeds
of a cotton cultivar with white fibers (St 406)
For comparisons, the glycerol content of the wax fraction is also
indicated.
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Cotton seed coats and green cotton fibers have a qualitatively similar
suberin monomer composition. However, the seed coats have been found
previously to contain higher amounts of 1-alkanols (Ryser and Holloway,
1985). This observation was confirmed in the present study. In
addition, larger amounts of hydroxycinnamic acids and to a lesser
extent of -hydroxyalkanoic acids were determined in the seed coats
(results not shown). Among the hydroxycinnamic acids of the seed coat
(Table I), caffeate was the most prominent monomer (38.4%),
followed by p-coumarate (16.6%) and ferulate (4.7%).
C18 ,alkenedioic acid was determined in the hydrolysates of
the seed coats of the cotton cultivar with white fibers (St 406), but
this monomer was not found in the fibers and seed coats of cv Green
Lint.
The fatty acid composition of the transesterification reaction mixtures
of the four tissues (seed coat, stem periderm, chalazal region, and
nucellar cuticle) was clearly different. Using the discriminant score
of Matzke and Riederer (1991) to distinguish suberin and cutin, typical
suberins dominate in the seed coat, the chalaza, and the periderm,
whereas the nucellar cuticle fraction contains cutin, as expected
(bottom of Table I). It can be deduced from these results that the
chalazal suberin was not significantly contaminated with nucellar cutin
or with suberin from the seed coats, as the predominating chain length
of the ,-dicarboxylic and -hydroxy fatty acids in the chalazal
suberin was C18, whereas in the seed coat and in cotton fibers chain
lengths of C22 predominated.
The suberin of the stem periderm contained about 74 µmol/g glycerol,
15 µmol/g alkanols, 54 µmol/g ,-dicarboxylic fatty acids, and
54 µmol/g -hydroxy fatty acids. Because the wax of the stem
periderm contained about 10 times less glycerol than the cell wall
residue of the same tissue, the glycerol determined in the suberin
cannot be attributed to wax contamination. Assuming that the monobasic
alkanoic and alkenoic acids with chain lengths of up to C18 are not
bona fide suberin or cutin monomers, a conservative estimate can be
made concerning a possible contamination of the purified cell wall
fractions with membrane lipids or triglycerides. Less than 11% of the
glycerol could be attributed to membrane lipids in the seed coat
fraction, and less than 2% in the periderm, whereas in the chalaza
this value could reach 97%. Accordingly, the contamination of the
nucellar cuticle with triglycerides from contaminating endosperm
cells must be less than 50%. It should be stressed, however, that in
addition to the results presented above for green cotton fiber suberin,
no clear-cut scientific evidence is available to rule out the
possibility that short-chain alkanoic and alkenoic fatty acids may be
bona fide suberin monomers.
As in the transesterification reaction mixtures of green cotton fibers
(Fig. 1), roughly equimolar concentrations of glycerol and of the sum
of the ,-dicarboxylic and -hydroxy fatty acids were recovered
in the transesterification reaction mixtures of exhaustively extracted
cell wall powders of the periderm of cotton stems and of suberized and
cutinized tissues of cotton seeds.
Glycerol Accumulation during the Purification of the Suberin of
Wound-Healing Potato Tuber Storage Parenchyma
In a preliminary experiment, 1.2% (of the dry weight) of bound
glycerol was determined in the purified periderm of steam-cooked potato
tubers (Schmutz et al., 1993). In the present experiment, 0.47%
glycerol was determined in the periderm of wound-healing potato tuber
storage parenchyma, and 1.04% in the purified suberin preparation.
Solid-state 13C-NMR spectroscopy (Fig.
4) was used to monitor the purification steps as follows: The cold-water-washed crude periderm preparation was
further purified by enzymatic treatments to remove polysaccharides; the
exhaustive solvent extraction was used to remove waxes; and the
dioxane:water extraction was used to remove soluble lignins and
residual sugars.
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| Figure 4.
Solid-state 13C-NMR spectra showing
the progressive purification of the suberin from the wound-healing
periderm of potato tuber slices. a, Cold-water-washed periderm
preparation showing resonances from suberin, wax, and cell wall
components. b, Periderm preparation treated with cellulase and
pectinase, drastically reducing carbohydrate peaks centered at 72 ppm. c, Suberin fraction after exhaustive extraction of the wax
with organic solvents, reducing mainly the methylene peaks at 33 ppm.
d, Suberin fraction after dioxane:water extraction to remove residual
sugars and soluble lignins, resulting in a better definition of the
carbohydrate peaks. The glycerol resonances expected at 66 and 75 ppm
were obscured by large cell wall peaks near 72 ppm and possibly by
signals from suberin esters of primary alcohols at 65 ppm.
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The crude suberin preparation and the residues obtained after the three
purification steps were each depolymerized by acid-catalyzed transesterification, and the concentration of the liberated suberin monomers was determined as described above. The results of this experiment are summarized in Figure 5.
Glycerol is present in all four suberin fractions, and its proportion
increases at about the same rate as the ,-dicarboxylic fatty
acids, the most diagnostic suberin monomers (Matzke and Riederer,
1991). In the purified suberin fraction (after dioxane:water
extraction), 113 mM glycerol, 244 mM
,-dicarboxylic fatty acids, and 97 mM -hydroxy
fatty acids were determined per gram dry weight. The -hydroxy fatty acids accumulate at different rates than glycerol and
,-dicarboxylic fatty acids during suberin purification. The
reason for this difference is not known. The concentration of
hydroxycinnamic acids drops markedly after the extraction with
pectinase and cellulase and accumulates subsequently at about the same
rate as glycerol and the ,-alkanedioic acids. This result
indicates that hydroxycinnamic acids are covalently linked to the cell
wall layers removed by enzymatic treatments, as well as to suberin. In
a control experiment unsuberized tissue from potato tuber slices was
purified and depolymerized exactly as the suberized periderm
preparations. No glycerol was detected in these preparations. This
observation rules out the possibility that glycerol or
glycerol-containing compounds could artifactually fix to cell walls or
starch during the purification procedures.
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| Figure 5.
Concentration of the major suberin monomers during
the purification of suberin from the periderm of wound-healing potato
tuber slices. A, Glycerol; B, ,-dicarboxylic acids; C,
-hydroxy fatty acids; D, hydroxycinnamic acids. The values in A are
means of three determinations, and the values in B to D are means of
two determinations. The isolated periderm was purified as in Figure 4
by 1, a cold water wash; 2, cellulase and pectinase treatments; 3, exhaustive extractions with organic solvents; and 4, extraction with
dioxane:H2O. The figures above the bars indicate the
relative changes in percent, with the enzyme-treated periderm sample
corresponding to 100%.
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Making a conservative estimate as above, and assuming that all of the
monobasic fatty acids with chain lengths of up to C18 are derived from
membrane lipids, less than 18% of the glycerol in the potato periderm
could be accounted for by membrane lipids (results not shown). Thus,
the presence of glycerol was confirmed in sterile suberin preparations
from wound-healing potato tubers, and the observed glycerol
concentrations were compatible with a structural role of glycerol in
potato suberin.
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DISCUSSION |
Kügler (1884; cited by Gilson, 1890) obtained the first
preliminary evidence for the presence of bound glycerol in the cork of
oak. After extraction of the wax with boiling chloroform, boiling ethanol, and water, suberin was depolymerized with boiling ethanolic KOH for 48 h. The resulting extract contained 2.65% glycerol and 30% fatty acids. Kügler concluded that suberin is a
triglyceride-like substance, rendered insoluble by the presence of
cellulose molecules. Gilson (1890) confirmed the presence of glycerol
in crude oak cork preparations using improved analytical methods.
However, he did not quantify the glycerol content of his preparations
and he firmly rejected the idea that suberin is a triglyceride, as proposed by Kügler. His conclusion was based on the absence of cellulose in the microscopically thin suberin layers and the different solubilities of suberin and triglycerides in organic solvents. Ribas
and Blasco (1940) showed that extractions with water, ethanol, ether,
or chloroform for 5 h in a Soxhlet apparatus did not change the
glycerol content (6.2% of the dry weight) of a finely powdered oak
cork preparation, and high concentrations of bound glycerol were later
reported also to occur in exhaustively extracted Douglas fir bark
(4.65%) by Hergert and Kurth (1952) and again in stopper cork (4.7%)
by Rosa and Pereira (1994). However, Ribas (1942, 1952) was the first
to propose that suberin may be a water-insoluble polyester that can be
easily saponified. According to Ribas, the ease of saponification also
explains why other authors (von Schmidt, 1904; Zetsche and Rosenthal,
1927), using prolonged extractions with weakly alkaline solutions
before saponification, concluded that glycerol was not a suberin
monomer.
With the advent of GC-MS for the identification of organic compounds,
suberin and cutin monomers, or their methylesters, were extracted after
the depolymerization step into organic solvents before further
derivatization and injection into the GC system, leaving small polar
molecules in the aqueous phase. Therefore, at most traces of glycerol
derivatives can be detected in such chromatograms.
The aliphatic monomer composition of the suberin of green cotton fibers
allows only the formation of linear polymers (Ryser at al., 1983; Yatsu
et al., 1983). Similar observations were made for most suberins, as
well as for some cutins (Kolattukudy, 1980a; Matzke and Riederer,
1991). Following the ideas outlined by Kolattukudy, we decided to study
in more detail the phenolic components of cotton fiber suberin. The
resulting purification of a caffeoyl-fatty acid-glycerol ester from the
wax of green cotton fibers by Schmutz et al. (1994b) then prompted us
to test the idea that glycerol may be a suberin monomer, connecting
aliphatic and aromatic suberin constituents.
Taking white cotton fibers as a control tissue, it was readily shown
that glycerol is specifically associated with the suberin layers of the
green fibers (Schmutz et al., 1993, 1996). Changes in the glycerol
concentration of the wax and suberin fractions induced by two specific
inhibitors of suberin biosynthesis were correlated with changes in the
concentrations of the aliphatic suberin monomers (Schmutz et al., 1993,
1994a, 1994b, 1996).
Using boiling SDS for the extraction of the wax of green cotton fibers,
we show in this work that the glycerol content of suberin is not a
contaminant from the wax or from membrane lipids. A developmental study
of differentiating green cotton fibers clearly indicates that glycerol
is incorporated into suberin at the same rate as the established
monomers, -hydroxyalkanoic acids and ,-alkanedioic acids.
Therefore, it can be excluded that glycerol becomes fixed artifactually
to the suberin layers only during the phase of programmed cell death
and dehydration of the fibers.
A comparison of the glycerol content of the periderm of cotton stems
and of wound-healing potato tubers, in relation to the molar
concentration of other suberin monomers, is consistent with our
previous hypothesis that glycerol may be a monomer of suberins in
general. In addition, the glycerol content of the wax of these suberins
is low, indicating again that the glycerol determined in the suberin
fraction cannot be considered as a contaminant from the wax. As in
green cotton fibers, glycerol occurs in these tissues in sufficient
molar proportions to play the expected structural role in suberin. This
possibility could be further tested by the isolation, purification, and
chemical analysis of suberin oligomers.
Glycerol may have a structural role in suberin, linking the aliphatic
and aromatic suberin domains, corresponding to the electron-translucent and electron-opaque suberin lamellae, respectively. This hypothesis is
supported by the dimensions of the electron-translucent suberin lamellae of green cotton fibers (Schmutz et al., 1996). The esterified C22 acids representing 96% of the aliphatic suberin monomers with two
functional groups (,-alkanedioic and -hydroxyalkanoic acids) are about 2.95 nm long. The thickness of the electron-translucent suberin lamellae, determined after tilting the suberin layers into a
position parallel to the electron beam with a computer-controlled precision goniometer, corresponded to 3.4 ± 0.3 nm. Therefore, the electron-translucent suberin layers are somewhat thicker than the
extended fatty acid chains; they could easily incorporate perpendicularly oriented glycerol molecules, cross-linking the extended
aliphatic suberin monomers in the electron-translucent lamellae to the
aromatic monomers confined to the electron-opaque suberin layers.
Alternatively, glycerol might simply function as a plasticizer in the
wax fraction and be more or less randomly incorporated into suberin.
However, the ultrastructure of the suberin layers suggest that suberin
is not a random polymer. The observed lamellation is typical of block
copolymers. These polymers, exhibiting different types of
ultrastructure, are characterized by the presence of two covalently
bound compounds differing in their polarities. The linkages between the
two compounds inhibit phase separation, which typically occurs in
mixtures of lipids of different polarity (Strobl, 1996).
In conclusion, convincing experimental evidence is presented that
glycerol is a suberin monomer, occurring in sufficient molar concentrations to play a structural role in the polymer. The three hydroxyl groups of glycerol are ideally suited to link and stabilize the three axes of lamellated structures. Therefore, the identification of glycerol as a suberin monomer will probably help us to fully understand the chemical basis of the lamellated ultrastructure of this
biopolymer. The identification of a new suberin monomer also creates an
argument for a reevaluation of the biochemical pathways leading to the
biosynthesis of suberin and associated waxes (von Wettstein-Knowles,
1993) as also proposed by Davin and Lewis (1992).
| |
FOOTNOTES |
1
This work was supported by the Swiss National
Science Foundation (grant nos. 31-39648.93 and 31-49305.96 to U.R.) and
by the U.S. National Science Foundation (grant nos. MCB-9406354 and
MCB-9728503 to R.E.S.).
*
Corresponding author; e-mail ulrich.ryser{at}unifr.ch; fax
41-26-300-97-40.
Received July 7, 1998;
accepted December 5, 1998.
| |
ACKNOWLEDGMENTS |
We thank Martine Schorderet for technical assistance and Ruth
Bosshard for cultivating the cotton plants.
| |
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Abstract
Introduction