The Arabidopsis DESPERADO AtWBC11 Transporter Is · The Arabidopsis DESPERADO/AtWBC11 Transporter Is Required for Cutin and Wax Secretion1[C][W] David Panikashvili, Sigal Savaldi-Goldstein,

The Arabidopsis DESPERADO/AtWBC11 Transporter IsRequired for Cutin and Wax Secretion1[C][W]

David Panikashvili, Sigal Savaldi-Goldstein, Tali Mandel, Tamar Yifhar, Rochus B. Franke, Rene Hofer,Lukas Schreiber, Joanne Chory, and Asaph Aharoni*

Department of Plant Sciences, Weizmann Institute of Science, Rehovot 76100, Israel (D.P., T.M., T.Y., A.A.);Plant Biology Laboratory and Howard Hughes Medical Institute, Salk Institute, La Jolla, California 92037(S.S.-G., J.C.); and Institute of Cellular and Molecular Botany, Department of Ecophysiology, University ofBonn, D–53115 Bonn, Germany (R.B.F., R.H., L.S.)

The cuticle fulfills multiple roles in the plant life cycle, including protection from environmental stresses and the regulation oforgan fusion. It is largely composed of cutin, which consists of C16-18 fatty acids. While cutin composition and biosynthesis havebeen studied, the export of cutin monomers out of the epidermis has remained elusive. Here, we show that DESPERADO(AtWBC11) (abbreviated DSO), encoding a plasma membrane-localized ATP-binding cassette transporter, is required for cutintransport to the extracellular matrix. The dso mutant exhibits an array of surface defects suggesting an abnormally functioningcuticle. This was accompanied by dramatic alterations in the levels of cutin monomers. Moreover, electron microscopy revealedunusual lipidic cytoplasmatic inclusions in epidermal cells, disappearance of the cuticle in postgenital fusion areas, and alteredmorphology of trichomes and pavement cells. We also found that DSO is induced by salt, abscisic acid, and wounding stresses andits loss of function results in plants that are highly susceptible to salt and display reduced root branching. Thus, DSO is not onlyessential for developmental plasticity but also plays a vital role in stress responses.

One of the most critical adaptations of plants to aterrestrial environment 450 million years ago was theformation of their surface, the cuticle. The cuticularlayer plays multiple roles in plants, including the reg-ulation of epidermal permeability and nonstomatal wa-ter loss and protection against insects, pathogens, UVlight, and frost (Sieber et al., 2000). It also functions innormal plant developmental processes, including theprevention of postgenital organ fusion and pollen-pistilinteractions (Lolle et al., 1998).

The major component of the cuticle is cutin, which isa polyester insoluble in organic solvents consisting of

oxygenated fatty acids with a chain length of 16 or 18carbons. Embedded in the cutin matrix are cuticularwaxes, which are complex mixtures of very-long-chainfatty acid (VLCFA; .C24) derivatives: aldehydes, ke-tones, primary and secondary alcohols, fatty acids,and wax esters (Kunst and Samuels, 2003). In manyspecies, they also include triterpenoids and other sec-ondary metabolites, such as sterols, alkaloids, phenyl-propanoids, and flavonoids. The cuticular waxes arearranged into an intracuticular layer in close associationwith the cutin matrix, as well as an epicuticular filmexterior to this, which may include epicuticular waxcrystals (Jetter et al., 2000). Recently, 2-hydroxy- anda,v-dicarboxylic fatty acids have been reported as thecharacteristic monomers of cutin in Arabidopsis (Arab-idopsis thaliana; Bonaventure et al., 2004; Franke et al.,2005). This cutin monomer composition is similar tothe aliphatic domain present in the Arabidopsis su-berin polymer (Franke et al., 2005). Suberin is part ofthe plant apoplastic barrier that prevents uncontrollednutritional and water loss, strengthens cell walls, andprovides protection from pathogens. In Arabidopsis,suberin depositions were detected in the endodermisof primary roots and the periderm of mature roots.

Postgenital fusion is a unique phenomenon that oc-curs when alterations in cuticle properties cause aug-mentation of the contact responsiveness. During plantdevelopment, organ fusion is tightly regulated, and thecuticle plays a vital role in either preventing or permit-ting fusions. Postgenital organ fusion occurs most com-monly during reproductive development (e.g. duringcarpel formation in angiosperms). One of the character-

1 This work was supported by the Adolfo and Evelyn BlumCareer Development Chair (to A.A.), by the Israeli Ministry ofScience (Eshkol fellowship for postdocs to D.P.), by the William Z.and Eda Bess Novick Young Scientist Fund (grant to A.A.), by the Y.Leon Benoziyo Institute for Molecular Medicine (to A.A.), by theUnited States-Israel Binational Agricultural Research and Develop-ment Fund (fellowship to S.S.-G.), by the Salk Institute (fellowship toS.S.-G.), by the National Research Initiative of the U.S. Department ofAgriculture Cooperative State Research, Education, and ExtensionService (grant to J.C.), and by the German Research Society (grant no.SCHR506/7–1 to L.S. and R.F.).

* Corresponding author; e-mail [email protected] author responsible for the distribution of materials integral to

the findings presented in this article in accordance with the policydescribed in the Instructions for Authors (www.plantphysiol.org) is:Asaph Aharoni ([email protected]).

[C] Some figures in this article are displayed in color online but inblack and white in the print edition.

[W] The online version of this article contains Web-only data.www.plantphysiol.org/cgi/doi/10.1104/pp.107.105676

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istic features of organ fusions is that adhesion of cellwalls is often accompanied by disappearance of thecuticle in the contact area (Lolle et al., 1998).

To date, nearly 20 Arabidopsis mutants displayingpostgenital fusions have been identified and for lessthan one-half of them has a corresponding gene prod-uct been associated (Lolle et al., 1998; Tanaka andMachida, 2006). One of the most known organ fusionmutant is the Arabidopsis fiddlehead (fdh). The FDHgene encodes a lipid biosynthetic enzyme that actsthrough the fatty acid elongation pathway and mightbe involved in cutin monomer biosynthesis. The fdhmutant leaves supported wild-type pollen germinationon their surfaces and showed increased permeability ofthe cuticle to the toluidine blue dye. In addition, fdhmutants exhibited an enhanced rate of chlorophyllleaching from leaves submerged in alcoholic solution(Lolle and Cheung, 1993; Yephremov et al., 1999; Pruittet al., 2000). Another mutant, abnormal leaf shape1 (ale1),showed defective cuticle in embryos and juvenileplants and, as a result, exhibited excessive water lossand organ fusion. The corresponding gene belongs to alarge family of subtilisin-like Ser proteases in Arabi-dopsis that are typically involved in intercellular sig-naling, converting their substrates to active or inactiveforms (Tanaka et al., 2001). The phenotypes of the ale1mutants depend on the genetic background, and theycould be observed in the Landsberg erecta backgroundbut not in the Columbia and Wassilewskija ecotypebackgrounds (Watanabe et al., 2004). The double mu-tant of ale1 (in Wassilewskija) and the Arabidopsishomolog of crinkly4 (acr4) resulted in one-half of theseedlings showing deformed cotyledons and severelyfused leaves. The authors suggested that ACR4 andALE1 synergistically affected the epidermis and thatACR4 plays a major role in the differentiation of epi-dermal cells in both vegetative and reproductive tis-sues. The maize crinkly4 (cr4) mutation shows graft-likefusions between organs, and the CR4 gene encodes aputative receptor kinase that might generate a signal forepidermal cell differentiation (Jin et al., 2000; Becraftet al., 2001; Tanaka et al., 2002).

A Cyt P450 monooxygenase, CYP86A8, catalyzes thev-hydroxylation of C12-18 fatty acids when assayed invitro. The CYP86A8 loss-of-function mutant, lacerata(lcr), showed severe cuticle defects, as evidenced byepidermal ruptures and postgenital fusions (Wellesenet al., 2001). A different gene, HOTHEAD (HTH), puta-tively encoding an oxidoreductase, was suggested tobe involved in the formation of a,v-dicarboxylic fattyacids because the hth-12 mutant allele showed de-creased load of these acids (Kurdyukov et al., 2006b).In the hth mutant, the majority of organ fusion eventsoccur during floral development (Lolle et al., 1998;Krolikowski et al., 2003). Interestingly, the HTH gene isnot epidermis specific, and its involvement in metab-olism of additional compounds not essential for con-struction of the cuticle is not yet clear.

Chen et al. (2003) reported the isolation of the WAX2gene and interpreted it to be required for both cutin

and cuticular wax deposition. The cuticular membraneof wax2 weighed less and was thicker, disorganized,and less opaque. The total wax load on leaves andstems was decreased to nearly 80%, showing a reduc-tion in the decarbonylase pathway products and anincrease in the acyl reduction pathway products. TheWAX2 protein contains certain regions with homologyto sterol desaturases and short-chain dehydrogenases/reductases. It was suggested that it plays a metabolicrole in both cutin and wax synthesis. The cloning andcharacterization of the same gene (termed YORE-YORE)was described by Kurata et al. (2003), and the yre mu-tant showed organ adhesion. The authors suggestedthat YRE might encode an enzyme catalyzing the for-mation of aldehydes in the wax decarbonylation path-way. Alterations to the fatty acid precursor pool couldalso result in plants showing organ fusion phenotypes.The enzyme acetyl-CoA carboxylase catalyses the ATP-dependant formation of malonyl-CoA. Acetyl-CoAcarboxylase activity in the cytosol generates a malonyl-CoA pool that is required for a wide range of reactions,including VLCFA elongation, that are incorporatedinto cutin and waxes. Weak gurke and pasticcino3 mu-tant alleles that correspond to a defect in the ACC1gene showed abnormal fused leaves that were oftenvitrified when plants were grown in vitro (Faure et al.,1998). A strong organ fusion phenotype was also seenin transgenic plants raised by Sieber et al. (2000) thatexpressed a fungal cutinase in Arabidopsis. Their resultssuggest that an intact cutin layer is crucial for prevent-ing organ fusions.

The synthesis of cuticle constituents occurs in theepidermis layer from which they are transported outto the plant surface. Recently, the first clue to the ex-port mechanism of cuticular lipids through the plasmamembrane was provided by the characterization of thecer5 Arabidopsis mutant (Pighin et al., 2004). The CER5gene encodes an ATP-binding cassette (ABC) trans-porter localized in the plasma membrane. Apart fromthe typical reduction in stem cuticular wax load (cerphenotype), the cer5 knockout mutant accumulatedsheet-like inclusions in the wax-secreting cells. Epi-dermal peel staining and observation with light mi-croscopy suggested that they are lipidic in nature. Inaddition, the overall levels of fatty acid were not al-tered, and this provided evidence that only wax trans-port was affected and not VLCFA biosynthesis. TheCER5 gene expression was detected in all plant organsexamined, including stems, leaves, siliques, flowers, androots. This was unexpected because the cer5 pheno-type is confined to leaves and stems. It was thereforesuggested that additional transporters must be in-volved in delivering wax components. In Arabidopsis,there are over 120 putative ABC transporters; 29 ofthem, including CER5, belong to the white-browncomplex (WBC) subfamily (Sanchez-Fernandez et al.,2001). In human and Drosophila, members of this fam-ily secrete cholesterol and plant sterols and play a rolein the steroid hormone pathway, respectively (Berge et al.,2000; Hock et al., 2000). In Arabidopsis, ABC transporters

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are implicated in the transport of a wide range of sub-strates, including auxin, Suc, and mono- and divalentions. They play a fundamental role in heavy metalstransport, resistance to xenobiotics, and different aspectsof plant development, including regulation of stomatalopening and closure (Schulz and Kolukisaoglu, 2006).

While the activity of CER5 could explain the trans-port of wax monomers out of the epidermal cells, themechanism responsible for cutin monomer transportremained unknown. In this article, we describe theDESPERADO (DSO) gene putatively encoding a WBC-subtype ABC transporter. We provide several lines ofevidence showing that DSO is vital for the export ofboth cutin and wax monomers to the surface of Arab-idopsis plants. The various DSO phenotypes and thefact that its expression was not confined to vegetativeorgans but was also detected in the root suggest that itmight also be involved in the transport of other typesof lipids. Such a lipid molecule could be, for example,suberin that shows chemical analogy to cutin and istypically deposited in plant roots. The results obtainedthrough this study also demonstrate that DSO is notonly vital for proper plant development but also toplants’ response to various stresses such as salinityand mechanical wounding.

RESULTS

Phenotypes of the DSO Loss-of-Function Lines

In human and Drosophila, members of the ABC trans-porters family play a role in the transport of lipid sub-strates (Pohl et al., 2005). To unravel the possible roleof ABC transporters in plant lipid transport, we sys-tematically generated, in Arabidopsis, RNA interfer-ence (RNAi) lines for more than 20 ABC transportersgenes. All members investigated belonged to the WBCsubfamily (Sanchez-Fernandez et al., 2001). One RNAiline (dso-1), targeted to silence the At1g17840 gene (Fig.1A), showed an array of morphological phenotypesincluding inter-organ postgenital fusions (Fig. 1, B–D)that resembled those of mutant plants altered in theircuticle (Yephremov et al., 1999; Chen et al., 2003;Krolikowski et al., 2003; Kurata et al., 2003; Schnurret al., 2004; Kurdyukov et al., 2006a). The DSO protein(AtWBC11; Sanchez-Fernandez et al., 2001) is a memberof a small group of WBC transporters that includes thepreviously described CER5 wax transporter (AtWBC12;Pighin et al., 2004; Supplemental Fig. S2), AtWBC15/22,AtWBC13, and AtWBC3. DSO shows the highest iden-tity to CER5 (52% at the amino acid level) and slightlyless homology to AtWBC15/22 (51% identity) andAtWBC13 (48% identity). The closest homolog of theCER5 wax transporter is AtWBC15/22 (85% identity;Supplemental Table S3).

The dso-1 plant phenotype was extremely severe asmost of the plants were strongly retarded in growthand upon maturation produced multiple, thin, and shortinflorescence stems (a bushy phenotype) probably due

to the loss of apical dominance. The fusion of organs indso-1 mutants often resulted in rosette leaves that weremisshapen and torn. A toluidine dye uptake test (Tanakaet al., 2004) suggested malformation of the dso-1 mu-tant cuticle as they displayed strong coloration after2 min of immersion in the dye, whereas no staining wasobserved in wild-type plants (Fig. 1, G and H). The leafsurfaces of several cuticular mutants were previouslyshown to support wild-type pollen germination (Lolleand Cheung, 1993; Lolle et al., 1998; Sieber et al., 2000;Wellesen et al., 2001; Kurdyukov et al., 2006b). Scan-ning electron microscopy (SEM) revealed that fullyexpanded rosette leaves of dso-1 plants do not supportwild-type pollen germination (data not shown). InArabidopsis rosette leaves, the vascular tissue is com-posed of the main vein in the middle of the leaf bladethat is interconnected by secondary and higher orderveins, forming a complex network. The vascular pat-terns in the dso-1 mutant appeared to be altered com-pared to the wild-type leaf patterns (Fig. 1, J and K). Inthe dso-1 leaf blade, fewer tertiary and quaternary veinswere observed. Moreover, the veins along the edge ofthe leaf formed a discontinuous circle.

We also generated misexpression lines by expressingDSO under the control of the 35S cauliflower mosaicvirus (CaMV) promoter (lines termed dso-2) and iden-tified a T-DNA insertional line (SALK_072079; Fig. 1A)in the DSO gene (lines termed dso-3). Semiquantitativereverse transcription (RT)-PCR was performed for allthree loss-of-function mutant genotypes to determinethe levels of the DSO transcript. In the dso-1 and dso-2lines, a significant reduction in transcript levels wasevident, whereas in the dso-3 line no transcript wasdetected, indicating that dso-3 is a null mutant (Sup-plemental Fig. S1). The results with misexpressionsuggested that instead of overexpression we obtainedcosuppression (detected in one-third of the primarytransformants).

In Supplemental Table S2, we depicted the differentphenotypes and their degree of penetration among thethree DSO loss-of-function genotypes. Overall, the dso-2and dso-3 plants showed similar phenotypes to the onesobserved in the dso-1 RNAi line (Fig. 1) but with dif-ferent levels of penetration. The cosuppression dso-2lines had a relatively mild phenotype compared to dso-1and dso-3, as they developed almost regular inflores-cence stems that frequently had a glossy cer-type phe-notype (Jenks et al., 1995; Fig. 1F). In some cases, cer-typephenotypes were observed only in certain stem partsbut not others in the same plant. The dso-2 plantsshowed notched rosette leaves. The majority of dso-1and dso-3 seedlings grown in tissue culture developedunusual, callus, or stigmatic-like protrusions from epi-dermal cells (Fig. 1E). A similar phenotype was ob-served previously in transgenic Arabidopsis plantsexpressing a fungal cutinase (Sieber et al., 2000). Leaf(Fig. 1M) and flower morphology was affected in allthree 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

DESPERADO/AtWBC11 Transporter, Cutin and Wax Secretion

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toluidine blue dye solution, they showed increasedstaining compared to wild-type seeds (data not shown),suggesting impaired integrity of the seed surface. Thestem cer-type phenotype detected by visual inspectionwas in agreement with the dramatic decrease in loadof epicuticular wax crystals on dso-2 mutant stem sur-faces observed by SEM (Fig. 2, A and B).

SEM examination of leaf surfaces uncovered notablephenotypes in the dso lines. Apart from random rup-tures in the epidermis (Fig. 2C), they developed dehy-drated trichomes with shortened stalks and irregularbranching patterns and they were often collapsed (Fig.2, D–F). Misshapen, asymmetric stomata and abnor-mal leaf pavement cell patterns were regularly observed(Fig. 2, I and J).

Alterations in reproductive organ morphology werealso detected in dso plants examined by SEM. Flowershad curved petals and distorted anther filaments (Fig.2, G and H). Moreover, the typical petal abaxial epi-dermis conical cells were variable in size and misshapen

in the petal folding area (Fig. 2, K and L). Pollen grainswere often absent from the stigmatic papillary cells,and they were often shriveled (Fig. 2, M–P). Alexanderstain for a pollen viability test showed that 23% of dso-3pollen is unviable and shriveled (data not shown). Wealso performed reciprocal crosses between dso-3 plantsand wild-type plants. When dso-3 plants were used asmale, short semi-sterile siliques were obtained. On theother hand, when dso-3 flowers were pollinated withwild-type pollen, no fertilization occurred. From theseobservations, we can conclude that the dso-3 sterilityoriginates from both defective male and female repro-ductive organs. The descendants of the backcrossed dso-3homozygous plants displayed the same phenotype, ex-cluding the possibility that the phenotype originatedfrom the background mutations. The effect of decreas-ing DSO transcript levels was also evidenced in below-ground tissues, as dso plants displayed reduced amountsof lateral roots compared to wild-type plants (Fig. 3; Sup-plemental Fig. S3).

Figure 1. The dso loss-of-function mutant phenotypes. A, Scheme of the DSO locus (At1g17840) depicting the RNAi targetsequence in the second exon and T-DNA insertion located in the fifth exon of the dso-3 line. B, Inter-organ postgenital fusions inthe dso-1 mutant plant. A fusion area between inflorescence and leaf is indicated by an 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 tissueculture. F, A cer phenotype in dso-2 plant stem (left) versus wild-type (WT) stem (right). G and H, dso-1 (G) and the wild type (H),1-month-old plants after 2-min immersion in toluidine blue staining solution. Sites of the dye penetration are indicated byarrows. I, A dso-2 flower phenotype. Underdeveloped petal is indicated by an arrow. J, Vasculature phenotype detected in dso-3rosette leaf. K, Wild-type rosette leaf vasculature. L, Siliques of dso-3 (1), dso-2 (2), and the wild type (3). M, Leaf phenotype of asevere dso-2 mutant line (1), mild dso-2 mutant line (2), dso-3 (3), and the wild type (4). N, The cer5-1/dso-2 (mild) doublemutant phenotype. Fusion between leaves is indicated by an arrow. Bars 5 2 mm in B and 1 mm in C.

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DSO and CER5 Might Function in the Same Pathway

To evaluate the interaction between CER5 and DSO,we crossed the mild phenotype dso-2 plants (withglossy stems, curved petals, and without organ fusionphenotype) with the cer5-1 mutant. As mentionedabove, the cer5-1 mutant does not display any addi-tional visual phenotype apart from its glossy cer-typestem. The dso-2/cer5 double mutant showed severefusion already at early developmental stages (Fig. 1N),suggesting an additive phenotype and providingevidence that these two genes could act in the samepathway.

Analysis of DSO and CER5 gene expression profilesusing Genevestigator (https://www.genevestigator.ethz.ch/at/) revealed similar expression patterns forthe two genes. Their expression is highest in seedlings,young leaves, and in the inflorescence. In the rootorgans, both genes display highest expression in lat-eral roots. Moreover, using the PRIME coexpressionsearch tool, we found that expression of both genes ishighly correlated (http://prime.psc.riken.jp/?action5coexpression_index).

DSO Loss-of-Function Lines Lack a Cuticular Layer inOrgan Fusion Areas and Contain Unusual LipidicCytoplasmatic Inclusions in Epidermal Cells

We used transmission electron microscopy (TEM) toexamine the changes in cuticle formation when twodso-1 rosette leaves are fused. These observations re-vealed that when complete fusion between the twoleaf surfaces occurred, absolute disappearance of thecuticular layer was detected, suggesting copolymeri-zation of adjacent cell walls (Fig. 4, A–C).

Unlike other cuticular mutants (Chen et al., 2003;Kurata et al., 2003; Xiao et al., 2004; Kurdyukov et al.,2006a, 2006b), dso mutant and transgenic line stemsand leaves were not altered in the cuticle ultrastruc-ture. On the other hand, detailed TEM inspection ofboth leaf and stems cells of dso-2 and dso-3 indicatedthat the transport of cuticular components might becompromised in the dso mutants. Unusual trilamellarcytoplasmatic inclusions were observed in epidermalcells of both leaves and stems of the mutants (Fig. 4,D–I). These structures could not be detected in epider-mis cells of wild-type leaves and stems and not in

Figure 2. SEM pictures. A and B, Stem wax load of dso-2 (A) and wild-type (B) plants. C, Occasional ruptures in the epidermis ofdso-2. D, Distorted and underdeveloped trichomes of dso-3. E, Collapsed and underdeveloped trichome of dso-1. Misshapensupport cells are indicated by an arrow. F, A wild-type trichome. G, Abnormal anther filament of a dso-3 flower. H, Curved petalsof a dso-3 flower. I and J, Light microscopy images showing aberrant pavement cell pattern and abnormal stomatal cells(indicated by arrows) in dso-3 (I) and abaxial leaf epidermis of wild-type plants (J). K and L, SEM of abnormal conical cells in theabaxial epidermis of a dso-3 flower petal (K) and conical cells in the abaxial epidermis of a wild-type flower petal (L). M and N,SEM micrographs of shriveled pollen grains in dso-3 lines (M) and pollen grains in the wild type (N). O and P, Stigmata papillae ofdso-3 (O) and those of the wild type (P); grains could not be detected in dso-3. Bars 5 20 mm in C and 50 mm in I and J.





other cell types of the mutants. To verify the natureof the cytoplasmatic inclusions, epidermal peels ofdso-3 stem were stained with Nile Red and observedwith fluorescence microscopy (Figs. 4, J and K). Theresults 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 examinedthe expression of GUS and GFP reporter genes underthe control of the DSO 5# upstream sequence. For GUSand GFP expression, 2,294 and 4,417 bp of the DSO 5#upstream region were transcriptionally fused to eitherone of these reporters, respectively, and reporter ac-tivity was evaluated in different tissues of T2 plants.Reporter GUS and GFP expression indicated that DSOis expressed in the seed coat and the endosperm (datanot shown) and later during embryo development inthe radical tip and vasculature (Fig. 5A). DSO expres-sion was detected in seedlings in the cotyledons, roottip, and young leaves (Fig. 5, B, C, F, and G). In bothyoung and mature leaves, expression was detected intrichomes and stomatal cells (Fig. 5, J and K) andweaker in the rest of the blade. The strongest expres-sion was detected in the main vein and the expandingbasal portion of the leaf (Fig. 5G). In roots of matureplants, DSO expression was clearly observed in lateralroot primordia and developing lateral roots but wasalso detected throughout the vasculature (Fig. 5, D–F).

Figure 3. The dso mutants exhibit a root-branching phenotype. Root-branching number of 3-week-old dso and wild-type (WT) plants isshown. Error bars represent SD. **, P , 0.01.

Figure 4. TEM pictures. A, The fusion areabetween dso-1 leaves in which the cuticles(cut) of either leaf align next to each other. Band C, Areas of leaf fusions in dso-1; redarrowheads mark areas in which the cuticleis absent; cuticles are indicated by arrows.D, F, G, and H, Unusual inclusions (arrows)in the epidermal cell cytoplasm of a dso-3leaf (D and F), in the epidermal cell of a dso-2stem (G), and in an epidermal cell of a dso-3stem (H). E, Epidermal cell cytoplasm ofwild-type leaf. I, Epidermal cell cytoplasmof wild-type stem. Fluorescence images in Jand K show Nile Red staining of the stemepidermis tissue isolated from dso-3 andthe wild type, respectively. Arrows in Jindicate the inclusions. Bars 5 200 nm inA, 1 mm in B and E, 2 mm in D, 500 nm in Gand H, 1 mm in Fand I, and 100 mm in J andK. [See online article for color version ofthis figure.]

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In the inflorescence, expression could be detected in allfloral organs, predominantly in the anthers, styles, andin young siliques (Fig. 5H). In the developing siliques,the strongest expression was detected in young si-liques (in the base and tip; Fig. 5I). Cross sections of the

inflorescence stem DSO expression showed that DSOexpression was not confined to epidermis, as it wasdetected in epidermal and mesophyll cells (Fig. 5L).Overall, developing rather than mature organs appearto express DSO.

Figure 5. DSO 5# upstream region directed expression. A, GFP expression driven by the DSO 5# upstream region in the embryo.Expression in radical tip is indicated by an arrow. B to I, GUS expression driven by the DSO 5# upstream region detected (after24-h incubation) in: 3-d-old seedling, expression in root cap is indicated by an arrow (B), a higher magnification image of astained root cap (C); in the vasculature (v) of developing root and at the lateral root primordia (lrp; D); in the emerging lateral root(E); in a 7-d-old seedling (arrow marks lateral root emergence sites; F); in a 15-d-old seedling, expression in the lateral root (lr),main vein (mv), and basal segment (bs) of the leaf is indicated by arrows (G); in the inflorescence (H); and in the developingsiliques (I). 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 GFPsignal is indicated by arrow. The blue signal marks autofluorescence in the cuticular ledges. K, Confocal microscopy images ofGFP 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 free-hand stemcross section. Arrows indicate GFP signal in epidermis. GFP signal was also detected in other stem tissues. Bar 5 50 mm in A.





DSO Protein Subcellular Localization

We also examined the subcellular localization of DSOby generating transgenic plants harboring a constructin which GFP was fused in frame to the N termini ofthe full-length DSO gene and expression was drivenby the DSO 5# upstream region. Two-thirds of the trans-formants displayed a cosuppression phenotype (datanot shown). Five T2 GFP positive plants with no signsof cosuppression were used for whole-mount confocalmicroscopy of leaves (Fig. 6, E–H), protoplast prepa-ration from stem epidermis-enriched tissues (Fig. 6,A–D), and preparation of free-hand stem cross sec-tions (Fig. 6, I–L). In protoplasts, the GFP signal was de-tected in the periphery of the cells, suggesting plasmamembrane localization. The use of protoplasts allowedus to exclude cell wall-specific expression. To determinewhether the observed fluorescence was associated withthe plasma membrane, we used a plasma membrane-specific marker (FM4-64). The results showed thatDSO is colocalized with FM4-64 (Fig. 6, E–H). Analysisof the stem cross sections of the same plants indicatedthat, contrary to the promoter-directed expression, GFPwas detected exclusively in the epidermis. Moreover,it was localized in a polar manner in the epidermisside facing the extracellular matrix (Fig. 6, I–L).

Changes in Cutin and Wax Monomer Composition inthe dso Mutants

To gain more knowledge on the precise role of DSO inthe transport of cuticle-associated lipids, we performedgas chromatography-mass spectrometry (GC-MS) anal-ysis of epicuticular waxes on the surface of dso-3 andwild-type plants. Chemical analysis revealed a 3-folddecrease in total stem wax load in dso-3 compared towild-type stems (6.77 6 1.02 mg/cm2 versus 19.66 62.22 mg/cm2; Table I). The C29 monomers, particularly,alkanes (11-fold decrease), ketone (2.6-fold decrease),and secondary alcohol (1.8-fold decrease), were largelyresponsible for this decrease (Fig. 7A).

The data gathered to this point suggested that DSOmight not only be required for wax but also for cutinmonomer transport. Consequently, we conducted GC-MS analysis of the previously reported Arabidopsis cutinconstituents, including regular fatty acids, 2-hydroxyfatty acids, v-hydroxy acids, and a,v-dicarboxylic acids(Bonaventure et al., 2004; Xiao et al., 2004; Franke et al.,2005). The results showed that total cutin monomerload per leaf area in dso-3 was reduced 3.3-fold com-pared to the wild type (63.96 6 4.38 ng/cm2 versus211.58 6 11.63 ng/cm2; Table II; Supplemental TableS1). Moreover, levels of all 22 detected cutin monomerswere dramatically decreased in dso-3 plants (Fig. 7B;Supplemental Table S1). Interestingly, chemical analy-sis of dso-2 leaf cuticular lipids showed reduction onlyin wax load, while no significant difference in cutinmonomer load was noted. However, cutin composi-tion in dso-2 was different from the wild type. Levels of2-hydroxy and v-hydroxy fatty acid levels were up-

regulated, whereas the levels of a,v-dicarboxylic acids(Bonaventure et al., 2004; Franke et al., 2005) were sig-nificantly reduced (Supplemental Fig. S4). Thus, DSOloss of function results in altered level and composi-tion of both cutin and wax monomers in the cuticle.

DSO Is Induced under Salt, Abscisic Acid, and

Wound Stresses

Salinity is a polymorphous stress that impedes plantdevelopment and viability through two shared effects:osmotic and nutritional. High salinity affects plantsthrough ion toxicity as well. To assess how DSO is im-plicated in these environmental stresses, we subjecteddso-1 seedlings to salinity stress (200 mM NaCl). We foundthat dso-1 seedlings are more susceptible to salinitystress than the wild type (Fig. 8, C and D). Using semi-quantitative RT-PCR, we evaluated DSO transcriptlevels in wild-type plants under the same conditionsand found up-regulation in DSO expression upon saltstress (24-h exposure; Fig. 8A).

Abscisic acid (ABA) is a universal plant hormonewidely implicated in adaptation to stress. It regulatesstomatal closure, and increasing evidence suggests thatit is involved in root branching (De Smet et al., 2006).Semiquantitative RT-PCR showed up-regulation of DSOtranscripts in RNA derived from seedlings treated for24 h with 50 mM ABA (Fig. 8A). DSO expression wasnot only induced by salinity and increased ABA levelsbut also upon mechanical wounding, as detected inleaves expressing GUS driven by the DSO 5# upstreamregion (Fig. 8B). We also examined the expression ofCER5 (AtWBC12) and AtWBC13 genes under the samesalt and ABA treatments (Fig. 8A). The results showthat while CER5 expression is induced in salt and ABAtreatments (similar to DSO), expression of WBC13 isinduced by salt but not by ABA.

DISCUSSION

Cutin is the third most abundant biopolymer onearth after lignin and cellulose. It is a major componentof the cuticle that covers all plant surfaces exposed toair. Despite its significance, little is known about thetransport of cutin monomers from their synthesis site inthe epidermal cells to the extracellular domain wherethe cuticle is assembled. This study demonstrates thatin Arabidopsis, DSO, a plasma membrane-localizedABC transporter, is required for proper export of cutinmonomers through the plasma membrane to the cuticle.It is also required for the transport of wax monomers, adifferent set of cuticular components, as reported ear-lier 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 becausea large number of the ABC transporters characterizedto date were able to handle several structurally differ-ent compounds (Yazaki, 2006). Moreover, the informa-tion regarding DSO gene expression and the array of

Panikashvili et al.




loss-of-function phenotypes suggest that the DSO pro-tein might also be associated with transport of otherwax- or cutin-like molecules.

The DSO protein sequence shares 52% identity withCER5 but, interestingly, an even higher level of sim-ilarity with the cotton (Gossypum hirsutum) GhWBC1protein (84% identity). GhWBC11 is highly expressedin developing cotton fiber cells, and its overexpressionin Arabidopsis resulted in plants with short siliquescontaining severely shriveled embryos and with onlyseveral seeds per silique (Zhu et al., 2003). A variableamount of suberin could be found at the cotton fiberbase that is typically deposited in concentric layers,

alternating with polysaccharides (Ryser, 1992). The rel-atively high similarity in sequence between DSO andGhWBC1, the overexpression phenotype, and the pres-ence of suberin and a thin cuticle with wax and cutincomponents in cotton fibers (Schmutz et al., 1996) sug-gest a similar role to the two transporter proteins inArabidopsis and cotton.

DSO expression was not confined to the epidermisof aerial parts as anticipated for a transporter of cu-ticular components. Its expression was also detected inother aerial plant organs and cell types as, for example,in the stem mesophyll cells. Interestingly, relativelystrong DSO expression was detected in lateral root

Figure 6. Localization of DSO-GFP protein fusion to the plasma membrane of epidermal cells. A to D, Confocal microscopy ofepidermal protoplasts derived from plants harboring the pDSOTGFP-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 wereacquired through: GFP filter (A), chlorophyll filter (B), transmission (C), and a merge (D) between A and C. E to H, Whole-mountconfocal microscopy of leaves derived from plants harboring the pDSOTGFP-DSO construct (GFP fused in the N termini).Images were acquired through: GFP filter (E), chlorophyll filter (F), transmission (G), and a merge (H) 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 colocalization with the GFP signal.Arrows indicate GFP in E and H and FM4-64 in F and H. I to L, Confocal microscopy of stem cross sections of plants harboring thepDSOTGFP-DSO construct. Images were acquired through: GFP filter (I), chlorophyll filter (J), transmission (K), and a merge (L)between I, J, and K. M to O, Whole-mount confocal microscopy of stems derived from plants harboring the pDSOTGFP-DSOconstruct (GFP fused in the N termini). Images were acquired through: GFP filter (M), chlorophyll filter (N), and a merge (O)between M and N.





primordia, the developing lateral root, and in the rootvasculature. Moreover, we also detected reduced rootbranching in the dso lines. In a different cuticular mu-tant, bodyguard (bdg), an increase in root branching wasobserved (Kurdyukov et al., 2006a). This is not the firststudy in which root expression of genes associated withcutin and wax metabolism is reported. Root expres-sion was also detected for the b-KETO ACYL-COASYNTHASE1 (KCS1; Todd et al., 1999), YRE (Kurataet al., 2003), BDG (Kurdyukov et al., 2006a), SHINE3(SHN3; Aharoni et al., 2004), HTH (Kurdyukov et al.,2006b), and CER5 (Pighin et al., 2004) genes. Fatty acid

analysis of the kcs1-1 mutant roots revealed a 2-foldincrease in a,v-dicarboxylic acids, and this result ledthe authors to suggest that KCS1 is not only implicatedin wax metabolism but also in the suberin biosynthesispathway (Todd et al., 1999). Apart from expression ofDSO in roots, two more lines of evidence suggest thatDSO is involved in transport of other lipid-derivedchemicals (such as suberin) that are similar in struc-ture to cuticular lipids. The first supporting evidence isthe altered leaf vascular patterns in the dso-3 mutantthat might be a result of changes in suberin depositionduring secondary growth in the vascular tissue.

A second point supporting this argument is the strik-ing similarity between the aliphatic monomer compo-sition of Arabidopsis cutin and suberin (Bonaventureet al., 2004; Franke et al., 2005). Indeed, a few recent stud-ies suggested that the long-chain a,v-dicarboxylic fattyacids are not only constituents of the cutin polyester inArabidopsis (Bonaventure et al., 2004; Kurdyukov et al.,2006b). They might play additional roles as a ‘‘suberin-like’’ network in the secondary cell wall or be requiredfor the cross linking that ensures the integrity of theprimary epidermis cell wall. Evolutionarily, it is feasi-ble that during the course of adaptation to terrestrialenvironments, plants modified the substrate specificityof a lipid transporter to a protein that could fulfill therequirements for constructing interface layers usingthree different building blocks, namely, cutin, wax, andsuberin. We therefore anticipate that in the near future,

Figure 7. Reduced epicuticularwax and cutin monomers load indso-3 plants. A, Stem wax load ofdso-3 plants versus the wild type. *,P , 0.05; **, P , 0.01. B, Cutinmonomer load of dso-3 plants ver-sus the wild type. The differenceswere significant between all barswith P , 0.01. Error bars are SD inboth A and B. For identities of majorcutin monomers identified in leafcuticles of dso-3 and wild-typeplants, see Table II.

Table I. GC-MS analysis of wax monomers load of dso-3 mutantsand wild-type plants

Data presented here are the average of three replicates.

Compound Class Wild Type dso-3

mean mg/cm2SD mean mg/cm2

SD

Alkanes 11.46 1.02 2.05 0.19Secondary alcohols 1.87 0.17 1.12 0.20Ketones 3.79 0.90 1.45 0.43Primary alcohols 1.56 0.07 1.43 0.07Fatty acids 0.03 0.01 0.00 0.00Aldehydes 0.25 0.02 0.10 0.02Wax esters 0.30 0.02 0.31 0.05Unknown 0.41 0.01 0.31 0.06Total wax load 19.66 2.22 6.77 1.02

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as suggested here for DSO and previously for KCS1(Todd et al., 1999), more genes associated with wax andcutin metabolism will also be recognized as playing arole in the biosynthesis and transport of other plantinterface components (e.g. suberin).

Full-size ABC transporters contain two ABC andtwo transmembrane domains in a single polypeptidechain (Schulz and Kolukisaoglu, 2006). On the otherhand, half-size transporters, such as the one encodedby DSO, achieve their functionality by combining twoABC-transmembrane domain units as homo- or het-erodimers in a membrane-bound transporter complex.One possible dimerization candidate for wax transportis CER5, although two other proteins in the same phy-logenetic clade (At3g21090, WBC15; and At1g51460,WBC13; see Sanchez-Fernandez et al., 2001) might alsoact as partners for the transport of cutin and othermolecules. For example, in Drosophila, dimerization ofthree half-transporter ABC proteins related in sequence,White with Brown and White with Scarlet, is requiredfor the transport of different eye pigment precursors

into pigment cells (Mackenzie et al., 2000). Further ex-periments should identify the interacting partners be-tween the transporters, their substrate specificity, andtheir mode of action.

With respect to their mode of action, ABC transportersmight actively expel the substrates into the extramem-brane space or through a side port of the transporter intothe upper leaflet of the plasma membrane bilayer in anATP-dependant process. Alternatively, they might actby turning over the substrates from the inner to the outerleaflet of the plasma membrane acting as an ‘‘hydro-phobic vacuum cleaner’’ (flippase activity; Chang andRoth, 2001). Another important question is: How docuticular lipids reach the ABC transporter localized inthe plasma membrane? The possible routes include: (1)they are picked up by fatty acid-binding proteins andrelocated to the transporter; or (2) relocation through avesicular pathway either by the formation of oleosomebodies coated by oleosin-like proteins or the formationof uncoated vesicles that contain the cuticular lipids inlipid rafts (Schulz and Frommer, 2004).

Table II. List of cutin monomers identified after GC-MS analysis with their respective concentrations in dso-3 mutants and wild-type plants

Data presented here are the average of three replicates.

n Compound Class Wild Type SD dso-3 SD P Value Change versus Wild Type

mean ng/cm2 mean ng/cm2

Alkan-1-oic acids1 C18 octadecenoic acid(1) 1.48 0.10 0.76 0.11 P , 0.01 Y2 C18 octadeca(dien1trien)oic acid(2,3) 17.37 1.54 6.78 1.01 P , 0.01 Y3 C22 docosanoic acid 3.53 0.43 1.25 0.07 P , 0.01 Y4 C24 tetracosanoic acid 4.20 0.36 1.94 0.03 P , 0.01 Y

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 Y6 C20 2-hydroxy-eicosanoic acid 1.51 0.12 0.53 0.06 P , 0.01 Y7 C22 2-hydroxy-docosanoic acid 7.92 0.08 3.49 0.15 P , 0.01 Y8 C23 2-hydroxy-tricosanoic acid 1.20 0.11 0.56 0.06 P , 0.01 Y9 C24 2-hydroxy-tetracosenoic acid(1) 23.77 0.56 5.98 0.38 P , 0.01 Y

10 C24 2-hydroxy-tetracosanoic acid 28.79 0.64 10.17 0.09 P , 0.01 Y11 C25 2-hydroxy-pentacosenoic acid(1) 0.94 0.12 0.27 0.01 P , 0.01 Y12 C25 2-hydroxy-pentacosanoic acid 1.40 0.08 0.66 0.03 P , 0.01 Y13 C26 2-hydroxy-hexacosenoic acid(1) 2.30 0.11 0.74 0.02 P , 0.01 Y14 C26 2-hydroxy-hexacosanoic acid 9.45 0.04 4.31 0.13 P , 0.01 Y

Total 81.85 1.94 28.46 1.01v-Hydroxy acids

15 C16 16-hydroxy-hexadecanoic acid 1.95 0.09 0.38 0.04 P , 0.01 Y16 C17 17-hydroxy-heptadecanoic acid 1.90 0.19 0.31 0.01 P , 0.01 Y17 C18 18-hydroxy-octadecadienoic acid(2) 1.80 0.11 0.60 0.10 P , 0.01 Y18 C18 18-hydroxy-octadecatrienoic acid(3) 1.95 0.16 0.42 0.09 P , 0.01 Y

Total 7.59 0.55 1.71 0.25a,v-Dicarboxylic acids

19 C16 hexadecane-(1,16)-dioic acid 13.89 1.41 1.91 0.12 P , 0.01 Y20 C18 octadecane-(1,18)-dioic acid 2.98 0.26 0.59 0.02 P , 0.01 Y21 C18 octadecen-(1,18)-dioic acid(1) 9.16 0.89 1.69 0.06 P , 0.01 Y22 C18 octadecadien-(1,18)-dioic acid(2) 29.94 1.88 6.69 0.73 P , 0.01 Y

Total 55.96 4.43 10.88 0.94Midchain oxygenated fatty acids 21.03 1.35 5.57 0.31Unknown 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





Wax load in, particularly, the C29 alkanes was dra-matically reduced in stems of both the dso and the cer5mutants. It should be noted that in the case of dso, levelsof several stem wax components were significantlyincreased in the mutant compared to the wild type (i.e.C27 alkanes, C31 secondary alcohols, and C24 primaryalcohols), suggesting compensation for the loss of othercuticle components. Leaves and fruit of a transposoninsertion mutant in the tomato LeCER6 encoding aVLCFA elongase (KCS) were deficient in n-alkanes andaldehydes with chain lengths beyond C30. In the sameplants, much higher levels of pentacyclic triterpenoids(a-, b-, and d-amyrin) were detected, suggesting com-pensation for the reduction in aliphatics (Vogg et al.,2004). In contrast to the cer5 knockout mutant showingthe typical glossy/cer-like stem phenotype with noeffect on plant architecture, the dso mutants exhibited arange of dramatic phenotypes in nearly every plantorgan examined. The lesion in DSO had a dramaticeffect on epidermal cell development, including alter-ations to trichomes, stomata, and pavement cells. In thecase of trichomes, they were collapsed and underde-veloped. Similar phenotypes were observed in trichomesof several mutants and transgenic plants involved inwax and cutin metabolism, including SHN1 overex-pression lines (Aharoni et al., 2004), fdh (Yephremovet al., 1999), lcr (Wellesen et al., 2001), cer10 (Zheng et al.,2005), and bdg (Kurdyukov et al., 2006a). It is intriguingthat SHN3, one of three AP2 domain transcriptionfactors suggested to act as activators of the wax bio-synthetic pathway in Arabidopsis, showed strong andspecific expression in the trichome support cells thatsurround the base of the trichome (Aharoni et al., 2004).The collapse of trichome phenotype detected in dso andbdg might be a result of altered development of thesupport cells and suggests that these cells might con-tain a unique component such as suberin that providesthem with the strength to hold the trichomes. This hy-

pothesis is further supported by an earlier report on thepresence of suberized cell walls in the boundary be-tween plants and secretory organs such as trichomes(Kolattukudy, 2001).

Basal or support cells of trichomes, cuticular ledges,and cuticles over anticlinal cell walls together provideaqueous pores for plants cuticles (Schonherr, 2006).The exact biological impact of the collapsed trichomessupporting cells of the dso mutants with regard to theaqueous solutes movement requires further investiga-tion. Because aqueous pores serve as a main gate forfoliar penetration of exogenously applied compoundsincluding agricultural chemicals, deciphering the ex-act role of cutin metabolism genes on cuticular aque-ous pores assembly in plants will have paramountsignificance for agriculture and ecophysiology.

A major phenotype of the dso loss-of-function geno-types was the occurrence of postgenital fusions thatinvolve surface contact between organs that have al-ready developed as individual entities (Verbeke, 1992).In dso, fusions could be noticed between different typesof vegetative and reproductive organs, including be-tween distal parts (even tips) of rosette leaves. Althoughmultiple mutants have been described that possespostgenital organ fusion (see introduction), one cannotidentify the factor(s) mediating this phenomenon. Theassortment of mutants exhibiting postgenital fusionsdescribed up to date differ in most of the parametersused to phenotype cuticular mutants, including in:permeability to a cationic dye, rate of chlorophyll leach-ing from leaves in alcoholic solution, rate of water loss,defects in pollen hydration, male sterility recovery un-der high humidity, glossy appearance, alterations to thecuticle ultrastructure, stomatal index, trichome num-ber and branching, and changes in the cuticle chemicalcomposition. This difference in phenotypes in the classof postgenital organ fusion mutants might be simplydue to the nature of mutations, gene redundancy, and

Figure 8. Induction of DSO and relatedgene expression by different stresses andsensitivity of the dso-1 lines to salinity. A,Semiquantitative RT-PCR experimentsshowing DSO (WBC11), CER5 (WBC12),and WBC13 expression under 200 mM

NaCL and 50 mM ABA treatments. Theb-actin gene served as a control for equalloading of cDNA. B, Wound inductiondetected in plants expressing GUS drivenby the DSO 5# upstream region. C and D,The decrease in DSO expression results insalt susceptibility as detected in 2-week-old dso-1 seedlings exposed to 200 mM

NaCl for 4 d (C) and a wild-type seedlingexposed to the same treatment (D).

Panikashvili et al.




the degree of phenotype penetration. However, it mayalso be that a single factor, either a not-yet-identifiedsignaling lipid-based molecule or a specific structuralchange in the cuticle or the epidermal cell wall (Nawrath,2006), could trigger this phenomenon. A more detailedcomparison between the dso and cer5 mutant pheno-types might aid in identifying the factor promotingpostgenital organ fusions because cer5 shows similaralterations in waxes compared to dso but does not ex-hibit fusion phenotypes.

In recent years, an increasing number of cuticle-related phenotypes and processes have been described(Nawrath, 2006). It is apparent that cuticle-associatedproteins not only play a role in plant development, butthey are also very active in the plant response to dif-ferent stress conditions. Our study suggests that DSOplays a vital role in stress response programs mediatedby the cuticle, including salt stress and wounding, asits expression was induced under these conditions.Supportive evidence for the importance of DSO func-tion in response to stress, particularly salt stress, wasprovided by experiments showing that dso-1 seedlingsare highly sensitive to salt treatment. Preliminary char-acterization of a gene trap line corresponding to DSOalso showed that it is also induced by multiple stresses,including ABA, high salt, and Glc (Alvarado et al.,2004). Like DSO, the expression of CER6 encoding aVLCFA-condensing enzyme was enhanced by osmoticstress and the presence of ABA (Hooker et al., 2002).

The role of DSO in stress response could be explainedby the need to alter surface structure upon water, salinity,and mechanical stress (Shepherd and Wynne Griffiths,2006). Leaf transpiration has stomatal and cuticularcom-ponents. Transpiration through the cuticle is largelydetermined by surface characteristics such as wax thick-ness and wax microstructure. Wax deposition that oc-curs rapidly within a few days is often a response towater stress, and stress-resistant plants often have thickerwaxes compared to susceptible ones. Increased wax de-position upon exposure to salinity stress was reportedfor several plants, including salt-sensitive jojoba (Sim-mondsia chinensis), and seems to be primarily a responseto water deficit (Mills et al., 2001). Moreover, in leaves ofsalt-sensitive jojoba, wax deposition is induced by ex-ogenous ABA (Mills et al., 2001). Finally, mechanicalstress such as wounding due to strong wind, rain drops,and leaf-to-leaf contact could also induce the formationof wax for reforming leaf structure (Shepherd and WynneGriffiths, 2006). More in relation to ABA and stressresponse, dso-1 mutant plants displayed reduced rootbranching number. Lateral root formation is essentialfor the adaptation of plants to changing environmentalchallenges such as increased osmotic stress and salinity.Growing evidence suggests that ABA is involved in theregulation of root branching (De Smet et al., 2006). In-terestingly, GUS expression driven by the DSO 5# up-stream region indicated DSO expression throughoutlateral root development.

This study adds another piece to the puzzle of howplants assemble their outermost surface. Nevertheless,

the mechanism of cuticle monomer transport from theirsite of synthesis to the membrane and further to theextracellular domain remains unclear.

MATERIALS AND METHODS

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-h-light/8-h-dark cycle and were in the

Arabidopsis (Arabidopsis thaliana) ecotype Columbia. 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 Web site).

DSO RNAi F1 plant seeds were stratified for 2 to 3 d at 4�C and subsequently

sown on Murashige and Skoog plates supplemented with 50 mg/mL kanamy-

cin and grown in a culture room under continuous light conditions at 20�C.

Two-week-old seedlings were subsequently transferred to soil. For the root-

branching experiment, 1-week-old transgenic plants displaying the typical

fusion phenotypes were transferred to vertically placed Murashige and Skoog

plates for additional 2 weeks growth. Only lateral roots branching out from the

main root were counted. For the salt stress assay, 2-week-old seedlings were

transferred to Murashige and Skoog plates supplemented with 200 mM NaCl

for an additional 1 week growth. Plant survival was monitored during 1 week

after application of the salt stress.

Generation of Plant Transformation Constructs andTransgenic Arabidopsis

For generating the DSO RNAi construct, a 298-bp genomic fragment was

amplified with sense (5#-AAAAAGCAGGCTCATATGTGACCCAAGACGA-

TAAC-3#) and antisense (5#-AGAAAGCTGGGTGCAGAAGCACTATCAAG-

ACCAC-3#) oligonucleotides, 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 in-

serted into BJ36 (Moore et al., 1998), under control of the 35S CaMV promoter,

and subsequently cloned into the pMLBART binary vector. For plant trans-

formation, inflorescences were dipped into Agrobacterium tumefaciens strain

GV3101 carrying the transgene construct as described (Clough and Bent, 1998).

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-week-old plants were bleached for 24 h in ethanol:acetic acid (6:1),

immersed in 70% ethanol for 1 h, mounted in chloralhydrate mixture (chloral-

hydrate: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 UsingSemiquantitative RT-PCR

Two to four rosette leaves of 3- to 4-week-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 (500 ng/1 mg) was transcribed to cDNA using oligo(dT)15 and AMV

reverse transcriptase (Chimerix). For the PCR reaction, 1 to 5 mL of was used

as a template for a 20-mL PCR reaction with the following primers: DSO sense

(5#-ATGTTACTCCTTGGGTCAGAG-3#), antisense (5#-ATTTCGGCACAATG-

CAAAC-3#) with expected band size of 399 bp; CER5 sense (5#-TGGGATG-

GAAGTGAGAAAGG-3#), antisense (5#-GAGCCAAGATCGATGTGTAG-3#)

with expected band size 193 bp; and WBC13 sense (5#-GGGGATTGTCACA-

GAAAGGA-3#), antisense (5#-TGACCCGACACAAATGGATA-3#) with ex-

pected band size 156 bp. PCR was started with a 94�C denaturation step for

2 min, followed by 45 s at 94�C, 45 s at 58�C, 1 min at 72�C, and 5 min 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 asReporter Gene

The DSO 5# upstream region (2,394 bp, termed pDSO) was amplified using

antisense (5#-NcoI-AAACCATGGCTCTTAAACCAAAACAGAGGATT-3#) and

sense (5#-BamHI-TTTAAGAATTAATTGTCTAAATAAC-3#) oligonucleotides,

excised with appropriate restriction enzymes, and subcloned into the pMAX

vector containing the GUS coding sequence and the NOS terminator. The

pDSO fragment together with the GUS gene was excised with PacI and AscI

and cloned into the binary vector pBIN1 (van Engelen et al., 1995). GUS

staining and embedding was performed according to Pekker et al. (2005). For

the 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 (4,417 bp) was amplified using sense (5#-BamHI-

AGGATCCCTCTTAAACCAAAACAGAGG-#3) and antisense (5#-PstI-CTG-

CAGGGTAAGTAATTTAGCAATTG-#3) oligonucleotides, and placed 5# to

the LHG4 gene in BJ36. The BJ36 was cut with NotI and subcloned into pART27.

To transactivate 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 488 nm, emission was at 500–530 nm for

GFP and 620–750 nm for chlorophyll).

SEM

Stems (second internodes from the bottom) were collected from wild-type

and dso-1, dso-2, or dso-3 plants after 5 to 6 weeks of growth. Leaves were

collected and fixated with glutaraldehyde using standard protocols and dried

using critical point drying. Samples were mounted on aluminum stubs and

sputter-coated with gold. SEM was performed using an XL30 ESEM FEG

microscope (FEI) at 5 to 10 kV.

TEM

Leaves and stems from 50-d-old plants were collected and processed using

a standard protocol (Weigel and Glazebrook, 2002). The spurr resin-embedded

samples were sectioned (70 nm) using an ultramicrotome (Leica) and ob-

served with a Tecnai T12 TEM (FEI).

Subcellular Localization of DSO and

Confocal Microscopy

For examination of the DSO protein subcellular localization, the fragment

containing the DSO 5# upstream region (4,417 bp) and the DSO cDNA were

translationally fused to GFP at N# termini. The DSO cDNA was amplified

using sense (5#-BamHI-TTTGGATCCCTACCATCTGCGAGCTCCATC-#3) and

antisense (5#-XhoI-AAACTCGAGAATGGAGATAGAAGCAAGCAG-#3) oli-

gonucleotides, excised with BamHI and XhoI, and cloned into the 10OPT

N#-GFP in the BJ36 vector (Moore et al., 1998). The DSO 5# upstream region was

amplified using sense (5#-KpnI-AAAGGTACCTAAGAATTAATTGTCTAAAT-

AAC-#3) and antisense (5#-XhoI-TTTCTCGAGCTCTTAAACCAAAACA-

GAGGATT-#3) oligonucleotides, and cut with KpnI and XhoI. The 10OPT

N#-GFP:DSO in BJ36 was cut with SalI, NotI, and a modified pBlueScript SK1

vector (Stratagene Cloning Systems) cut with NotI and KpnI. After a triple

ligation, the obtained vector was cut with PacI and AscI, and cloned into the

pBIN1 binary vector (van Engelen et al., 1995). Protoplasts from epidermis-

enriched stem segments were prepared as described elsewhere (Sheen, 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 to 525 nm for GFP and 620 to 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 recov-

ery of the GFP signal (Supplemental Movie S1).

Wax and Cutin Analysis

For wax analysis, the second internodes of 7-week-old plants (n 5 5) or

leaves of 4- to 5-week-old plants were cut and immersed twice in 5 mL of hexane

for 30 s at room temperature. The obtained solution containing the cuticular

waxes was spiked with 2 mg of tetracosane (Fluka) as an internal standard and

analyzed as described (Kurdyukov et al., 2006a). The extracted stem area was

calculated based on the measurements of stem length and upper and lower

diameter. Leaf area was assessed using the NIH ImageJ software. For cutin

analysis, 4-week-old mature leaves (n 5 3–4 for the wild type 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 10 mL of

methanol:chloroform (1:1, v/v) mixture for 14 d (solvent was changed daily).

Leaf material was dried, weighed (about 25–30 mg), 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 following oligonu-

cleotides: sense, 35S-out-CAATCCCACTATCCTTCG, and antisense, DSO

TGTCTGCTTGCTTCTATCTC (expected band size 361 bp).

Statistical Analysis

Data are presented as means 6 SD. Statistical significance was determined by

a Student’s t test. P values ,0.05 were considered to be statistically significant.

Sequence data from this article can be found in the GenBank/EMBL data

libraries under accession number 30685555.

Supplemental Data

The following materials are available in the online version of this article.

Supplemental Figure S1. Gene expression analysis of DSO and related

sequences in dso lines.

Supplemental Figure S2. Phylogenetic tree of the Arabidopsis WBC

subfamily.

Supplemental Figure S3. Root-branching phenotype of dso plants.

Supplemental Figure S4. Cuticular lipids analysis of dso-2 and wild-type

plants.

Supplemental Table S1. GC-MS analysis of cutin monomers of dso-3 and

wild-type plants.

Supplemental Table S2. Phenotype penetration in the DSO loss-of-

function lines.

Supplemental Table S3. Homology between the DSO/CER5 subclade of

WBC-type ABC transporters.

Supplemental Movie S1. GFP signal recovery in pDSOTGFP-DSO trans-

genic plants.

ACKNOWLEDGMENTS

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.

Received July 17, 2007; accepted October 11, 2007; published October 19, 2007.

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