The Arabidopsis DESPERADO AtWBC11 Transporter is · PDF fileThus, DESPERADO is not only...

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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



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



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



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



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.



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'), antisense (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).



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



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



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



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.



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. .



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.



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



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



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.



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.



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



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



Figure 2

a

a

j

l

A

FE

DCB

M N O P

G

LKJI

H

I J

n



Figure 3

WT dso-1 WT dso-2 WT dso-30

5

10

15

20

**

root

bra

nchi

ng n

umbe

r

****



Figure 4

▼▼ ▼cutcut

▼

▼ cutcut cut

A B C

D E

G

F

IH

cut

cut

KJ



Figure 5

lrp

v

C D

E F Glr

bs

mv

BA

N

O

H

LK

I J

i ii

iii iv



Figure 6I

A B

C D

E F

G Hc

C D

N

K L

M

O

JIBA



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



Figure 8A

DSO / WBC11

WBC13

CER5 / WBC12

no stress NaCl 200mM ABA 50µM

β-actin

B C D

w

ww

.plantphysiol.orgon M

ay 20, 2018 - Published by

Dow

nloaded from

Copyright ©

2007 Am

erican Society of P

lant Biologists. A

ll rights reserved.


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