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Running head:
The Arabidopsis DESPERADO/AtWBC11 Transporter is
Required for Cutin and Wax Secretion
Corresponding author: Asaph Aharoni, Department of Plant Sciences, Weizmann
Institute of Science P.O. Box 26, Rehovot 76100, Israel. Tel.: +972 8 934 3643; Fax:
+972 8 934 4181; E-mail: [email protected]
Research category: Biochemical processes and Macromolecular Structures
Plant Physiology Preview. Published on October 19, 2007, as DOI:10.1104/pp.107.105676
Copyright 2007 by the American Society of Plant Biologists
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The Arabidopsis DESPERADO/AtWBC11 Transporter is
Required for Cutin and Wax Secretion
David Panikashvili,1 Sigal Savaldi-Goldstein,2 Tali Mandel,1 Tamar Yifhar,1 Rochus B.
Franke,3 René Höfer,3 Lukas Schreiber,3 Joanne Chory2 and Asaph Aharoni1*
1Department of Plant Sciences, Weizmann Institute of Science, P.O. Box 26, Rehovot
76100, Israel
2Plant Biology Laboratory and Howard Hughes Medical Institute, The Salk Institute, La
Jolla, CA 92037, USA 3Institute of Cellular and Molecular Botany (IZMB), Department of Ecophysiology,
Kirschallee 1, University of Bonn, D-53115 Bonn, Germany
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1A.A. is an incumbent of the Adolfo and Evelyn Blum Career Development Chair and
D.P. is a recipient of the Israeli Ministry of Science Eshkol fellowship for post-docs. The
research in A.A. laboratory was supported by a grant from the William Z. and Eda Bess
Novick Young Scientist Fund and the Y. Leon Benoziyo Institute for Molecular
Medicine. 2S.S-G was supported by a fellowship from BARD and The Salk Institute and
by a grant from the National Research Initiative of the USDA Cooperative State
Research, Education and Extension Service to J.C. 2J.C. is an investigator of the Howard
Hughes Medical Institute. 3The work of L.S. and R.F. was supported by a grant from the
DFG (German Research Society; SCHR506/7-1).
*Corresponding author: Asaph Aharoni
Tel.: +972 8 934 3643
Fax: +972 8 934 4181;
E-mail: [email protected]
The author responsible for the distribution of materials integral to the findings presented in
this article in accordance with the policy described in the Instructions for Authors
(www.plantphysiol.org) is Asaph Aharoni, [email protected]
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ABSTRACT
The cuticle fulfills multiple roles in the plant life cycle including protection from
environmental stresses and the regulation of organ fusion. It is largely composed of cutin,
which consists of C16-18 fatty acids. While cutin composition and biosynthesis has been
studied, the export of cutin monomers out of the epidermis has remained elusive. Here we
show that DESPERADO (AtWBC11), encoding a plasma membrane localized ABC
transporter, is required for cutin transport to the extracellular matrix. The desperado
mutant exhibits an array of surface defects suggesting an abnormally functioning cuticle.
This was accompanied by dramatic alterations in the levels of cutin monomers.
Moreover, electron microscopy revealed unusual lipidic, cytoplasmatic inclusions in
epidermal cells, disappearance of the cuticle in postgenital fusion areas and altered
morphology of trichomes and pavement cells. We also found that DESPERADO is
induced by salt, ABA and wounding stresses and its loss-of-function results in plants that
are highly susceptible to salt and display reduced root branching. Thus, DESPERADO is
not only essential for developmental plasticity but also plays a vital role in stress
responses.
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INTRODUCTION
One of the most critical adaptations of plants to a terrestrial environment 450 million
years ago was the formation of their surface, the cuticle. The cuticular layer plays
multiple roles in plants including the regulation of epidermal permeability and non-
stomatal water loss, protection against insects, pathogens, UV light and frost (Sieber et
al., 2000). It also functions in normal plant developmental processes including the
prevention of postgenital organ fusion and pollen-pistil interactions (Lolle et al., 1998).
The major component of the cuticle is cutin that is a polyester insoluble in organic
solvents, consisting of oxygenated fatty acids with a chain length of 16 or 18 carbons.
Embedded in the cutin matrix are cuticular waxes, which are complex mixtures of very
long chain fatty acid (VLCFA; >C24) derivatives: aldehydes, ketones, primary and
secondary alcohols, fatty acids and wax esters (Kunst and Samuels, 2003). In many
species they also include triterpenoids and other secondary metabolites, such as sterols,
alkaloids, phenylpropanoids and flavonoids. The cuticular waxes are arranged into an
intracuticular layer in close association with the cutin matrix, as well as an epicuticular
film exterior to this, which may include epicuticular wax crystals (Jetter et al., 2000).
Recently, 2-hydroxy- and α,ω-dicarboxylic fatty acids have been reported as the
characteristic monomers of cutin in Arabidopsis (Bonaventure et al., 2004; Franke et al.,
2005). This cutin monomer composition is similar to the aliphatic domain present in the
Arabidopsis suberin polymer (Franke et al., 2005). Suberin is part of the plant apoplastic
barrier which prevents uncontrolled nutritional and water loss, strengthens cell walls and
provides protection from pathogens. In Arabidopsis, suberin depositions were detected in
the endodermis of primary roots and the periderm of mature roots.
Postgenital fusion is a unique phenomenon which occurs when alterations in
cuticle properties cause augmentation of the contact responsiveness. During plant
development organ fusion is tightly regulated and the cuticle plays a vital role in the
either preventing or permitting fusions. Postgenital organ fusion occurs most commonly
during reproductive development as for example during carpel formation in
Angiosperms. One of the characteristic features of organ fusions is that adhesion of cell
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walls is often accompanied by disappearance of the cuticle in the contact area (Lolle et
al., 1998).
To date, nearly twenty Arabidopsis mutants displaying postgenital fusions have
been identified and only for less than a half of them has a corresponding gene product
been associated (Lolle et al., 1998; Tanaka and Machida, 2006). One of the most known
organ fusion mutant is the Arabidopsis fiddlehead (fdh). The FIDDLEHEAD gene
encodes a lipid biosynthetic enzyme that acts through the fatty acids elongation pathway
and might be involved in cutin monomers biosynthesis. The fdh mutant leaves supported
wild-type pollen germination on their surfaces and showed increased permeability of the
cuticle to the toluidine blue dye. In addition, fdh mutants exhibited enhanced rate of
chlorophyll leaching from leaves submerged in alcoholic solution (Lolle and Cheung,
1993; Yephremov et al., 1999; Pruitt et al., 2000). Another mutant, abnormal leaf shape 1
(ale1), showed defective cuticle in embryos and juvenile plants and as a result exhibited
excessive water loss and organ fusion. The corresponding gene belongs to a large family
of subtilisin-like serine proteases in Arabidopsis that are typically involved in
intercellular signaling, converting their substrates to active or inactive forms (Tanaka et
al., 2001). The phenotypes of the ale1 mutants depend on the genetic background and
they could be observed in the Landsberg erecta background but not in the Columbia and
Wassilewskija (Ws) ecotypes backgrounds (Watanabe et al., 2004). Double mutant of
ale1 (in Ws) and the Arabidopsis homolog of crinkly4 (acr4) resulted in half of the
seedlings showing deformed cotyledons and severely fused leaves. The authors suggested
that ACR4 and ALE1 synergistically affected the epidermis and that ACR4 plays a major
role in the differentiation of epidermal cells in both vegetative and reproductive tissues.
The maize crinkly4 (cr4) mutation shows graft-like fusions between organs and the CR4
gene encodes a putative receptor kinase that might generate a signal for epidermal cells
differentiation (Becraft et al., 2001; Jin et al., 2000; Tanaka et al., 2002).
A cytochrome P450 monooxygenase, CYP86A8, catalyzes the ω–hydroxylation of
C12-18 fatty acids when assayed in-vitro. The CYP86A8 loss-of-function mutant, lacerata
(lcr), showed severe cuticle defects as evidenced by epidermal ruptures and postgenital
fusions (Wellesen et al., 2001). A different gene, HOTHEAD (hth), putatively encoding
an oxidoreductase was suggested to be involved in the formation of α,ω-dicarboxylic
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fatty acids since the hth-12 mutant allele showed decreased load of these acids
(Kurdyukov et al., 2006b). In the hth mutant, the majority of organ fusion events occur
during floral development (Lolle et al., 1998; Krolikowski et al., 2003). Interestingly, the
HTH gene is not epidermis-specific and its involvement in metabolism of additional
compounds, not essential for construction of the cuticle, is not yet clear.
Chen et al. (2003) reported the isolation of the WAX2 gene and interpreted it to be
required for both cutin and cuticular wax deposition. The cuticular membrane of wax2
weighed less, was thicker, disorganized and less opaque. The total wax load on leaves
and stems was decreased to nearly 80%, showing a reduction in the decarbonylase
pathway products and an increase in the acyl reduction pathway products. The WAX2
protein contains certain regions with homology to sterol desaturases and short-chain
dehydrogenases/reductases. It was suggested that it plays a metabolic role in both cutin
and wax synthesis. The cloning and characterization of the same gene (termed YORE-
YORE) was described by Kurata et al. (2003) and the yre mutant showed organ adhesion.
The authors suggested that YRE might encode an enzyme catalyzing the formation of
aldehydes in the wax decarbonylation pathway. Alterations to the fatty acid precursor
pool could also result in plants showing organ fusion phenotypes. The enzyme Acetyl-
CoA Carboxylase (ACCase) catalyses the ATP-dependant formation of malonyl-CoA.
ACCase activity in the cytosol generates a malonyl-CoA pool that is required for a wide
range of reactions including VLCFA elongation which are incorporated into cutin and
waxes. Weak gurke (gk) and pasticcino3 (pas3) mutant alleles that correspond to a defect
in the ACC1 gene, showed abnormal fused leaves that were often vitrified when plants
were grown in-vitro (Faure et al., 1998). A strong organ fusion phenotype was also seen
in transgenic plants raised by Sieber et al. (2000) that expressed a fungal cutinase in
Arabidopsis. Their results suggest that an intact cutin layer is crucial for preventing organ
fusions.
The synthesis of cuticle constituents occurs in the epidermis layer from which
they are transported out to the plant surface. Recently the first clue to the export
mechanism of cuticular lipids through the plasma membrane was provided by the
characterization of the cer5 Arabidopsis mutant (Pighin et al., 2004). The CER5 gene
encodes an ATP-Binding Cassette (ABC) transporter localized in the plasma membrane.
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Apart from the typical reduction in stem cuticular wax load (cer phenotype), the cer5
knockout mutant accumulated sheet-like inclusions in the wax secreting cells. Epidermal
peel staining and observation with light microscopy suggested that they are lipidic in
nature. In addition, the overall levels of fatty acid were not altered and this provided
evidence that only wax transport was affected and not VLCFAs biosynthesis. The CER5
gene expression was detected in all plant organs examined including stems, leaves,
siliques, flowers and roots. This was unexpected since the cer5 phenotype is confined to
leaves and stems. It was therefore suggested that additional transporters must be involved
in delivering wax components. In Arabidopsis, there are over 120 putative ABC
transporters, 29 of them including CER5, belong to the White Brown Complex (WBC)
subfamily (Sanchez-Fernandez et al., 2001). In human and Drosophila, members of this
family secrete cholesterol and plant sterols and play a role in steroid hormone pathway,
respectively (Berge et al., 2000; Hock et al., 2000). In Arabidopsis, ABC transporters are
implicated in the transport of wide range substrates including auxin, sucrose, mono- and
divalent ions. They play a fundamental role in heavy metals transport, resistance to
xenobiotics and different aspects of plant development including regulation of stomatal
opening and closure (Schulz and Kolukisaoglu, 2006).
While the activity of CER5 could explain the transport of wax monomers out of
the epidermal cells, the mechanism responsible for cutin monomers transport remained
unknown. In this paper we describe the DESPERADO (DSO) (outlaw in Spanish) gene
putatively encoding a WBC subtype ABC transporter. We provide several lines of
evidence showing that DSO is vital for the export of both cutin and wax monomers to the
surface of Arabidopsis plants. The various DSO phenotypes and the fact that its
expression was not confined to vegetative organs but was also detected in the root
suggest that it might also be involved in the transport of other type of lipids. Such a lipid
molecule could be for example suberin that shows chemical analogy to cutin and is
typically deposited in plant roots. The results obtained through this study also
demonstrate that DSO is not only vital for proper plant development but also to plants
response to various stresses such as salinity and mechanical wounding.
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RESULTS
Phenotypes of the DESPERADO Loss-of-Function Lines
In human and Drosophila, members of the ABC transporters family play a role in the
transport of lipid substrates (Pohl et al., 2005). To unravel the possible role of ABC
transporters in plant lipid transport, we systematically generated, in Arabidopsis, RNAi
lines for over twenty ABC transporters genes. All members investigated belonged to the
WBC subfamily (Sanchez-Fernandez et al., 2001). One RNAi line (desperado-1; dso-1),
targeted to silence the At1g17840 gene (Fig. 1A), showed an array of morphological
phenotypes including inter-organ postgenital fusions (Fig. 1B, C, D) that resembled those
of mutant plants altered in their cuticle (Yephremov et al., 1999; Chen et al., 2003;
Krolikowski et al., 2003; Kurata et al., 2003; Schnurr et al., 2004; Kurdyukov et al.,
2006a). The DSO protein (AtWBC11; Sanchez-Fernandez et al., 2001) is a memebr of a
small group of WBC transporters that includes the previously described CER5 wax
transporter (AtWBC12; Pighin et al., 2004; Suppl. Fig. S2), AtWBC15/22, AtWBC13
and AtWBC3. DSO shows the highest identity to CER5 (52% at the amino acid level)
and slightly less homology to AtWBC15/22 (51% identity) and AtWBC13 (48%
identity). The most close homolog of the CER5 wax transporter is AtWBC15/22 (85%
identity; Suppl. Table SIII).
The dso-1 plants phenotype was extremely severe as most of the plants were
strongly retarded in growth and upon maturation produced multiple, thin and short
inflorescence stems (a bushy phenotype) probably due to the loss of apical dominance.
The fusion of organs in dso-1 mutants often resulted in rosette leaves that were
misshapen and torn. A toluidine dye uptake test (Tanaka et al., 2004) suggested
malformation of the dso-1 mutants cuticle as they displayed strong coloration after two
min of immersion in the dye whereas no staining was observed in wild-type (WT) plants
(Fig. 1G and H). The leaf surfaces of several cuticular mutants were previously shown to
support WT pollen germination (Lolle and Cheung, 1993; Lolle et al., 1998; Sieber et al.,
2000; Wellesen et al., 2001; Kurdyukov et al., 2006b). Scanning electron microscopy
(SEM) revealed that fully expanded rosette leaves of dso-1 plants do not support WT
pollen germination (data not shown). In Arabidopsis rosette leaves, the vascular tissue is
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composed of the main vein in the middle of the leaf blade that is interconnected by
secondary and higher order veins, forming a complex network. The vascular patterns in
the dso-1 mutant appeared to be altered compared to the WT leaf patterns (Fig. 1J and K).
In the dso-1 leaf blade less tertiary and quaternary veins were observed. Moreover, the
veins along the edge of the leaf formed a discontinuous circle.
We also generated misexpression lines by expressing DSO under the control of
the 35S CaMV promoter (lines termed dso-2) and identified a T-DNA insertional line
(SALK_072079; Fig. 1A) in the DSO gene (lines termed dso-3). Semi-quantitative RT-
PCR was performed for all three loss-of-function mutant genotypes in order to determine
the levels of the DSO transcript. In the dso-1 and dso-2 lines a significant reduction in
transcript levels was evident whereas in the dso-3 line no transcript was detected
indicating that dso-3 is a null mutant (Suppl. Fig. S1). The results with misexpression
suggested that instead of overexpression we obtained cosuppression (detected in one third
of the primary transformants).
In Suppl. Table SII we depicted the different phenotypes and their degree of
penetration among the three DSO loss-of-function genotypes. Overall, the dso-2 and dso-
3 plants showed similar phenotypes to the ones observed in the dso-1 RNAi line (Fig. 1)
but with different levels of penetration. The cosuppression dso-2 lines had a relatively
mild phenotype compared to dso-1 and dso-3 as they developed an almost regular
inflorescence stems that frequently had a glossy cer-type phenotype (Jenks et al., 1995;
Fig. 1F). In some cases, cer-type phenotypes were observed only in certain stem parts but
not others in the same plant. The dso-2 plants showed notched rosette leaves. The
majority of dso-1 and dso-3 seedlings grown in tissue culture developed unusual, callus
or stigmatic-like, protrusions from epidermal cells (Fig. 1E). A similar phenotype was
observed previously in transgenic Arabidopsis plants expressing a fungal cutinase (Sieber
et al., 2000). Leaves (Fig. 1M) and flowers morphology was affected in all three dso
genotypes as petals were folded and twisted (Fig. 1I) and they produced short almost
seedless siliques (Fig. 1L). When seeds of dso-2 lines were immersed in toluidine blue
dye solution they showed increased staining compared to WT seeds (data not shown)
suggesting impaired integrity of the seed surface. The stem cer-type phenotype detected
by visual inspection was in agreement with the dramatic decrease in load of epicuticular
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wax crystals on dso-2 mutant stem surfaces observed by scanning electron microscopy
(SEM) (Fig. 2A and B).
SEM examination of leaf surfaces uncovered notable phenotypes in the dso lines.
Apart from random ruptures in the epidermis (Fig. 2C), they developed dehydrated
trichomes with shortened stalks and irregular branching patterns and they were often
collapsed (Fig. 2D, E, F). Misshapen, asymmetric stomata and abnormal leaf pavement
cells pattern were regularly observed (Fig. 2I, J).
Alterations in reproductive organ morphology were also detected in dso plants
examined by SEM. Flowers had curved petals and distorted anther filaments (Fig. 2G and
H). Moreover, the typical petal abaxial epidermis conical cells were variable in size and
misshapen in the petal folding area (Fig. 2K and L). Pollen grains were often absent from
the stigmatic papillary cells and they were often shriveled (Fig. 2M-P). Alexander stain
for a pollen viability test showed that 23% of dso-3 pollen is unviable and shriveled (data
not shown). We also performed reciprocal crosses between dso-3 plants and WT plants.
When dso-3 plants were used as male, short semi-sterile siliques were obtained. On the
other hand, when dso-3 flowers were pollinated with WT pollen no fertilization occurred.
From these observations we can conclude that the dso-3 sterility originates from both
defective male and female reproductive organs. The descendants of the backcrossed dso-
3 homozygous plants displayed the same phenotype excluding the possibility that the
phenotype originated from the background mutations. The effect of decreasing DSO
transcript levels was also evidenced in below-ground tissues as dso plants displayed
reduced amounts of lateral roots compared to WT plants (Fig. 3; Suppl. Fig. S3).
DSO and CER5 Might Function in the Same Pathway
In order to evaluate the interaction between CER5 and DSO we crossed the mild
phenotype dso-2 plants (with glossy stems, curved petals and without organ fusion
phenotype) with the cer5-1 mutant. As mentioned above, the cer5-1 mutant does not
display any additional visual phenotype apart from its glossy cer-type stem. The dso-
2/cer5 double mutant showed severe fusion already at early developmental stages (Fig.
1N), suggesting an additive phenotype and providing evidence that these two genes could
act in the same pathway.
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Analysis of DSO and CER5 gene expression profiles using GENEVESTIGATOR
(https://www.genevestigator.ethz.ch/at/) revealed similar expression patterns for the two
genes. Their expression is highest in seedlings, young leaves and in the inflorescence. In
the root organs, both genes display highest expression in lateral roots. Moreover, using
the PRIME co-expression search tool we found that expression of both genes is highly
correlated (http://prime.psc.riken.jp/?action=coexpression_index).
DSO Loss-of-Function Lines Lack a Cuticular Layer in Organ Fusion Areas and
Contain Unusual Lipidic Cytoplasmatic Inclusions in Epidermal Cells
We used Transmission Electron Microscopy (TEM) to examine the changes in cuticle
formation when two dso-1 rosette leaves are fused. These observations revealed that
when complete fusion in between the two leaves surface has occurred, absolute
disappearance of the cuticular layer was detected suggesting copolymerization of
adjacent cell walls (Fig. 4A, B, C).
Unlike other cuticular mutants (Chen et al., 2003; Kurata et al., 2003; Xiao et al.,
2004; Kurdyukov et al., 2006a; Kurdyukov et al., 2006b), dso mutant and transgenic lines
stems and leaves were not altered in the cuticle ultrastructure. On the other hand, detailed
TEM inspection of both leaves and stems cells of dso-2 and dso-3 indicated that the
transport of cuticular components might be compromised in the dso mutants. Unusual
trilamellar cytoplasmatic inclusions were observed in epidermal cells of both leaves and
stems of the mutants (Fig. 4D-I). These structures could not be detected in epidermis cells
of WT leaves and stems and not in other cell types of the mutants. In order to verify the
nature of the cytoplasmatic inclusions epidermal peels of dso-3 stem were stained with
Nile Red and observed with fluorescence microscopy (Figs. 4J and K). The results
indicate that as in the case of the cer5 mutant these inclusions are lipidic in nature (Pighin
et al., 2004).
Expression of Reporter Genes Driven by the DSO 5' Region
To study the tissue specificity of DSO we examined the expression of GUS and GFP
reporter genes under the control of the DSO 5'-upstream sequence. For GUS and GFP
expression, 2294bp and 4417bp of the DSO 5'-upstream region were transcriptionaly
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fused to either one of these reporters, respectively, and reporter activity was evaluated in
different tissues of T2 plants. Reporter GUS and GFP expression indicated that DSO is
expressed in the seed coat and the endosperm (data not shown) and later during embryo
development in the radical tip, and vasculature (Fig. 5A). DSO expression was detected
in seedlings in the cotyledons, root tip and young leaves (Fig. 5B, C, F, and G). In both
young and mature leaves, expression was detected in trichomes and stomatal cells (Fig.
5J and K) and weaker in the rest of the blade. The strongest expression was detected in
the main vein and the expanding basal portion of the leaf (Fig. 5G). In roots of mature
plants, DSO expression was clearly observed in lateral root primordia and developing
lateral roots, but was also detected throughout the vasculature (Fig. 5D, E, F). In the
inflorescence, expression could be detected in all floral organs predominantly in the
anthers, styles and in young siliques (Fig. 5H). In the developing siliques the strongest
expression was detected in young siliques (in the base and tip; Fig. 5I). Cross section of
the inflorescence stem DSO expression showed that DSO expression was not confined to
epidermis as it was detected in epidermal and mesophyll cells (Fig. 5L). Overall,
developing rather than mature organs appear to express DSO.
The DSO Protein Subcellular Localization
We also examined the subcellular localization of DSO by generating transgenic plants
harboring a construct in which GFP was fused in frame to the N-termini of the full-length
DSO gene and expression was driven by the DSO 5'-upstream region. Two thirds of the
transformants displayed a cosupression phenotype (data not shown). Five T2 GFP
positive plants, with no signs of cosupression were used for whole mount confocal
microscopy of leaves (Figs. 6E-H), protoplasts preparation from stem epidermis enriched
tissues (Figs. 6A-D) and preparation of free hand stem cross sections (Figs. 6I-L). In
protoplasts, the GFP signal was detected in the periphery of the cells suggesting plasma
membrane localization. The use of protoplasts allowed us to exclude cell wall specific
expression. To determine whether the observed fluorescence was associated with the
plasma membrane we used a plasma membrane specific marker (FM4-64). The results
showed that DSO is co-localized with FM4-64 (Figs. 6E-H). Analysis of the stem cross
sections of the same plants indicated that in contrary to the promoter directed expression
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GFP was detected exclusively in the epidermis. Moreover, it was localized in polar
manner in the epidermis side facing the extracellular matrix (Figs. 6I-L).
Changes in Cutin and Wax Monomers Composition in the dso Mutants
To gain more knowledge on the precise role of DSO in the transport of cuticle associated
lipids we performed Gas Chromatography-Mass Spectrometry (GC-MS) analysis of
epicuticular waxes on the surface of dso-3 and WT plants. Chemical analysis revealed a
3-fold decrease in total stem wax load in dso-3 compared to WT stems (6.77 ± 1.02
µg/cm² versus 19.66 ± 2.22 µg/cm²; Table I). The C29 monomers, particularly alkanes
(11-fold decrease), ketone (2.6-fold decrease) and secondary alcohol (1.8-fold decrease)
were largely responsible for this decrease (Fig. 7A).
The data gathered to this point suggested that DSO might not only be required for
wax but also to cutin monomers transport. Consequently, we conducted GC-MS analysis
of the previously reported Arabidopsis cutin constituents including regular fatty acids, 2-
hydroxy fatty acids, ω-hydroxy acids and α,ω-dicarboxylic acids (Bonaventure et al.,
2004; Xiao et al., 2004; Franke et al., 2005). The results showed that total cutin
monomers load per leaf area in dso-3 was reduced 3.3-fold compared to WT (63.96 ±
4.38 ng/cm² versus 211.58 ± 11.63 ng/cm²; Table II and Suppl. Table SI). Moreover,
levels of all the twenty two detected cutin monomers were dramatically decreased in dso-
3 plants (Fig. 7B; Suppl. Table SI). Interestingly, chemical analysis of dso-2 leaf
cuticular lipids showed reduction only in wax load while no significant difference in
cutin monomers load was noted. However, cutin composition in dso-2 was different from
WT. Levels of 2-hydroxy and ω-hydroxy fatty acid levels were upregulated, whereas the
levels of α,ω-dicarboxylic acids (Bonaventure et al., 2004; Franke et al., 2005) were
significantly reduced (Suppl. Fig. S4). Thus, DSO loss-of-function results in altered level
and composition of both cutin and wax monomers in the cuticle.
DSO is Induced under Salt, ABA and Wound Stresses
Salinity is a polymorphous stress which impede plant development and viability through
two shared effects: osmotic and nutritional. High salinity affects plants through ion
toxicity as well. In order to assess how DSO is implicated in these environmental stresses
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we subjected dso-1 seedlings to salinity stress (200 mM NaCl). We found that dso-1
seedlings are more susceptible to salinity stress than WT ones (Fig. 8C and D). Using
semi-quantitative RT-PCR we evaluated DSO transcript levels in WT plants under the
same conditions and found up-regulation in DSO expression upon salt stress (24h
exposure) (Fig. 8A).
ABA is a universal plant hormone widely implicated in adaptation to stress. It
regulates stomatal closure and increasing evidence suggests that it is involved in root
branching (De Smet et al., 2006). Semi-quantitative RT-PCR showed up-regulation of
DSO transcripts in RNA derived from seedlings treated for 24h with 50µM ABA (Fig.
8A). DSO expression was not only induced by salinity and increased ABA levels but also
upon mechanical wounding as detected in leaves expressing GUS driven by the DSO 5'-
upstream region (Fig. 8B). We also examined the expression of CER5 (AtWBC12) and
AtWBC13 genes under the same salt and ABA treatments (Fig. 8A). The results show that
while CER5 expression is induced in salt and ABA treatments (similar to DSO),
expression of WBC13 is induced by salt but not by ABA.
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DISCUSSION
Cutin is the third most abundant biopolymer on earth after lignin and cellulose. It is a
major component of the cuticle that covers all plant surfaces exposed to air. Despite its
significance only little is known about the transport of cutin monomers from their
synthesis site in the epidermal cells to the extracellular domain where the cuticle is
assembled. This study demonstrates that in Arabidopsis, DSO, a plasma membrane
localized ABC transporter is required for proper export of cutin monomers through the
plasma membrane to the cuticle. It is also required for the transport of wax monomers, a
different set of cuticular components, as reported earlier for the Arabidopsis CER5
transporter (Pighin et al., 2004). The transport of these two compound classes (i.e. cutin
and wax) by the same transporter is likely since a large number of the ABC transporters
characterized to date were able to handle several structurally different compounds
(Yazaki, 2006). Moreover, the information regarding DSO gene expression and the array
of loss-of-function phenotypes suggest that the DSO protein might also be associated
with transport of other, wax or cutin-like molecules.
The DSO protein sequence shares 52% identity with CER5 but interestingly even
much higher level of similarity with the cotton (Gossypum hirsutum) GhWBC1 protein
(84% identity). GhWBC11 is highly expressed in developing cotton fiber cells and its
overexpression in Arabidopsis resulted in plants with short siliques containing severely
shriveled embryos and with only several seeds per silique (Zhu et al., 2003). A variable
amount of suberin could be found at the cotton fiber base which is typically deposited in
concentric layers, alternating with polysaccharides (Ryser, 1992). The relatively high
similarity in sequence between DSO and GhWBC1, the overexpression phenotype and
the presence of suberin and a thin cuticle, with wax and cutin components, in cotton
fibers (Schmutz et al., 1996) suggests a similar role to the two transporter proteins in
Arabidopsis and cotton.
DSO expression was not confined to the epidermis of aerial parts as anticipated
for a transporter of cuticular components. Its expression was also detected in other aerial
plant organs and cell types as for example in the stem mesophyll cells. Interestingly,
relatively strong DSO expression was detected in lateral root primordia, the developing
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lateral root and in the root vasculature. Moreover, we also detected reduced root
branching in the dso lines. In a different cuticular mutant, bodyguard (bdg), an increase
in root branching was observed (Kurdyukov et al., 2006a). This is not the first study in
which root expression of genes associated with cutin and wax metabolism is reported.
Root expression was also detected for KCS1 (Todd et al., 1999), YRE (Kurata et al.,
2003), BODYGUARD (BDG; Kurdyukov et al., 2006a), SHINE3 (SHN3; Aharoni et al.,
2004), HTH (Kurdyukov et al., 2006b) and the CER5 (Pighin et al., 2004) genes. Fatty
acid analysis of the kcs1-1 mutant roots revealed a two-fold increase in α,ω-dicarboxylic
acids and this result led the authors to suggest that KCS1 is not only implicated in wax
metabolism but also in the suberin biosynthesis pathway (Todd et al., 1999). Apart from
expression of DSO in roots, two more lines of evidence suggest that DSO is involved in
transport of other lipid derived chemicals (such as suberin) that are similar in structure to
cuticular lipids. The first supporting evidence is the altered leaves vascular patterns in the
dso-3 mutant which might be a result of changes in suberin deposition during secondary
growth in the vascular tissue.
A second point supporting this argument is the striking similarity between the
aliphatic monomer composition of Arabidopsis cutin and suberin (Bonaventure et al.,
2004; Franke et al., 2005). Indeed, a few recent studies suggested that the long-chain α-
,ω-dicarboxylic fatty acids are not only constituents of the cutin polyester in Arabidopsis
(Bonaventure et al., 2004; Kurdyukov et al., 2006b). They might play additional roles as
a "suberin-like" network in the secondary cell wall or required for the cross-linking that
ensures the integrity of the primary epidermis cell-wall. Evolutionarily, it is feasible that
during the course of adaptation to terrestrial environments, plants modified the substrate
specificity of a lipid transporter to a protein which could fulfill the requirements for
constructing interface layers using three different building blocks, namely, cutin, wax and
suberin. We therefore anticipate that in the near future, as suggested here for DSO and
previously for KCS1 (Todd et al., 1999), more genes associated with wax and cutin
metabolism will also be recognized as playing a role in the biosynthesis and transport of
other plant interface components (e.g. suberin).
Full-size ABC transporters contain two ABC and two transmembrane domains
(TMD) in a single polypeptide chain (Schulz and Kolukisaoglu, 2006). On the other
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18
hand, half-size transporters such as the one encoded by DSO, achieve their functionality
by combining two ABC-TMD units as homo- or heterodimers in a membrane bound
transporter complex. One possible dimerization candidate for wax transport is CER5
although two other proteins in the same phylogenetic clade (At3g21090, WBC15 and
At1g51460; WBC13; see Sanchez-Fernandez et al., 2001) might also act as partners for
the transport of cutin and other molecules. For example, in Drosophila, dimerization of
three half-transporter ABC proteins related in sequence, White with Brown and White
with Scarlet, is required for the transport of different eye pigment precursors into pigment
cells (Mackenzie et al., 2000). Further experiments should identify the interacting
partners in between the transporters, their substrate specificity and their mode of action.
With respect to their mode of action, ABC transporters might actively expel the
substrates into the extramembrane space or through a side port of the transporter into the
upper leaflet of the plasma membrane bilayer in an ATP-dependant process.
Alternatively, they might act by turning over the substrates from the inner to the outer
leaflet of the plasma membrane acting as an "hydrophobic vacuum cleaner" (flippase
activity; Chang and Roth, 2001). Another important question is how do cuticular lipids
reach the ABC transporter localized in the plasma membrane? The possible routes
include: a) they are picked up by fatty acid binding proteins and relocated to the
transporter, b) relocation through a vesicular pathway either by the formation of
oleosome bodies coated by oleosin-like proteins or the formation of uncoated vesicles
that contain the cuticular lipids in lipid rafts (Schulz and Frommer, 2004).
Wax load, in particularly the C29 alkanes, was dramatically reduced in stems of
both the dso and the cer5 mutants. It should be noted that in the case of dso, levels of
several stem wax components were significantly increased in the mutant compared to the
wild-type (i.e. C27 alkanes, C31 secondary alcohols and C24 primary alcohols), suggesting
compensation for the loss of other cuticle components. Leaves and fruit of a transposon
insertion mutant in the tomato LeCER6 encoding a VLCFA elongase (β-keto acyl-CoA
synthase; KCS) were deficient in n-alkanes and aldehydes with chain lengths beyond C30.
In the same plants, much higher levels of pentacyclic triterpenoids (α-,β-, and δ-amyrin)
were detected, suggesting compensation for the reduction in aliphatics (Vogg et al.,
2004). In contrast to the cer5 knockout mutant showing the typical glossy/cer like, stem
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19
phenotype with no effect on plant architecture, the dso mutants exhibited a range of
dramatic phenotypes in nearly every plant organ examined. The lesion in DSO had a
dramatic effect on epidermal cells development, including alterations to trichomes,
stomata and pavement cells. In the case of trichomes, they were collapsed and
underdeveloped. Similar phenotypes were observed in trichomes of several mutants and
transgenic plants involved in wax and cutin metabolism including SHINE1
overexpression lines (Aharoni et al., 2004), fdh (Yephremov et al., 1999), lcr (Wellesen
et al., 2001), cer10 (Zheng et al., 2005) and bdg (Kurdyukov et al., 2006a). It is intriguing
that SHN3, one of three AP2 domain transcription factors suggested to act as activators of
the wax biosynthetic pathway in Arabidopsis, showed strong and specific expression in
the trichome support cells that surround the base of the trichome (Aharoni et al., 2004).
The collapse of trichomes phenotype detected in dso and bdg might be a result of altered
development of the support cells and suggests that these cells might contain a unique
component such as suberin that provides them with the strength to hold the trichomes.
This hypothesis is further supported by an earlier report on the presence of suberized cell
walls in the boundary between plants and secretary organs such as trichomes
(Kolattukudy, 2001).
Basal or support cells of trichomes, cuticular ledges and cuticles over anticlinal
cell walls together provide aqueous pores for plants cuticles (Schonherr, 2006). The exact
biological impact of the collapsed trichomes supporting cells of the dso mutants with
regard to the aqueous solutes movement requires further investigation. Since aqueous
pores serve as a main gate for foliar penetration of exogenously applied compounds
including agricultural chemicals, deciphering the exact role of cutin metabolism genes on
cuticular aqueous pores assembly in plants will have paramount significance for
agriculture and ecophysilogy.
A major phenotype of the dso loss-of-function genotypes was the occurrence of
postgenital fusions which involve surface contact between organs that have already
developed as individual entities (Verbeke, 1992). In dso, fusions could be noticed in
between different types of vegetative and reproductive organs including between distal
parts (even tips) of rosette leaves. Although multiple mutants have been described that
posses postgenital organ fusion (see Introduction), one cannot identify the factor/s
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20
mediating this phenomenon. The assortment of mutants exhibiting postgenital fusions
described up to date differ in most of the parameters used to phenotype cuticular mutants
including in: permeability to a cationic dye, rate of chlorophyll leaching from leaves in
alcoholic solution, rate of water loss, defects in pollen hydration, male sterility recovery
under high humidity, glossy appearance, alterations to the cuticle ultrastructure, stomatal
index, trichome number and branching, and changes in the cuticle chemical composition.
This difference in phenotypes in the class of postgenital organ fusion mutants might be
simply due to the nature of mutations, gene redundancy and the degree of phenotype
penetration. However, it may also be that a single factor, not yet identified either a
signaling lipid-based molecule or a specific structural change in the cuticle or the
epidermal cell wall (Nawrath, 2006), could trigger this phenomenon. A more detailed
comparison between the dso and cer5 mutant phenotypes might aid in identifying the
factor promoting postgenital organ fusions since cer5 shows similar alterations in waxes
compared to dso but does not exhibit fusion phenotypes.
In recent years an increasing number of cuticle related phenotypes and processes
have been described (Nawrath, 2006). It is apparent that cuticle associated proteins do not
only play a role in plant development but they are also very active in the plant response to
different stress conditions. Our study suggests that DSO plays a vital role in stress
response programs mediated by the cuticle, including salt stress, and wounding as its
expression was induced under these conditions. Supportive evidence for the importance
of DSO function in response to stress, particularly salt stress was provided by
experiments showing that dso-1 seedlings are highly sensitive to salt treatment.
Preliminary characterization of a gene trap line corresponding to DSO also showed that it
is also induced by multiple stresses including ABA, high salt, and glucose (Alvarado et
al., 2004). Like DSO, the expression of CER6 encoding a VLCFA condensing enzyme
was enhanced by osmotic stress and the presence of ABA (Hooker et al., 2002).
The role of DSO in stress response could be explained by the need to alter surface
structure upon water, salinity and mechanical stress (Shepherd and Wynne Griffiths,
2006). Leaf transpiration has stomatal and cuticular components. Transpiration through
the cuticle is largely determined by surface characteristics such as wax thickness and wax
microstructure. Wax deposition that occurs rapidly within a few days is often a response
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21
to water stress and stress-resistant plants often have thicker waxes compared to
susceptible ones. Increased wax deposition upon exposure to salinity stress was reported
for several plants as for example salt sensitive jojoba and seems to be primarily a
response to water deficit (Mills et al., 2001). Moreover, in leaves of salt sensitive jojoba,
wax deposition is induced by exogenous ABA (Mills et al., 2001). Finally, mechanical
stress such as wounding due to strong wind, rain drops, and leaf to leaf contact could also
induce the formation of wax for reforming leaf structure (Shepherd and Wynne Griffiths,
2006). More in relation to ABA and stress response, dso-1 mutant plants displayed
reduced roots branching number. Lateral root formation is essential for the adaptation of
plants to changing environmental challenges such as increased osmotic stress and
salinity. Growing evidence suggest that ABA is involved in the regulation of root
branching (De Smet et al., 2006). Interestingly, GUS expression driven by the DSO 5'-
upstream region indicated DSO expression throughout lateral root development.
This study adds another piece to the puzzle of how plants assemble their
outermost surface. Nevertheless, the mechanism of cuticle monomers transport from their
site of synthesis to the membrane and further to the extracellular domain remains unclear.
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22
MATERIALS AND METODS
Plant Material and Growth Conditions
All plants, including the transgenic lines were grown in the climate room at 20°C, 70%
relative humidity and a 16/8 hr light/dark cycle and were in the Arabidopsis thaliana
ecotype Col-0. Salk T-DNA insertion line, SALK_072079, was obtained from the
European Arabidopsis Stock Centre (Alonso et al., 2003). T-DNA insertion was
identified in the fifth exon using oligonucleotides designed by the iSECT tool (Signal T-
DNA Express website). DSO RNAi F1 plants seeds were stratified for 2-3d at 4°C and
subsequently sown on MS plates supplemented with kanamycin 50µg/ml and grown in a
culture room under continuous light conditions at 20°C. Two weeks old seedlings were
subsequently transferred to soil. For the roots branching experiment, 1 week old
transgenic plants displaying the typical fusion phenotypes were transferred to vertically
placed MS plates for additional 2 weeks growth. Only lateral roots branching out from
the main root were counted. For the salt stress assay, two weeks old seedlings were
transferred to MS plates supplemented with 200mM NaCl for additional 1 week growth.
Plants survival was monitored during 1 week after application of the salt stress.
Generation of Plant Transformation Constructs and Transgenic Arabidopsis
For generating the DSO RNAi construct, a 298bp genomic fragment was amplified with
the following oligonucleotides: sense (5'-AAAAAGCAGGCTCATATGTGACCCAAG-
ACGATAAC-3'), anti- sense (5'-AGAAAGCTGGGTGCAGAAGCACTATCAAGACC-
AC-3') and integrated into pDONR201 using the Gateway cloning system
(INVITROGEN). The LR Clonase (INVITROGEN) was then used to recombine this
fragment into pK7GWIWG2(I) binary vector (Karimi et al., 2002). For overexpression,
the full length DSO cDNA was amplified and inserted into BJ36 (Moore et al., 1998) the
under control of the 35S CaMV promoter and subsequently cloned into the pMLBART
binary vector. For plants transformation inflorescences were dipped into Agrobacterium
tumefaciens strain GV3101 carrying the transgene construct as described (Clough et al.,
1998).
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23
Toluidine Blue and Nile Red Staining and Visualization of the Leaf Vasculature
The method for examination of cuticular integrity was performed as described (Tanaka et
al., 2004). For the leaf vasculature visualization, leaves of 4 weeks old plants were
bleached for 24h in ethanol: acetic acid (6:1), immersed in 70% ethanol for 1h, mounted
in chloralhydrate mixture (chloralhydrate/ glycerol/ water- 8: 1: 2) and observed using
standard light microscopy. Nile Red staining was performed as described previously
(Pighin et al., 2004).
Gene Expression Analyses Using Semi-Quantitative RT-PCR
Two to four rosette leaves of 3 to 4 weeks old plants were used for total RNA isolation.
Total RNA was isolated using the RNAeasy Plant Mini Kit (QIAGEN) according to the
manufacturer’s protocol. Isolated total RNA was treated by DNAse according to the
manufacturer’s protocol (PROMEGA). Total RNA (500ng-1µg) was transcribed to
cDNA using oligo(dT)15 and AMV reverse transcriptase (CHIMERIX, INC.). For the
PCR reaction, 1-5µl of was used as a template for a 20µl PCR reaction with the following
primers: DSO sense (5'-ATGTTACTCCTTGGGTCAGAG-3'), antisense (5’-
ATTTCGGCACAATGCAAAC-3’) with expected band size of 399bp; CER5 sense (5’-
TGGGATGGAAGTGAGAAAGG-3'), antisense (5'-GAGCCAAGATCGATGTGTAG-
3') with expected band size 193bp; WBC13 sense (5'-GGGGATTGTCACAGAAAGGA-
3') antisense (5'-TGACCCGACACAAATGGATA-3') with expected band size 156bp.
PCR was started with a 94ºC denaturation step for 2 minutes, followed by 45 seconds at
94ºC, 45 seconds at 58ºC, 1 minute at 72ºC and 5 minutes of final elongation. The
following amounts of amplification cycles were used: 27 for DSO, 33 for WBC13 and 36
for CER5.
DSO Promoter Analysis Using GUS or GFP as Reporter Genes
The DSO 5'-upstream region (2394bp, termed pDSO) was amplified using the following
oligonucleotides: antisense (5'- NcoI-AAACCATGGCTCTTAAACCAAAACAGAGG-
ATT-3'), sense (5'-BamHI-TTTAAGAATTAATTGTCTAAATAAC-3'), excised with
appropriate restriction enzymes and subcloned into the pMAX vector containing the GUS
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24
coding sequence and the NOS terminator. The pDSO fragment together with the GUS
gene was excised with Pac1 and Asc1 and cloned into the binary vector pBIN+ (van-
Engelen et al., 1995). GUS staining and embedding was performed according to Pekker
et al. 2005. For promoter directed GFP expression experiment, a two-component LHG4-
10OP transactivation system was used as described (Moore et al., 1998). The DSO 5'-
upstream region (4417bp) was amplified using the oligonucleotides: sense (5’-BAMH1-
AGGATCCCTCTTAAACCAAAACAGAGG-‘3), antisense (5'-PST1-CTGCAGGGTA-
AGTAATTTAGCAATTG-‘3) and placed 5' to the LHG4 gene in BJ36. The BJ36 was
cut with Not1 and subcloned into pART27. In order to trans-activate the GFP, this
construct was transformed into a 10OP:GFP line. Seeds of the F2 GFP-positive plants
were dissected for observation of embryos and GFP signal was observed either using
standard fluorescent microscopy or confocal microscopy (excitation at 488nm, emission
was at 500-530nm for GFP and 620-750nm for chlorophyll).
Scanning Electron Microscopy
Stems (second internodes from the bottom) were collected from wild-type and dso-1,
dso-2 or dso-3 plants after 5-6 weeks of growth. Leaves were collected and fixated with
glutaraldehyde using standard protocols and dried using critical point drying (CPD).
Samples were mounted on aluminum stubs and sputter-coated with gold. Scanning
electron microscopy was performed using an XL30 ESEM FEG microscope (FEI) at 5-
10kV.
Transmission Electron Microscopy
Leaves and stems from 50 days old plants were collected and processed using a standard
protocol (Weigel and Glazebrook, 2001). The spurr resin embedded samples were
sectioned (70nm) using an ultramicrotome (LEICA INC., IL) and observed with a Tecnai
T12 transmission electron microscope (FEI).
Subcellular Localization of DSO and Confocal Microscopy
For examination of the DSO protein subcellular localization, the fragment containing the
DSO 5'-upstream region (4417bp) and the DSO cDNA were translationaly fused to GFP
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25
at N’-termini. The DSO cDNA was amplified using the following oligonucleotides: sense
(5’-BamH1- TTTGGATCCCTACCATCTGCGAGCTCCATC), antisense (5’-Xho1-
AAACTCGAGAATGGAGATAGAAGCAAGCAG-‘3), excised with BamHI and Xho1
and cloned into the 10OP::N’-GFP in the BJ36 vector (Moore et al., 1998). The DSO 5'-
upstream region was amplified using the following oligonucleotides: sense (5’-Kpn1–
AAAGGTACCTAAGAATTAATTGTCTAAATAAC-‘3), antisense (5’-Xho1-
TTTCTCGAGCTCTTAAACCAAAACAGAGGATT-‘3), cut with Kpn1 and Xho1. The
10OP::N’-GFP:DSO in BJ36 was cut with Sal1, Not1 and a modified pBlueScript vector
(STRATAGENE CLONING SYSTEMS, La Jolla, CA) cut with Not1 and Kpn1. After a
triple ligation the obtained vector was cut with Pac1, Asc1 and cloned into the pBIN+
binary vector (van Engelen et al., 1995). Protoplasts from epidermis enriched stem
segments were prepared as described elsewhere (Sheen et al., 2001). Fluorescence was
observed by an Olympus CLSM500 microscope with an argon laser at 488 nm for
excitation and images for GFP and chlorophyll signals were collected through 505-525
nm for GFP and 620-750 nm for chlorophyll and FM4-64. FM4-64 plasma membrane
staining was performed as described elsewhere (Zheng et al., 2005). When the stem cross
sections were analyzed using confocal microscopy, the presence of the GFP signal was
verified using the laser photobleach approach. Time-lapse images were taken before the
bleach and immediately after photobleaching, to assess the recovery of the GFP signal
(Suppl. Movie M1).
Wax and Cutin Analysis
For wax analysis, the second internodes of 7-week-old plants (n=5) or leaves of 4-5
weeks old plants were cut and immersed twice in 5ml of hexane for 30s at room
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26
temperature. The obtained solution containing the cuticular waxes was spiked with 2µg
of tetracosane (FLUKA) as an internal standard and analyzed as described (Kurdyukov et
al., 2006a). The extracted stems area was calculated based on the measurements of stem
length, upper and lower diameter. Leaves area was assessed using the NIH ImageJ
software. For cutin analysis, 4 week-old mature leaves (n=3-4 for WT and 25 for dso-2 or
dso-3) were photographed and their areas measured using the NIH ImageJ software.
Soluble lipids were extracted from samples by dipping in 10ml of methanol/chloroform
(1:1, v/v) mixture for 14 days (solvent was changed daily). Leaf material was dried,
weighed (about 25-30mg) and used for analysis as described (Kurdyukov et al., 2006a).
Generation of cer5-1/dso-2 Double Mutants
Plants exhibiting a mild dso-2 phenotype were crossed with cer5-1 plants. Seeds from F2
plants with a dso-2 phenotype were selected and the double mutants were identified in the
F3 generation based on the additive phenotype that segregated in close to a 3:16 ratio and
PCR to verify the presence of the 35S CaMV promoter fragment and the DSO cDNA
using the oligonucleotides: sense, 35S-out- CAATCCCACTATCCTTCG, antisense DSO
TGTCTGCTTGCTTCTATCTC (expected band size 361bp).
Statistical Analysis
Data are presented as mean ± sd (standard deviation). Statistical significance was
determined by a student t-test. Probability values (P) smaller than 0.05 were considered
to be statistically significant. One star means that P<0.05 and two stars P<0.01.
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27
Acknowledgements
We thank Dr. Eyal Shimoni and Hanna Levanony for assistance with TEM; Dr. Eugenia
Klein for help with SEM; Alexander Goldschmidt for the 10OP:GFP line and kind help
with fluorescence microscopy; Max Itkin for the pMAX construct; Vladimir Kiss for
assistance with confocal microscopy and Guy Gafni for technical assistance. .
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28
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35
FIGURE LEGENDS
Figure 1. The dso loss-of-function mutants phenotypes. (A) Scheme of the DSO locus
(At1g17840) depicting the RNAi target sequence in the second exon and T-DNA
insertion located in the fifth exon of the dso-3 line. (B) Inter-organ postgenital fusions in
the dso-1 mutant plant. A fusion area between inflorescence and leaf is indicated by
arrow. (C) Fusion between a leaf blade (lb) and a floral bud (fb) in dso-1. (D) Fusion
between two leaves of dso-2 plant. (E) Unusual protrusions in dso-3 plants grown in
tissue culture. (F) A cer phenotype in dso-2 plant stem (left) versus wild-type (WT) stem
(right). (G) dso-1 and (H) WT, one month old plants after 2 min immersion in toluidine
blue staining solution. Sites of the dye penetration are indicated by arrows. (I) A dso-2
flower phenotype. Underdeveloped petal is indicated by arrow. (J) Vasculature
phenotype detected in dso-3 rosette leaf. (K) WT rosette leaf vasculature. (l) Siliques of
dso-3 (1), dso-2 (2) and WT (3). (M) Leaf phenotype of a severe dso-2 mutant line (1),
mild dso-2 mutant line (2), dso-3 (3) and WT (4). (N) The cer5-1/dso-2 (mild) double
mutant phenotype. Fusion between leaves is indicated by arrow. Bars are 2 mm in (B)
and 1 mm in (C).
Figure 2. Scanning electron microscopy pictures. (A) Stem wax load of dso-2 and (B) of
wild-type (WT) plants. (C) Occasional ruptures in the epidermis of dso-2. (D) Distorted
and underdeveloped trichomes of dso-3. (E) Collapsed and underdeveloped trichome of
dso-1. Misshapen support cells are indicated by an arrow. (F) A WT trichome. (G)
Abnormal anther filament of a dso-3 flower. (H) Curved petals of a dso-3 flower. (I)
Light microscopy images showing aberrant pavement cells pattern and abnormal stomatal
cells (indicated by arrows) in dso-3 and (J) abaxial leaf epidermis of WT plants. (K)
Scanning electron micrographs (SEM) of abnormal conical cells in the abaxial epidermis
of a dso-3 flower petal and conical cells in the abaxial epidermis of a WT flower petal
(L). (M) SEM micrographs of shriveled pollen grains in dso-3 lines and pollen grains in
WT (N). (O) Stigmata papillae of dso-3 and those of WT (P), grains could not be
detected in dso-3. In: (C) bar is 20 µm; (I) and (J) 50 µm.
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36
Figure 3. The dso mutants exhibit a root branching phenotype. Root branching number
of three weeks old dso and wild-type (WT) plants. Error bars represent standard
deviation. Two asterisks mean P<0.01
Figure 4. Transmission electron microscopy pictures illustrate: (A) The fusion area
between dso-1 leaves in which the cuticles (cut) of either leaf align next to each other.
(B) and (C), areas of leaf fusions in dso-1, red arrow heads mark areas in which the
cuticle is absent, cuticles are indicated by arrows. Unusual inclusions (arrows) in the
epidermal cell cytoplasm of a dso-3 leaf (D), and (F), (G) in the epidermal cell of a dso-2
stem, (H) in an epidermal cell of a dso-3 stem. (E) Epidermal cell cytoplasm of WT leaf.
(I) Epidermal cell cytoplasm of WT stem. Fluorescence images (J) and (K) show Nile
Red staining of the stem epidermis tissue isolated from dso-3 and WT, respectively.
Arrows in (J) indicate the inclusions. Bars represent in: A - 200nm; B, E - 1µm; D- 2µm;
G, H - 500nm; F, I - 1µm; J, K – 100 µm.
Figure 5. DSO 5'-upstrem region directed expression. (A) GFP expression driven by the
DSO 5'-upstream region in the embryo. Expression in radical tip is indicated by arrow.
GUS expression driven by the DSO 5'-upstream region detected (after 24 h incubation)
in: (B) three days old seedling, expression in root cap is indicated by an arrow, (C) a
higher magnification image of a stained root cap, (D) in the vasculature (v) of developing
root and at the lateral root primordia (lrp), (E) in the emerging lateral root, (F) in a 7 days
old seedling (arrow marks lateral root emergence sites), (G) in a 15 days old seedling,
expression in the lateral root (lr), main vein (mv) and basal segment (bs) of the leaf are
indicated by arrows, (H) in the in the inflorescence, (I) in the developing siliques. (J)
Confocal microscopy images of GFP expression driven by the DSO 5'-upstream region in
the adaxial leaf epidermis (i- autofluorescence; ii- GFP signal; iii- merge of GFP with
transmission; iv- merge between autofluorescence and GFP). The GFP signal indicated
by arrow. The blue signal marks autofluorescence in the cuticular ledges. (K) Confocal
microscopy images of GFP expression driven by the DSO 5'-upstream region in the
adaxial leaf epidermis showing GFP signal in the trichome base (indicated by arrow) and
support cells. (L) Images of GFP expression driven by the DSO 5'-upstream region in a
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37
free hand stem cross section. Arrows indicate GFP signal in epidermis. GFP signal was
also detected in other stem tissues. The bar in (A) is 50 µm.
Figure 6. Localization of DSO-GFP protein fusion to the plasma membrane of epidermal
cells. In (A) to (D), confocal microscopy of epidermal protoplasts derived from plants
harboring the promoterDSO::GFP-DSO construct (GFP fused in the N-termini) are
shown. Protoplasts were prepared from stem epidermis enriched tissue and analyzed for
DSO subcellular localization. Images were acquired through: (A) GFP filter, (B)
chlorophyll filter, (C) transmission and (D) a merge between (A) and (C). In (E) to (H),
whole mount confocal microscopy of leaves derived from plants harboring the
promoterDSO::GFP-DSO construct (GFP fused in the N-termini). Images were acquired
through: (E) GFP filter, (F) chlorophyll filter, (G) transmission and (H) a merge between
(E) and (F). FM4-64 (red signal) is a plasma membrane marker and was used in (F).
FM4-64 was used in (H) for co-localization with the GFP signal. Arrows indicate GFP in
(E), (H) and FM4-64 in (F), (H). In (I) to (L), confocal microscopy of stem cross
sections of plants harboring the promoterDSO::GFP-DSO construct. Images were
acquired through: (I) GFP filter, (J) chlorophyll filter, (K) is the transmission and (L) a
merge between (I), (J) and (K). Whole mount confocal microscopy of stems derived
from plants harboring the promoterDSO::GFP-DSO construct (GFP fused in the N-
termini). Images were acquired through: (M) GFP filter, (N) chlorophyll filter, and (O) a
merge between (M) and (N).
Figure 7. Reduced epicuticular wax and cutin monomers load in dso-3 plants. (A) Stem
wax load of dso-3 plants versus WT, two asterisks signify P<0.01 and one is P<0.05. (B)
Cutin monomers load of dso-3 plants versus wild-type (WT). The differences were
significant between all bars with P<0.01. Error bars are standard deviation in both (A)
and (B). For identities of major cutin monomers identified in leaf cuticles of dso-3 and
WT plants see Table II.
Figure 8. Induction of DSO and related genes expression by different stresses and
sensitivity of the dso-1 lines to salinity. (A) Semi-quantitative RT-PCR experiments
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38
showing DSO (WBC11), CER5 (WBC12) and WBC13 expression under 200 mM NaCL
and 50 µM ABA treatments. The β-actin gene served as a control for equal loading of
cDNA. (B) Wound induction detected in plants expressing GUS driven by the DSO 5'-
upstream region. (C) The decrease in DSO expression results in salt susceptibility as
detected in two weeks old dso-1 seedlings exposed to 200 mM NaCl for 4 days and (D) a
wild-type seedling exposed to the same treatment.
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39
Table I: GC-MS analysis of wax monomers load of dso-3 mutants and WT plants. WT dso-3
Compound class mean µg/cm2 sd mean µg/cm2 sd
Alkanes 11.46 1.02 2.05 0.19
Secondary alcohols 1.87 0.17 1.12 0.20
Ketones 3.79 0.90 1.45 0.43
Primary alcohols 1.56 0.07 1.43 0.07
Fatty acids 0.03 0.01 0.00 0.00
Aldehydes 0.25 0.02 0.10 0.02
Wax esters 0.30 0.02 0.31 0.05
Unknown 0.41 0.01 0.31 0.06
Total wax load 19.66 2.22 6.77 1.02
Data presented here is the average of 3 replicates; sd- is the standard deviation.
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40
Table II: List of cutin monomers identified after GC-MS analysis with their respective
concentrations in dso-3 mutants and WT plants.
WT des-3
ng/cm2 ng/cm2N Alkan-1-oic acids mean sd mean sd P value change vs. WT1 C18 Octadecenoic acid(1) 1.48 0.10 0.76 0.11 P<0.01 ↓
2 C18 Octadeca(dien+trien)oic acid(2,3) 17.37 1.54 6.78 1.01 P<0.01 ↓
3 C22 Docosanoic acid 3.53 0.43 1.25 0.07 P<0.01 ↓
4 C24 Tetracosanoic acid 4.20 0.36 1.94 0.03 P<0.01 ↓
total 26.58 2.43 10.73 1.212-Hydroxy acids
5 C16 2-Hydroxy-hexadecanoic acid 4.57 0.06 1.75 0.08 P<0.01 ↓
6 C20 2-Hydroxy-eicosanoic acid 1.51 0.12 0.53 0.06 P<0.01 ↓
7 C22 2-Hydroxy-docosanoic acid 7.92 0.08 3.49 0.15 P<0.01 ↓
8 C23 2-Hydroxy-tricosanoic acid 1.20 0.11 0.56 0.06 P<0.01 ↓
9 C24 2-Hydroxy-tetracosenoic acid(1) 23.77 0.56 5.98 0.38 P<0.01 ↓
10 C24 2-Hydroxy-tetracosanoic acid 28.79 0.64 10.17 0.09 P<0.01 ↓
11 C25 2-Hydroxy-pentacosenoic acid(1) 0.94 0.12 0.27 0.01 P<0.01 ↓
12 C25 2-Hydroxy-pentacosanoic acid 1.40 0.08 0.66 0.03 P<0.01 ↓
13 C26 2-Hydroxy-hexacosenoic acid(1) 2.30 0.11 0.74 0.02 P<0.01 ↓
14 C26 2-Hydroxy-hexacosanoic acid 9.45 0.04 4.31 0.13 P<0.01 ↓
total 81.85 1.94 28.46 1.01ω-Hydroxy acids
15 C16 16-Hydroxy-hexadecanoic acid 1.95 0.09 0.38 0.04 P<0.01 ↓
16 C17 17-Hydroxy-heptadecanoic acid 1.90 0.19 0.31 0.01 P<0.01 ↓
17 C18 18-Hydroxy-octadecadienoic acid(2) 1.80 0.11 0.60 0.10 P<0.01 ↓
18 C18 18-Hydroxy-octadecatrienoic acid(3) 1.95 0.16 0.42 0.09 P<0.01 ↓
total 7.59 0.55 1.71 0.25
α,ω-Dicarboxylic acids19 C16 Hexadecane-(1,16)-dioic acid 13.89 1.41 1.91 0.12 P<0.01 ↓
20 C18 Octadecane-(1,18)-dioic acid 2.98 0.26 0.59 0.02 P<0.01 ↓
21 C18 Octadecen-(1,18)-dioic acid(1) 9.16 0.89 1.69 0.06 P<0.01 ↓
22 C18 Octadecadien-(1,18)-dioic acid(2) 29.94 1.88 6.69 0.73 P<0.01 ↓
total 55.96 4.43 10.88 0.94 mid-chain oxygenated fatty acids 21.03 1.35 5.57 0.31
Unknown aliphatics 10.42 0.46 3.67 0.28Unidentified compounds 8.16 0.46 2.94 0.38
sum total 211.58 11.63 63.96 4.38 Data presented here is the average of 3 replicates; sd- is the standard deviation.
dso-3
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Figure 1
b ca
f
lb
fb
B C E
G H
LK M
JI
N
kj
on
Locus At1g17840.1
5‘- 6142603 3‘- 6146514
SALK_line_72079RNAi target:871-1159bp
Exon II Exon V
A
dso-1 dso-1 dso-3
dso-1 WT dso-2 dso-3
WT dso-2 / cer5
1 2
3
1 2 3 4
D
dso-2
WT
F
dso-2
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Figure 2
a
a
j
l
A
FE
DCB
M N O P
G
LKJI
H
I J
n
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Figure 3
WT dso-1 WT dso-2 WT dso-30
5
10
15
20
**
root
bra
nchi
ng n
umbe
r
****
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Figure 4
▼▼ ▼cutcut
▼
▼ cutcut cut
A B C
D E
G
F
IH
cut
cut
KJ
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Figure 5
lrp
v
C D
E F Glr
bs
mv
BA
N
O
H
LK
I J
i ii
iii iv
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Figure 6I
A B
C D
E F
G Hc
C D
N
K L
M
O
JIBA
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Figure 7
B
1 1 2 2 3 3 4 4 5 5 6 6 7 7 8 8 9 9 10 10 11 11 12 12 13 13 14 14 15 15 16 16 17 17 18 18 19 19 20 20 21 21 22 220
5
10
15
20
25
30
35
40
alkan-1-oic acids 2-hydroxy acids ω-hydroxy acids α ,ω -dicarboxylic-acids
ng/ c
m2
cutin component
C27 C27C29C29C31C31 C29C29C31C31 C29C29 C24C24C26C26C28C28C30 C28C28C32C32 C30 C40C40C42C42C44C440.00
1.25
2.50
3.75
5.00
6.25
7.50
8.75
10.00
11.25
12.50
13.75
15.00
alkanes secondary alc. ketone primary alc. aldehydes f. acids wax esters
µ g /
cm2
wax component
A
wild-type dso-3
wild-type dso-3
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Figure 8A
DSO / WBC11
WBC13
CER5 / WBC12
no stress NaCl 200mM ABA 50µM
β-actin
B C D
w
ww
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