The ultrastructure of the cuticle in mature plant organs

The cuticle is typically organized into distinct layers of variable thickness. There is a large amount of structural diversity among different plant species and organs (Jeffree, 2006; Buda et al., 2009). The outermost layer of the cuticle is formed by epicuticular waxes that may be deposited as an amorphous film or in the form of crystals (Jetter et al., 2006). In many species, a specific hydrophobic layer called the cuticle proper can be distinguished beneath the epicuticular wax layer (Sitte and Rennier, 1963; Schreiber and Schönherr, 2009) (Figure 1B). This layer very hydrophobic and consists mainly of the aliphatic polyester cutin, which is the main structural component, and intracuticular wax. The layer below the cuticle proper is less hydrophobic and is often referred to as the cuticular layer (Sitte and Rennier, 1963; Schreiber and Schönherr, 2009). The cuticular layer contains additional polysaccharides and forms a continuum with the cell wall, potentially with gradually decreasing proportions of lipid material towards the inner side of the cell wall (Figure 1B) (Sitte and Rennier, 1963; Schreiber and Schönherr, 2009). The cuticle has an amorphous, lamellate or reticulate structure, and these features have long served as the primary characteristics for the classification of cuticles (Holloway, 1982b). In species with a thick cuticle, the cuticular layer may form several structurally distinct layers (Jeffree, 2006). The thickness and ultrastructure of the cuticle changes markedly during the growth and development of organs, and these changes correlate with changes in cutin and wax composition (Riederer and Schonherr, 1988; Hauke and Schreiber, 1998 ).

The ultrastructure of the cuticle can be visualized by conventional transmission electron microscopy (TEM) of embedded and sectioned tissues. Soluble components, such as wax and other molecules that are not cross-linked, are removed during the embedding procedure that is necessary for TEM (Figure 1B). The surface structure of the cuticle can be visualized by cryo- or environmental scanning electron microscopy (SEM), preserving the structure of the epicuticular wax (Figure 1A).

The ultrastructure of the cuticle in rosette leaves

In wild-type (wt) Arabidopsis, the ultrastructure of the cuticle has been characterized in the leaves, inflorescence stems and flower organs. In different accessions, the cuticle of the leaf blade is documented as a thin, electron-dense layer of 20–25 nm (Xiao et al., 2007; Voisin et al., 2009) (Figure 2A, D). A layered reticulate structure was observed at high magnification (Broun et al., 2004). Mutants with a substatial reduction in cutin polyester content (> 50%), such as the long-chain acyl-CoA synthethase 2 (lacs2) (At1g49430) mutant and the glycerol-3-phosphate acyltransferase (gpat) double mutant gpat4gpat8 (At1g01610, At4g00400) and transgenic plants expressing a fungal cutinase (CUTE-plants), may not have any an osmiophilic layer on the surface of their cell walls (Sieber et al., 2000; Bessire et al., 2007; Li et al., 2007b) (Figure 2F). The lacs2 mutant exhibits an unusual gradual darkening of the cell wall, as well as osmiophilic structures within the cell wall. It has been speculated that these structures represent residual cutinization in the absence of a distinct cuticle layer on the surface of the cell wall (Bessire et al., 2007; Li et al., 2007b) (Figure 2B, 2C). Arabidopsis mutants that exhibit alterations in the composition of cutin in the leaves, such as the lacerata (lcr) (At2g45970) and bodyguard (bdg) (At1g64670) mutants, typically display visible modifications to the ultrastructure of the leaf cuticle (Kurdyukov et al., 2006a; Voisin et al., 2009). In the aberrant induction of type three genes 1 (att1) (At4g00360) mutant, the cuticle is broader and more electron-translucent than that of wt Arabidopsis (Xiao et al., 2004) (Figure 2E), while WAX INDUCER1/SHINE1 (win1/shn1) (At1g15360)-overexpressing plants accumulate electron-dense material beneath a broad electron-translucent cuticle proper (Broun et al., 2004). In a few cases, structural changes were not detected by conventional TEM methods, even when the barrier properties were altered (e.g., in the fiddlehead (fdh) (At2g26250) mutant) (Voisin et al., 2009). Epicuticular wax is deposited as a thin film on the leaf blade of wt Arabidopsis leaves. However, the overproduction of wax, as in activation-tagged WIN1/SHN1-overexpressing plants, may lead to the formation of flakes on the leaf surface (Aharoni et al., 2004; Broun et al., 2004).

Figure 2.

Cuticles of Arabidopsis leaves.

Transmission electron micrographs of the cuticle of pavement cells from Col-0 (A), lacs2–3 (B, C) (from Bessire et al., 2007), Col-gl (D), att1 (E) (from Xiao et al., 2004) and transgenic plants expressing a fungal cutinase (F) (from Sieber et al., 2000). Scale bars represent 800 nm in A–C, F and 400 nm in D–E.

The ultrastructure of the cuticle of the inflorescence stem

The inflorescence stem of wt Arabidopsis plants has a more substantial cuticle of 80 -120 nm (Sieber et al., 2000; Chen et al., 2003; Bird et al., 2007; Li et al., 2007b; Lu et al., 2009) (Figure 3A, C, E). An outer, less electron-dense cuticle proper can be distinguished from an electron-dense inner cuticular layer at the cuticle/cell wall interface close to the base of the inflorescence stem of a fully developed wt plant (between the 1st and 2nd internode) (Chen et al., 2003; Lu et al., 2009) (Figure 3A, E). In other preparations, the cuticle is visualized as a single electron-dense layer (Sieber et al., 2000; Bird et al., 2007; Li et al., 2007b; Lu et al., 2011) (Figure 3C). Mutants that have significant reductions in the amount of cutin in the stem usually maintain a distinct cuticle proper. However, the structure of the cuticle proper may vary depending on the genetic lesion. In some mutants, such as the cer3/wax2 (At5g57800), abcg11–3 (wbc11-3) (At1g17840) and gpat4/gpat8 mutants and transgenic CUTE-plants, the cuticle proper may be thicker with a loose, less osmiophilic structure (Sieber et al., 2000; Chen et al., 2003; Bird et al., 2007; Li et al., 2007b) (Figure 3B, 3D). In other mutants, such as the glossyhead1 (gsh1) (A1g36160) mutant, the cuticle proper may be a thin, strongly osmiophic layer that is more irregular (Lu et al., 2011). When a bilayered cuticle structure could be resolved in TEM preparations of wt Arabidopsis, the organization of the inner cuticular layer was strongly affected by slight decreases in cutin, while the thickness of the entire cuticle was reduced by substantial decreases in polymer content (Lu et al., 2009) (Figures 3F, 3G, 3H). In Arabidopsis, epicuticular wax of inflorescent stems is deposited in the form of crystals that reflect its amount and composition (Jetter et al., 2006). In Arabidopsis and other species, a large number of mutants have been identified by the glossy appearance of their stems, which indicates that these mutants may have reduced wax crystal deposition (Kunst and Samuels, 2003) (Figure 4).

Figure 3.

Cuticles of inflorescence stems of Arabidopsis.

Transmission electron micrographs of the cuticle of pavement cells of inflorescence stems from C24 (A), wax2 (B) (from Chen et al, 2003), Col-0 (C), abcg11–3 (D) (from Bird et al., 2007), Col-0 (E), lacs2–3 (F), lacs1 (G), lacs1lacs2 (H) (from Lü et al., 2009). Scale bars represent 100 nm in A-D and 200 nm in E–H.

Figure 4.

Epicuticular waxes on inflorescence stems of Arabidopsis thaliana.

(A) Arabidopsis influorescent stems, with decreasing wax amounts from left to right: WS, cer3, cer4, cer1, cer2 (from Mc Nevin et al., 1993). (B–C) Cryo-SEM pictures of wax crystals of Col-0. (D) Residual wax on the cer2 mutant in the form of a wax film (from Haslam et al., 2012). Scale bars (C, D): 2 µm.

The Cuticle of the Embryo and in Emerging Organs

A cuticle is formed early during embryo development. In Arabidopsis, the presence of the cuticle has been demonstrated experimentally using fluorescent staining of cuticular lipids at the globular stage, when the embryo consists of a few dozen cells (Szczuka and Szczuka, 2003). Mutant phenotypes associated with impairments in cuticle formation have been reported at the torpedo stage in the gassoh1/gassoh2 (gso1/gso2) (At4g20140/At5g44700) double mutant, indicating that at this stage of development, the diffusion barrier is usually functional (Tsuwamoto et al., 2008). Similarly, the expression of ABCG11, a transporter that is involved in cutin and wax secretion, suggests that the cuticle is present in walking stick embryos (Panikashvili et al., 2010).

The embryo is further protected by additional permeability barriers that are present in the seed coat. The seed coat contains several polyesters of different compositions that are difficult to characterize separately (Molina et al., 2006). ATT1, a cytochrome P450-dependent fatty acid ω;-hydroxylase that is encoded by CYP86A2 (At4g00360) and necessary for cutin biosynthesis, is exclusively expressed in the inner integument, while GPAT5 (At3g11430), a glycerol-3-phosphate acyltransferase necessary for suberin formation, is expressed in the outer integument (Molina et al., 2008), suggesting that these polyesters are localized in different layers of the seed coat. The seed coat will not be discussed further in this chapter.

After the cuticle has been formed during the establishment of the L1 layer, it is maintained during plant development. Thus, when new organs are formed at the shoot and floral apical meristem, they are from the beginning covered by an electron-opaque layer called the procuticle (Heide-Jorgensen, 1991). The composition of the procuticle is not known in Arabidopsis, but this layer likely has a different composition than the cuticle of mature organs, as some cutin biosynthesis mutants form organ fusions, while others do not (see below). Once the cuticle is damaged during the life of a plant, it does not regenerate; instead, suberin is formed to heal the wound.

Figure 5.

Cuticles of epidermal cells of Arabidopsis petals.

SEM (A–E) and (H–I) and TEM (F–G) and (J–K) analyses of Arabidopsis petals. On the adaxial side of the petal (A–G) and the abaxial side of the petal (H–K), epidermal cells display nanoridges on their surface. Col-0 (A, C, F, J, H), pec1 (B, D, E, G) (from Bessire et al.), gpat6 (I, K) (from Li-Beisson et al., 2009). Scale bars (A, B, H, I): 10 µm; (C–E): 2 µm; (F, G, J, K): 500 nm.

After the emergence of the seedling, the cuticle undergoes compositional and structural changes that are highly coordinated with the growth and development of the plant. In barley leaves, cutin synthesis occurs during cell elongation, while wax synthesis occurs after the cells reach their final length (Richardson et al., 2007). The exact timing of the deposition of cutin and wax has not yet been determined in Arabidopsis leaves. However, the induction of the transcription factor SHINE leads to the initial expression of genes involved in cutin biosynthesis and the subsequent induction of genes involved in wax biosynthesis 10 hours later, suggesting that cutin is formed first and then impregnated with wax (Kannangara et al., 2007). In stems, cutin and wax are present 3 mm below the meristem of elongating stem segments, indicating that cutin and wax are likely deposited in close sequence (Suh et al., 2005). The wax load remains constant at 32 µg/cm² from 1 cm below the meristem to the base of the stem. The amount of polyester that is susceptible to depolymerization is much lower in the oldest segment (i.e., 2 µg/cm²) than in the youngest segment (i.e., 8 µg/cm²), suggesting that the polymer undergoes structural changes when elongation ceases, such as increased crosslinking within itself or to other molecules of the extracellular matrix (Suh et al., 2005).

The root epidermis and its elusive cutin/suberin fortifications

Less than a millimeter from the tip of the root meristem, the last root cap cells are sloughed off, and the epidermal cells constitute the outermost cell layer of the root. The Arabidopsis root epidermis is patterned in two distinct cell files of alternating root hair and non-root hair cells (Dolan et al., 1994). Root hairs emerge quickly upon the exit of cells from the meristem and begin to perform their role in nutrient uptake. Thus, in contrast to the shoot epidermis, the epidermis of primary roots is a cell layer that is actively involved in solute acquisition from the soil. As a result, the root epidermis cannot feature a pronounced, hydrophobic cell wall modification. However, it is conceivable that the soil-facing cell walls at the base of the root hairs contain polymers that alter the epidermal cell wall properties for increased resistance to pathogens or specialized diffusive functions. Consistent with their physiological function, Arabidopsis root epidermal cell walls do not represent a significant barrier to the diffusion of solutes, especially in early developmental stages. This characteristic is demonstrated by the rapid diffusion of fluorescent tracer molecules, such as propidium iodide (PI), into inner tissue layers, which is only blocked once the dye reaches the differentiated endodermal layer (see below). Modification of the root epidermal cell wall (e.g., by the presence of suberin or lignin-like polymers) may possibly increase resistance without completely blocking the diffusion of solutes. It has been reported that dyes that detect lignin/polyaromatic substances or suberin-like components stain epidermal cell walls. This finding was observed in a number of species (Wilson and Peterson, 1983) and has been investigated in more detail in maize, onion and soybean (Peterson et al., 1978; Schreiber et al., 1999; Thomas et al., 2007). Confirmation of the presence of lignin and/or suberin in the epidermis by chemical analysis is complicated, because many species possess a highly suberized hypodermal/exodermal cell layer immediately below the epidermis, which cannot be separated from the epidermis. Chemical analysis performed in soybean (a species without a hypodermis) and ultrastructural analysis of the onion epidermis support the theory that “diffuse suberin” is present in epidermal cell walls in the root (Thomas et al., 2007). In Arabidopsis, the chemical analysis of epidermal cells would be technically challenging, and we are unaware of ultrastructural studies reporting specialized cell wall features of the root epidermal cell wall. However, increased autofluorescence and an increased resistance to PI penetration can be observed in differentiated epidermal cells (Franke et al., 2005) (Naseer et al., unpublished observation). Thus, it is possible that Arabidopsis also deposits some aliphatic and/or aromatic polymers in its root epidermal cell walls. In addition, studies of the expression patterns of cutin biosynthetic genes detected the expression of a number of relevant genes in root epidermal cells. For example, CUTICLE DESTRUCTING FACTOR 1 (CDEF1) (At4g30140) is a gene that encodes an esterase involved in the degradation of the cuticle during pollen penetration. Promoter GFP-fusions revealed a specific upregulation of CDEF1 in the root epidermis during lateral root emergence (Takahashi et al., 2010). This process requires the separation of epidermal cells and is accompanied by an induction of cell wall remodeling enzymes (Swarup et al., 2008). Thus, the expression of CDEF1 in root epidermal cells suggests that the protein encoded by this gene functions in the remodeling or degradation of standard cell wall components in this tissue. Alternatively, this finding could suggest the presence of aliphatic polymers within root epidermal cell walls (Takahashi et al., 2010). Other examples of root epidermis-expressed genes involved in cell wall polyester formation are BDG, which encodes a hydrolase involved in cuticle production, LCR, which encodes a cytochrome P450 enzyme, and ADHESION OF CALYX EDGES/HOTHEAD (ACE/HTH) (At1g72970), which encodes a protein thought to be involved in the biosynthesis of dicarboxylic acids (Wellesen et al., 2001 ; Kurdyukov et al., 2006a; Kurdyukov et al., 2006b). Interestingly, although a polyester composition typical of suberin was found in other species, the proteins that were found to be upregulated in the Arabidopsis root epidermis were previously demonstrated to be involved in cutin biosynthesis (Schreiber et al., 1999). Arabidopsis root epidermal cells are likely reinforced by aliphatic polymers, but the exact nature of these polymers remains unknown (Figure 6A).

Figure 6.

Deposition of diffusion barrier polymers in Arabidopsis roots.

Schematic diagrams of Arabidopsis roots. Cross-sections through the roots of 5-day-old Arabidopsis plants approximately 1 mm and 3 mm from the root tip (A), illustrating the endodermal differentiation states I and II, respectively. Longitudinal sections of Arabidopsis roots (B) from different growth stages, representing different types of diffusion barriers. Root hairs and lateral roots are not depicted.

A systematic study of the expression of the gene families involved in cutin and suberin biosynthesis in the root epidermis is currently feasible. Such a study could facilitate studies that clarify the presence and function of these polymers in root epidermal cell walls in Arabidopsis.

The endodermis: the Casparian strip and suberin lamellae

As mentioned above, the role of the root in nutrient uptake does not allow for the formation of highly protective, essentially impermeable barriers. Instead, a compromise between protection and interaction with the environment must be attained. To overcome this problem, many multicellular organisms have developed epithelial sheets, in which cells within a layer are tightly attached to each other through ring-like junctions that seal the extracellular space (Cereijido et al., 2004). This strategy leaves the rest of the cellular surface unprotected, but it prevents the uncontrolled entry of substances across the cell layer through the intercellular spaces. The outer and inner cellular surfaces (relative to the ring) can then engage in the selective uptake of substances from the environment. This solution developed independently in both the polarized epithelia of animals and the root endodermis of vascular plants.

In the endodermis, the sealing of the extracellular cell wall space between cells is achieved by the Casparian strip, which was named after its discoverer, the German botanist Robert Caspary (Caspary, 1865) (Figure 6). The Casparian strip is a hydrophobic impregnation of the primary cell wall that is distinct from the widespread deposition of secondary cell walls that is observed in other cells (Esau, 1977) (Figure 7A). In addition, at the Casparian strip, the plasma membrane adheres tightly to the cell wall and suppresses the lateral diffusion of plasma membrane proteins (Behrisch, 1926; Karahara and Shibaoka, 1992; Alassimone et al., 2010; Bornette and Puijalon, 2011 ). This specialized cell wall-plasma membrane (PM) junction effectively blocks the penetration of substances through the extracellular space from the cortex into the central cylinder of the root and vice versa, performing a function similar to that of tight and adherens junctions in animals. Endodermal cell walls are further impregnated by suberin in their secondary developmental stage (Krömer, 1903) (Figure 7C).

The suberized periderm

The roots of dicots, such as Arabidopsis, undergo considerable radial thickening during their lifetime, leading to the elimination of the endodermis, cortex and epidermis and the replacement of these layers by a periderm. This secondary dermal tissue develops in the pericycle, representing the first cell layer in the central cylinder of the root (Figure 6B). In Arabidopsis, the root periderm is composed of 1–2 layers of suberized cells. In their secondary developmental state, roots must facilitate the longitudinal transport of water and solutes in the xylem from the root to the shoot and the transport of sugars in the phloem from the shoot to the root. Thus, in contrast to primary roots, roots in their secondary developmental state do not significantly contribute to the radial uptake of water and dissolved nutrients. Ultrastructural investigations of the root periderm in Arabidopsis revealed the occurrence of characteristic lamellae that were previously described in suberized cells of the potato periderm and in cork (Figure 8). Typical of suberin depositions, these lamellae are formed next to the plasma membrane at the inner side of the cell wall. The localization of the diffusion barrier within the extracellular matrix is one of the features that clearly distinguish cutin from suberin depositions.

Figure 7.

Casparian strips between endodermal cells of Arabidopsis thaliana.

(A, B) Casparian strip between two endodermal cells of a 5-day-old Arabidopsis root. (C) Casparian strip in the cell wall next to the periderm of a 5-week-old Arabidopsis root. (B) Localization of Casparian strip protein 1 (CASP1) in the plasma membrane beneath the Casparian strip (CS) (from Ropollo et al. 2011). (C) Casparian strips reinforced with suberin lamellae and secondary cell wall next to the periderm (picture by Martine Schorderet, University of Fribourg). Scale bars (A, B): 250 nm; (C):100 nm. CW: Cell wall, PM: plasma membrane, CSD: Casparian strip domain.

Cuticle preparation and cutin monomer composition

The standard procedure for the determination of cutin monomer composition in different plant species is based on the isolation of the cuticle by digestion of the tissue with cellulases and pectinases (Schönherr and Riederer, 1986). In addition, solvent treatments are necessary to extract the wax components of the cuticle (see below). The chloroform-insoluble fraction contains the cutin polyester and a residual fraction that is composed primarily of polysaccharides. After extensive solvent extraction, cutin can be depolymerized by base- or acid-catalyzed procedures that cleave ester bonds. While reductive cleavage with lithium aluminum hydride is the classic method (Walton and Kolattukudy, 1972; Kolattukudy, 2001a), the cutin of Arabidopsis has been characterized by transesterification with methanol containing boron trifluoride, sodium methoxide or hydrochloric acid (Bonaventure et al., 2004; Franke et al., 2005; Li-Beisson et al., 2013). Different methods produce similar results, although acid catalysis releases more 2-hydroxy acids, which may be derived from sphingolipids rather than polyester (Franke et al., 2005). The resulting polyester monomers must be converted into derivatives (e.g., with N,O-Bis(trimethylsilyl)trifluoroacetamide (BSTFA)) before being subjected to GC-MS for identification based on their characteristic fragmentation patterns (Walton and Kolattukudy, 1972).

Figure 8.

Ultrastructure of suberized root tissues of wild type Arabidopsis plants at the beginning of the secondary thickening of the root.

(A) Overview of suberized endodermal and peridermal cells in the root. Suberin deposition is visible as electron-opaque layer inside the primary cell wall. The fully suberized peridermal cell layer typically collapses during the dehydration and embedding procedures necessary for TEM because of the low permeability of the suberized cell walls. Scale bar: 2.5 µm. (B) Enlargement of (A). Fine structure of suberin. The structure of the lamellae, with an alternation of electron-opaque and electron-translucent suberin layers, is clearly visible when the specimen is cut perpendicularly to the suberin layers (concave arrowheads). However, the lamellate structure of suberin is barely visible when the specimen is not cut perpendicularly to the suberin layers (arrow). Scale bar: 500 nm. (C) Enlargement of (B). The thickness of the electron-opaque and electron-translucent suberin layers is very regular and characteristic for the tissue sample (pictures by Martine Schorderet, University of Fribourg). Scale bar: 100 nm. P: peridermal cell, E: endodermal cell: pericycle cell, CW: cell wall.

Because the cuticle of Arabidopsis leaves is very thin, the polyester composition of isolated cuticles has only been reported once (Franke et al., 2005). A comparison of the polyester composition of isolated cuticles with the aliphatic monomers released by the transesterification of total leaf material that has been extensively extracted with methanol-chloroform revealed qualitatively identical composition. The analysis of exclusively solvent-extracted plant material has the disadvantage of detecting monomers that include material that is solubilized from the cell walls of internal tissues. However, this procedure also has several advantages and is commonly used. The composition of the polyester can be obtained from nearly any tissue, even from mutants with reduced or fragmented cuticles. In addition, material at the interface between the cell wall and the cuticle is not lost during preparation.

Typical cutin monomers are hydroxy and epoxy fatty acids with a backbone of 16 or 18 carbons, such as 10(9),16-dihydroxyhexadecanoic acid, 9,10,18-trihydroxyoctadecanoic acid and 9,10-epoxy-18-hydroxyoctadecanoic acid. Minor monomers may also be fatty acids (FA), fatty alcohols, fatty acids that contain aldehyde and carbonyl groups, fatty diacids and hydroxycinnamic acids. In Arabidopsis, the generic compositional predominance of C16 and C18 in-chain hydroxylated FA derivatives applies only to floral cutin (Li-Beisson et al., 2013). The leaf and stem cutin is remarkably different, as it consists of more than 40% of unsaturated α,ω;-dicarboxylic acids (DCAs) (Bonaventure et al., 2004; Franke et al., 2005; Li-Beisson et al., 2013), which were previously reported as minor cutin monomers (Holloway, 1982a). Whether these compositional differences reflect differences in structure and functional properties remains to be determined. A comparison of the cutin load and composition in different organs, including rosette leaves, stems and flowers, is provided in Table 13 of the TAB chapter on acyl-lipid metabolism (Li-Beisson et al., 2013). Mutants that lack hydroxylated fatty acids in the cell wall-residue have not yet been identified, indicating that the incorporation of hydroxylated fatty acids in the epidermal extracellular matrix is necessary for the survival of the plant.

Cuticular wax extraction and composition

Suberin preparation and suberin monomer composition

Similar to cutin, the preferred sample preparation method for the analysis of the chemical composition of suberized cell walls of the root includes incubation with polysaccharide hydrolases and manual separation of suberin-containing cell walls, which are resistant to cellulases and pectinases, followed by a solvent extraction for the removal of unbound components. Suberized periderms, such as potato tuber peels, are well suited for this approach, whereas suberized root tissues, such as the mostly monolayered exodermis, have only been isolated from certain species (Schreiber et al., 2005a, 2007). In Arabidopsis, the isolation and separation of suberized tissues from roots has not yet been achieved, and hydrolase digestion and/or extraction of whole root sections is a commonly used method (Franke et al., 2005; Li-Beisson et al., 2013).

Standard depolymerization methods (i.e., base- or acid-catalyzed transmethylation) of suberin-enriched Arabidopsis root cell wall material revealed a monomer composition similar to that of potato suberin, the long-time suberin model. Bifunctional C18 monounsaturated ω;-hydroxyacids and α,ω;-DCA are the major monomers, followed by saturated ω;-hydroxyacids and α,ω;-DCAs that vary in chain-length from C16 to C24 (Franke et al., 2005). Monofunctional C18 to C24 FA and alcohols comprise 15% of the Arabidopsis root suberin polyester.

Suberin-rich roots and seeds can also be extracted without prior enzymatic digestion (Molina et al., 2006; Li-Beisson et al., 2013). Typically, in an optimized chromatography system for long-chain aliphatics, contamination with a small amount of carbohydrates does not cause significant interference. A systematic evaluation of the advantages, disadvantages and yields of the different base- and acid-catalyzed depolymerization methods for Arabidopsis root suberin has not yet been reported. However, suberin profiles generated using different methods are qualitatively similar (i.e., in the composition and ratio of the aliphatic monomers). Some quantitative differences that have been reported in the literature, even within a single publication, can be attributed to seasonal, developmental and cultivation variations. It is reasonable to assume that, like cutin, no significant differences in suberin composition will be observed between base- and acid-catalyzed transmethylation (Franke et al., 2005 suppl.). An overview of suberin monomer load and composition are presented in Tables 9 and 10 of the TAB chapter acyl-lipid metabolism (Li-Beisson et al., 2013).

Identification and characterization of root waxes

While cuticles, which cover leaf and fruit surfaces, always occur together with wax that efficiently seals the cutin polymer, suberin does not necessarily contain wax molecules (Schreiber, 2010). In suberized endodermal and hypodermal cells isolated from the roots of a number of different species, typical wax compounds are absent or present in trace amounts. Conversely, in suberized tissues directly exposed to the atmosphere (e.g., potato tubers, aerial roots, etc.), which experience a lower water potential than tissues in the soil, wax amounts and wax compositions similar to those observed in cuticles have been described. Like the cuticle, wax also efficiently seals the periderm, and wax extraction from the potato periderm increases the peridermal permeability by approximately 2 orders of magnitude (Schreiber et al., 2005b).

Extraction methods similar to the methods employed for the collection of total cuticular waxes have recently been applied to whole roots of fully developed Arabidopsis plants. In addition to sterols, free fatty acids, primary alcohols, secondary alcohols and alkanes that resemble aerial waxes, the soluble lipids in these extracts also included monoacylglycerols and alkyl hydroxycinnamates (Li et al., 2007a; Molina et al., 2009). These lipids have been termed “root waxes” and are located in the tap root (Kosma et al., 2012). The taproot wax is particularly rich in alkyl hydroxycinnamates, which comprise approximately 50% of the extracted lipids, including esters of C18-C22 alcohols with the hydroxycinnamic acids p-coumarate, caffeate and ferulate (Kosma et al., 2012). The presence of C18-C22 alkyl hydroxycinnamates on root surfaces was also confirmed using laser desorption-ionization mass spectrometry (Jun et al., 2010). Alkyl hydroxycinnamates in potato tuber periderms (e.g., C16-C32 ferulate esters) contribute substantially to the barrier properties of this suberized tissue (Schreiber et al., 2005b). The roles of these waxy compounds in diffusion barrier formation and other physiological functions in Arabidopsis remain to be determined. This complex mixture of waxy components may possibly also be the result of pooling compounds that originate from the apoplast of different cell types, such as the epidermis, endodermis and periderm; thus, these components may mediate multiple physiological functions. Cell type-specific expression analysis and the localization of biosynthetic enzymes in different cell types may provide answers to these questions (Kosma et al., 2012).

The composition of the Casparian strip

Despite extensive research, considerable uncertainty as to the chemical nature of this localized wall impregnation remains. Caspary himself was unable to conclude whether the Casparian strips are composed of “Holzstoff” (i.e., wood substance, or lignin) or “Korkstoff” (i.e., cork substance, or suberin). Krömer later concluded that the Casparian strip shows a typical “wood staining reaction” and found no convincing evidence for the presence of suberin (Krömer, 1903). Despite this finding, suberin was often proposed to be the major constituent of Casparian strips. In 1961, Vanfleet discussed these divergent views and concluded that although none of the studies “satisfy the question, lignin or suberin, they do suggest the presence of phenolic material” (Vanfleet, 1961). However, Vanfleet also cites studies that suggest that the “Casparian strip is probably made of lignin plus fatty acid.” Regardless, the textbook ”Anatomy of Seed Plants” by Esau describes the Casparian strip as being composed of suberin and does not mention lignin (Esau, 1977). One major obstacle that must be overcome when investigating this issue is that after the establishment of the Casparian strip, Arabidopsis endodermal cells rapidly form suberin lamellae on all cell surfaces, making it difficult to exclusively analyze Casparian strips. Pure samples of Casparian strips are difficult to obtain in sufficient amounts for chemical analysis. In addition, a survey by Wilson and Peterson that used a number of histochemical stains on more than 30 plant species suggested that there is a considerable degree of natural variation in the chemical composition of the Casparian strip (Wlson and Peterson, 1983).

Schreiber et al. used natural variations in endodermal development to analyze the endodermis in its primary developmental state, which is characterized by the sole presence of Casparian strips, in three species: Clivia miniata, Pisum sativum and Monstera deliciosa (Schreiber, 1996). This study demonstrated that lignin is the major polymer in the Casparian strips of these species. However, suberin was also detected in small amounts. Histochemical investigations of Casparian strips in a variety of species typically reveal the presence of both lignin (via staining with phloroglucinol/HCl) and suberin (via staining with Sudan). The lignin monomer ratios observed in the endodermal cell wall are similar to the lignin of xylem vessels, but a specialized form of Casparian strip lignin, which may represent an adaptation to the function of the Casparian strip as a solute diffusion barrier, cannot be excluded (Naseer et al., 2012). In another study, maize roots were dissected at different developmental stages (Zeier et al., 1999). The first stage, which included only Casparian strips, was observed to contain a distinct lignin component and smaller amounts of suberin (Schreiber et al., 1999). Recent developmental studies in Arabidopsis, with a higher spatial resolution along the length of the developing root, indicate that lignification of the primary cell wall occurs in parallel with the tight attachment of the plasma membrane to the cell wall site of the Casparian strip, approximately 12 cells after the onset of elongation. In contrast, suberization appears to occur much later, when functional Casparian strips have already been formed. Histochemically detectable suberin appears approximately 38 cells after the onset of elongation (Naseer et al., 2012). Importantly, this study employed genetic and pharmacological manipulations, which demonstrated that interference with suberin deposition has no observable effect on the development of functional Casparian strips. Conversely, blocking lignin biosynthesis using an inhibitor of the phenylpropanoid pathway abrogates the formation of Casparian strips and leads to a completely permeable endodermis. In addition, monolignol feeding experiments and chemical analysis of root tips that contain only Casparian strips as potentially lignified cell layers both indicated that the initial Casparian strips of Arabidopsis are composed of a polymer closely related to the “canonical” lignin of xylem vessels. These findings strongly suggest that in Arabidopsis, suberin is confined to the formation of suberin lamellae during the secondary developmental state of endodermal cells (Figure 6A, Figure 7A, C). Currently, the spatial resolution of sampling methods for the analysis of root cell walls does not allow for the chemical determination of whether suberin is added to older Casparian strips at the time of suberin lamellae formation.To answer this question, Casparian strip cell walls and the cell walls from the inner and outer tangential walls of the endodermis would have to be isolated in sufficient amounts for quantitative compositional analysis; such an experiment is not currently realistic in Arabidopsis.

In the past, suberin has been described as a polymer that not only contains ferulate esterified to the aliphatic polymer but also contains a phenylpropanoid-derived polyaromatic domain. The definition of the polyaromatic domain in suberin is specifically based on studies with the potato wound periderm (Bernards, 2002). Recent studies used an inhibitor of the phenylpropanoid pathway in Arabidopsis that blocks both monolignol formation and the formation of other monomers needed for the synthesis of the aromatic polymer of suberized tissues (Naseer et al., 2012). As mentioned above, the application of this drug to Arabidopsis seedling roots led to a complete block of lignin formation without affecting the formation of (aliphatic) suberin in distal endodermal cell walls. This finding demonstrated that in Arabidopsis, the aliphatic polyester (suberin) could form independently of an aromatic polymer. The finding that suberin can exist independently of the presence of a polyaromatic domain raises the question of whether suberin should be in general viewed as a polymer with an aliphatic and a polyaromatic domain or a polymer containing only an aliphatic domain with esterified aromatic compounds. The data described above clearly demonstrate that the Casparian strip of 5-day-old Arabidopsis seedlings is a functional diffusion barrier that is composed predominantly of a lignin polymer. This finding is also supported by the recent identification of cell wall biosynthetic genes involved in Casparian strip formation (Lee et al., 2013) (see below). Additional model systems must be investigated to answer important questions concerning the general structure of suberin and the importance of lignin for the formation of the Casparian strip in other species. It will also be important to obtain better read-outs for barrier functionality. The use of fluorescent tracers, such as PI, is very convenient. However, large, charged molecules might behave differently from relevant monovalent or divalent ions and polar water molecules. It seems reasonable to assume that a defect that allows for the penetration of PI would also allow for the penetration of elemental nutrients and water. The reverse conclusion, however, does not apply. Many mutants with subtle defects that do not exhibit penetration of PI may display significant penetration of other elements or water.

Core reactions common to cutin and suberin monomer biosynthesis

Based on the predominant classes of compounds detected in other plant species, namely ω;-hydroxy acids, α,ω;-DCA, alcohols, carboxylic acids and fatty acids, and the presence of glycerol in both cutin and suberin, it is reasonable to assume that the core reactions in cutin and suberin biosynthesis are catalyzed by the same or similar enzymes. The identification of candidate genes was facilitated by epidermal and endodermal transcriptomics (Birnbaum et al., 2003; Suh et al., 2005; Brady et al., 2007), which allowed for the identification of transcripts whose predicted functions aligned with the proposed metabolic activities deduced from the known chemical composition of these polyesters. Four biosynthetic steps are common to the biosynthetic pathways of these polyesters, namely the oxygenation, reduction and activation of fatty acids and their subsequent transfer to glycerol-3-phosphate.

Characteristic reactions in the biosynthesis of aliphatic cutin monomers

Characteristic reactions in the biosynthesis of aliphatic suberin monomers

The major chemical differences between cutin and suberin aliphatic polyesters are: i) the abundance of very long-chain fatty acid (VLCFA) derivatives and ii) the presence of up to 5% aromatic components, mainly ferulic acid, in Arabidopsis suberin.

Open questions regarding the biosynthesis of polyester intermediates

Many key enzymes involved in cutin and suberin monomer biosynthesis have been identified through the characterization of Arabidopsis mutants. However, inferring enzymatic activity from chemical phenotypes of the polymer is difficult, as plants are complex organisms that often compensate for the loss of a gene function by activating other pathways, and biochemical verification is only available for a subset of proteins. The organization of individual reactions into pathways, the in vivo substrates of the required enzymes, and the relevant metabolic intermediates must be elucidated. For example, it is not known whether FAE precedes ω;-hydroxylation. It also remains to be determined at which steps of FA modification CoA-activation and acylglycerol formation take place (Beisson et al., 2012).

The compositional analysis of cutin in cyp86a4 and cyp77a6 mutants and the finding that recombinant CYP77A6 hydroxylates 16-hydroxyhexadecanoic acids indicate that in flower cutin biosynthesis, ω;-hydroxylation is required for subsequent in-chain hydroxylation. In contrast, yeast-expressed CYP86B1 did not generate any hydroxylated products from the provided FAs. To determine whether conjugated FA, such as acyl-CoAs and glycerolipids, which are potentially generated by LACS or GPAT, and not free FA are the endogenous substrates further study is required. For CYP86A22 from Petunia sp., it has been demonstrated that saturated and unsaturated acyl-CoA derivatives are the substrates for ω;-hydroxylation (Han et al., 2010). A strong partnership of GPAT with CYP86 has been demonstrated by co-expression of the suberin-associated genes GPAT5 and CYP86A1 (Li et al., 2007a; Li-Beisson et al., 2009). Co-expressing lines incorporated C20 and C22 ω;-hydroxyacids and α,ω;-DCAs, which are typically present in root polyesters, into shoot polyesters (Li et al., 2007a). These findings and other metabolic evidence concerning the sequential order of reactions has been previously discussed (Yephremov and Schreiber, 2005; Li-Beisson et al., 2013; Beisson et al., 2012).

Many open questions concern the mechanisms by which the biosynthesis of cuticular lipids interacts with general acyl-lipid metabolism, as a number of mutants affected in general lipid biosynthesis have cuticular phenotypes, including fatty acid desaturase2 (fad2) (At3g12120), fatty acyl-ACP thioesterase b (fatb) (At1g08510), acyl carrier protein4 (acp4) (At4g25050), acyl CoA binding protein 3 (acbp3) (At4g24230), acbp4 (At3g05420), and acbp6 (At1g31812) (Bonaventure et al., 2004; Xia et al., 2009; Xia et al., 2012).

The biosynthesis of VLCFAs

FAE complexes are heterotetramers of monofunctional proteins that catalyze a series of four reactions to elongate plastid-generated C16 or C18 fatty acids by the sequential addition of two carbons from malonyl-CoA. The condensing enzyme, or β-ketoacylCoA synthase (KCS), catalyzes the first of the four reactions and is both rate-limiting and specific for the chain length of acyl-CoA synthesized (Millar and Kunst, 1997). Two highly divergent families of KCSs have been identified in Arabidopsis: an FAE1-type family homologous to the first condensing enzyme demonstrated to be involved in seed oil biosynthesis (Kunst et al., 1992; James et al., 1995), and ELOs, which have homology to the Saccharomyces cerevisiae ELO family that is responsible for sphingolipid synthesis (Dunn et al., 2004). Of the 21 FAE1-type KCS enzymes in Arabidopsis (Joubes et al., 2008), 11 have been shown by microarray analysis to be upregulated in the stem epidermis (Suh et al., 2005). Only one of these proteins, ECERIFERUM 6/3-KETOACYL-COA SYNTHETASE 6/CUTICULAR WAX 1, which is encoded by CER6/KCS6/CUT1 (At1g68530) (Millar et al., 1999; Fiebig et al., 2000; Joubes et al., 2008), has a dominant role in the elongation of VLCFAs for cuticular wax synthesis, as CER6 suppression causes a dramatic reduction in wax monomers longer than C24 and a 9095% depletion of cuticular wax on Arabidopsis inflorescence stems (Millar et al., 1999). The only Arabidopsis ELO protein functionally characterized to date is HIGH EXPRESSION OF OSMOTICALLY RESPONSIVE GENES 3, encoded by HOS3 (At4g36830), which was shown to be involved in the synthesis of VLCFAs ranging from C22 to C26 in length. Disruption of VLCFA production in the hos3 mutant causes ceramide deficiency and affects ABA-related stress responses, such as root growth inhibition, seed germination with ABA treatment and water loss from leaves. These results implicate HOS3 and VLCFA pathway products in several aspects of ABA-mediated stress signaling (Quist et al., 2009).

Recently, an additional protein has been demonstrated to be involved in the elongation of VLCFAs to 30 carbons, which is critical for the production of most wax constituents. Functional characterization of KCS enzymes from Arabidopsis in yeast revealed that the CER6 enzyme, the key KCS associated with cuticular wax synthesis, is unable to elongate VLCFAs beyond C28. Mutant screens have not identified other condensing enzymes involved in VLCFA elongation beyond C28. The only known mutant defective in the conversion of C28 to C30 VLCFAs is eceriferum2 (cer2) (At4g24510), indicating that CER2, annotated as a BAHD acyltransferase based on sequence homology, is required for C28 elongation. Biochemical analysis of CER2 demonstrated that CER2 does not share the catalytic mechanism that has been confirmed for other members of the BAHD family and that concerted activity of CER2 and CER6 in Saccharomyces cerevisiae is necessary and sufficient for C30 VLCFA synthesis (Haslam et al., 2012). Recent evidence also indicated that a CER2-like gene called CER26 encodes an enzyme involved in the elongation of C30 to C32 for leaf cuticular wax production (Pascal et al., 2013).

The synthesis of different wax components from VLCFAs

All aliphatic wax components are generated from VLCFAs by two biosynthetic pathways: the alcohol-forming acyl reduction pathway, and the alkane-forming decarbonylation pathway. The enzymes and reactions of the acyl reduction pathway that generate primary alcohols and alkyl esters have been well characterized in Arabidopsis (Samuels et al., 2008). The alkane formation pathway is more obscure, and it has only recently been demonstrated that co-expression of two well known waxrelated Arabidopsis proteins, ECERIFERUM 1, encoded by CER1 (At1g02205), and ECERIFERUM 3, encoded by CER3 (At5g57800) (Li-Beisson et al., 2013), in yeast results in the production of Arabidopsis-characteristic alkanes (Bernard et al., 2012). Although the exact biochemical role of these proteins in alkane formation remains unresolved, the CER1 and CER3 proteins were shown to physically interact in the endoplasmic reticulum in both yeast and Arabidopsis, suggesting that they associate in a complex. Both CER1 and CER3 contain tripartite His clusters, which are essential for the catalytic activity of hydroxylase and desaturase enzymes. Site-directed mutagenesis of these His-rich motifs revealed that all three His clusters were required for CER1 activity but not CER3 activity. CER1 was also demonstrated to interact with endoplasmic reticulum-localized isoforms of cytochrome b5 (CYTB5), a proposed electron donor to nonheme irons associated with His clusters. Co-expression of CYTB5 with CER1 and CER3 in yeast resulted in a two-fold increase in alkane yield, suggesting that CYTB5s are CER1 electron donors and that alkane synthesis is a redox-dependent process (Bernard et al., 2012). Alkanes are further metabolized in the Arabidopsis stem epidermis to yield secondary alcohols and ketones. A midchain alkane hydroxylase, which is encoded by MIDCHAIN ALKANE HYDROXYLASE1 (MAH1) (At1g57750), catalyzes the conversion of odd-chain alkanes to secondary alcohols and is likely to catalyze the conversion of secondary alcohols to ketones (Greer et al., 2007).

Transport of aliphatic compounds across the plasma membrane

One mechanism for the delivery of wax and polyester components to the PM involves a classical secretory pathway in which secretory vesicles carry the biosynthetic intermediates and export them into the cell wall by exocytosis (Franke and Schreiber, 2007; Pollard et al., 2008). In addition, direct transfer of lipids from the ER to the PM has been hypothesized (Kunst and Samuels, 2003). Interestingly, some of the enzymes of the BAHD family, such as DCR and DCF, which are involved in the formation of cutin intermediates, have been localized to the cytoplasm, which necessitates the presence of their substrates in this compartment or at the PM/cytoplasm interface (Panikashvili et al., 2009; Rautengarten et al., 2012). Thus, additional transport mechanisms may be involved (Panikashvili et al., 2009).

While the wax molecules that must be transported are relatively well characterized, the nature of the exported molecules for polyester formation (i.e., monomeric acyl lipids, esterified lipids, such as acylglycerol and feruloyl conjugates, or intracellular preformed oligomers) remains largely unknown. However, there is increasing evidence that free fatty acids and sn-2 monoacylgylcerols are exported (Li et al., 2007b). In GPAT5-overexpressing lines, new free fatty acids and suberin-specific monoacylglycerols appear in cuticular waxes and are incorporated into cutin when CYP86A1 is co-expressed with GPAT5 (Li et al., 2007a). Because it is unlikely that monoacylglycerols are the only precursors of polyester monomers, other intermediates are likely synthesized by additional acyltransferases, some of which may have broader substrate specificities and may also be involved in general acyl lipid metabolism.

Monolignol export

Several lines of evidence have demonstrated that the full-size ABC-transporter ABCG29 of Arabidopsis, which belongs to the PDR family (At3g16340), can act as a p-coumaryl alcohol transporter in yeast and in planta (Alejandro et al., 2012). abcg29 mutant lines have reduced levels of p-hydroxyphenyl and guaiacyl monomers and a slight reduction of synringyl monomers in the lignin, supporting this hypothesis. Although ABCG29 was expressed in endodermal cells at the plasma membrane, it was excluded from the Casparian strip, consistent with the theory that localized export of monolignols is not necessary for Casparian strip formation (Alejandro et al., 2012) (see below).

The regulation of cutin biosynthesis

The first transcription factors shown to regulate cuticle formation were WIN1/SHN1 and its homologs (Aharoni et al., 2004; Broun et al., 2004). These transcription factors primarily control cutin biosynthesis, as demonstrated by binding of the WIN1/SHN1 transcription factor to the promoter of the LACS2 gene, and indirectly affect wax accumulation (Kannangara et al., 2007) (Figure 9). The WIN/SHN family of transcription factors, encoded by SHN1 (At1g15360), SHN2 (At5g11190) and SHN3 (At5g25390), also regulates the expression of a number of genes involved in the formation of cell wall polysaccharides and proteins of the epidermal extracellular matrix in Arabidopsis flowers (Shi et al., 2011). Gene expression studies revealed that the WIN/SHN transcription factors are regulated by gibberellic acid (GA) via DELLA proteins (Shi et al., 2011). In addition, cuticle formation is integrated into the network of epidermal cell differentiation by TRANSPARENT TESTA GLABRA1 (GL1 ), which is encoded by TTG1 (At5g24520), and GLABRA3, which is encoded by GL3 (At5g41315). These transcription factors are key players in the network of proteins involved in trichome differentiation. The mutants gl1, ttg1 and gl3 have permeable cuticles that are likely due to a reduction in cutin amount, as determined for gl1 (Xia et al., 2010). The observed deficiencies in cuticle formation in the gl1 mutant were restored by the application of GA, supporting the notion that these factors act early in the cutin biosynthetic pathway (Shi et al., 2011) (Figure 9). In contrast, the lack of trichomes in these mutants was not restored by the application of GA, demonstrating an absolute requirement for GL1, TTG1 and GL3 during trichome differentiation.

Wu et al. (2011) reported the isolation of the CURLY FLAG LEAF 1 (CFL1) gene (At2g33510) and demonstrated that it encodes a WW domain protein involved in cuticle development in Arabidopsis and rice (Wu et al., 2011). This study provided biochemical evidence that AtCFL1 interacts with HOMEODOMAIN GLABROUS1 (HDG1), encoded by HDG1 (At3g61150), a class IV homeodomain-leucine zipper transcription factor that regulates the cuticle development-related genes BDG and FDH (Figure 9).

Figure 9.

Schematic diagram of the regulatory circuits involved in cuticle formation.

Cuticle formation is regulated during organ development and by environmental factors. The hormones GA and ABA (in rouge) are involved in this regulation. Regulatory elements that have been identified are shown. Genes encoding biosynthetic enzymes are also presented (red boxes). Postulated regulatory pathways are indicated by dashed lines.

Thus, the regulation of the deposition of cutin polyester is part of the transcriptional network involved in epidermal cell differentiation.

The regulation of wax biosynthesis

To date, no transcription factors have been demonstrated to be involved in the direct activation of wax biosynthetic genes during cuticle development. Instead, characterization of the waxdeficient cer7 mutant revealed that wax production in Arabidopsis stems is controlled by the CER7 ribonuclease, which is encoded by CER7 (At3g60500), a core subunit of the exosome that is responsible for the 3′ to 5′ degradation of RNA (Hooker et al., 2007) (Figure 9). Functional characterization of the CER7 enzyme demonstrated that it positively regulates mRNA levels of CER3, a wax biosynthetic gene whose protein product is required for wax formation via the alkane pathway (Hooker et al., 2007; Rowland et al., 2007). Because CER7 is a ribonuclease, it likely acts indirectly by degrading a repressor of CER3 transcription. To identify the putative repressor, a genetic screen for mutations that suppress the stem wax deficiency of cer7 was performed; a series of wax restorer (war) mutants were identified, which had mutations in genes distinct from CER7. Map-based cloning of two of the WAR genes identified them as RDR1 (At1g14790), which encodes RNA-DEPENDENT RNA POLYMERASE 1, and SGS3 (At5g23570), which encodes SUPPRESSOR OF GENE SILENCING 3. Both of these proteins are involved in the synthesis of small RNAs. These results led to the hypothesis that uncharacterized small RNA species directly or indirectly mediate posttranscriptional silencing of CER3 to control cuticular wax deposition in developing Arabidopsis inflorescence stems (Lam et al., 2012).

The regulation of polyester biosynthesis

Genes involved in both cutin and suberin formation are induced by environmental stimuli. For example, localized suberization is modified in response to drought, salinity or wounding. Consistent with these observations, the suberin biosynthetic genes are transcriptionally activated by the same environmental factors (Franke et al., 2009; Lee et al., 2009; Domergue et al., 2010). The BDG, LACS2 and PEC1 genes that are involved in cutin deposition are also transcriptionally upregulated by drought and abscisic acid (ABA) treatment (van den Brule and Smart, 2002; L'Haridon et al., 2011). Interestingly, cutin accumulation is only induced by drought stress and ABA, while wax deposition is also affected by water deficit and mild salt treatments, suggesting that the regulation of wax production may be more complex than the control of cutin biosynthesis (Kosma et al., 2009). The ABA biosynthetic mutants aba2 and aba3 have permeable cuticles, similar to those of the bdg and lacs2 mutants, underscoring the importance of ABA for cutin biosynthesis (L'Haridon et al., 2011) (Figure 9). A functional cuticle is also required for osmotic stress-mediated regulation of ABA biosynthesis by a mechanism that is impaired in a mutant with a lesion in 9-CIS EPOXYCAROTENOID DIOXYGENASE DEFICIENT1 (CED1/BDG) (At2g38110) (Kurdyukov et al., 2006a; Wang et al., 2011b) and in several other cutin biosynthesis mutants, including att1, lacs2 and lcr, but not in the wax-deficient mutants mahl and cer7 (Wang et al., 2011b). Surprisingly, this feedback regulation may be independent of cuticular permeability. Additional studies are required to elucidate the mechanism underlying this phenotype (Wang et al., 2011b).

The transcription factor MYB41, encoded by MYB41 (At4g28110), is specifically involved in the negative regulation of cutin biosynthesis, as indicated by reduced expression of LACS2 and ATT1 in MYB41-overexpressing lines (Cominelli et al., 2008). These transgenic lines show typical phenotypes associated with a permeable cuticle, including sensitivity to desiccation and abnormal cell expansion. MYB41 has been shown to be involved in general responses to osmotic stress (Lippold et al., 2009) (Figure 9).

The regulation of wax biosynthesis

Two MYB domain-containing transcription factors have been demonstrated to control cuticular wax biosynthesis in response to environmental stimuli. The MYB96 transcription factor, which is encoded by MYB96 (At5g62470), was demonstrated to promote cuticular wax accumulation under drought conditions by binding directly to conserved sequences in the promoters of wax biosynthetic genes and subsequently activating their transcription (Seo et al., 2011). MYB30, which is encoded by MYB30 (At3g28910), was demonstrated to activate the expression of wax biosynthetic genes in response to attacks by pathogens, but the extent to which this transcription factor participates in wax biosynthesis under normal conditions remains to be determined (Raffaele et al., 2008) (Figure 9).

A complex regulatory network is involved in cuticle formation, and the well-coordinated activity of all involved proteins is required for the generation of a functional diffusion barrier. Although some plants, including the bdg mutant and WIN/SHN-overexpressing plants, have a larger amount of cutin, this feature does not result in a better-sealed cuticle. In contrast, these plants have disorganized and more permeable cuticles (Aharoni et al., 2004; Broun et al., 2004; Kurdyukov et al., 2006a; Kannangara et al., 2007). In this context, the function of the CER9 protein, which is encoded by CER9 (At4g34100), a homologue of the E3 ubiquitin ligase DoA10 in yeast, which was previously demonstrated to target ER proteins for proteasomal degradation, is of great interest. Loss of CER9 activity leads to coordinated alterations in cutin and wax biosynthesis and improved sealing properties of the cuticle (Lu et al., 2012).

Diffusion processes across the cuticle

The plant cuticle acts as an efficient barrier that protects the shoot from uncontrolled water loss. Gas exchange and transpiration are primarily regulated by stomata. When stomata are closed, residual water loss from the plant surface occurs by cuticular transpiration and non-functional stomata (Kerstiens, 1996). To exclusively study cuticular transpiration, many researchers investigating cuticular water permeability have used enzymatically isolated astomatous cuticles as their experimental system. This approach allows for the quantitative determination of cuticular barrier properties, which are described by the permeance, P, in units of m s-¹. P is thus a measure of velocity, which describes the speed at which water diffuses across the cuticular membrane (Schreiber and Schönherr, 2009). On the basis of the P value of a barrier, cuticular transpiration can be directly compared with the water permeability of other biological and technical membranes (Riederer and Schreiber, 2001). In addition, P can also be measured for solutes dissolved in water (e.g., herbicides and sugars), which allows for the correlation of solute permeability with water permeability.

Quantitative assessment of cuticle permeability

Although quantitative characterization of cuticular barrier properties has been performed for a variety of species, this characterization has not been performed for Arabidopsis. Direct application of these well-established experimental approaches to Arabidopsis was not possible. Even if cuticle isolation were feasible, which is not the case, as the isolated cuticle is extremely fragile and easily disintegrates (Franke et al., 2005), Arabidopsis has stomata on both sides of the leaf, and astomatous cuticles are not available. However, because this problem also applies to the cuticles of many other species, approaches have been developed to allow for the quantitative characterization of the cuticular permeability of intact stomatous leaf surfaces. This characterization was possible because (i) it could be demonstrated experimentally that the permeability of isolated cuticles and intact leaves was similar (Kirsch et al., 1997), and (ii) the cuticular permeability of solutes (e.g., organic solutes and herbicides) was highly correlated with water permeability (Niederl et al., 1998). Thus, the cuticular transpiration of stomatous leaf surfaces can be predicted from the measured cuticular permeability of non-volatile, radiolabeled solutes dissolved in water. If radiolabeled water were used instead of radiolabeled solutes with these stomatous leaf surfaces, there is a possibility that a substantial fraction of the water that entered the leaf via open stomata could not be excluded, resulting in an erroneous determination of cuticular transpiration.

This approach has recently been successfully applied to Arabidopsis leaves (Ballmann et al., 2011). Experiments using isolated cuticles and astomatous intact leaf surfaces from a number of species demonstrated that the cuticular permeability of ¹⁴C-epoxiconazole was highly correlated with ³H₂O permeability. This correlation was used to predict the cuticular water permeability of intact Arabidopsis leaves from measurements of ¹⁴C-epoxiconazole permeability (Ballmann et al., 2011). The permeance of the Arabidopsis leaf cuticle was 4.55 x 10^-8 m·s^-1. This value can be directly compared to the permeances established for cuticles isolated from other species (Schreiber and Schönherr, 2009). The permeability of the Arabidopsis cuticle is one to two orders of magnitude higher than that of most other cuticles. Thus, it is in the upper range of permeances. However, previous permeance measurements were primarily obtained using cuticles isolated from evergreen species that had a thickness of approximately 1–3 µm (Schreiber, 2010). Considering that the Arabidopsis leaf cuticle is approximately 100-times thinner, the high permeance is not surprising. In addition, it has been speculated that the unusual chemical composition of the Arabidopsis cuticle, which is dominated by unsaturated monomers, contributes to its relatively high permeance.

Previous studies demonstrated that aliphatic wax molecules improve the sealing of the cuticle, while the role of other wax components was uncertain (Schreiber and Schönherr, 2009). The effect of triterpenoids on the sealing properties of the Arabidopsis cuticle has recently been addressed by overexpression of LUPEOL SYNTHASE 4 (LUP4) (At1g7895) in Arabidopsis leaves that do not naturally contain β-amyrin (Buschhaus and Jetter, 2012). The increased permeability of the cuticle in the transgenic lines, which accumulated up to 4% β-amyrin in their intracuticular wax layer, suggests that triterpenoids negatively affect the sealing of the cuticle. Alternatively, the unnatural β-amyrin accumulation could also disturb the cutin/wax association in the intracuticular wax layer. Thus, a general conclusion regarding the effect of triterpenoids on cuticle permeability cannot be drawn at this time.

Semi-quantitative assessment of cuticle permeability

Although the approach described above allows for quantitative comparisons of the cuticular barrier properties of Arabidopsis wt plants to mutants, it is not convenient for fast and efficient screening of cuticle mutants because it is time consuming and requires radioactivity. Water loss assays, drought susceptibility tests using intact plants or excised leaves, and assays of dye diffusion through the cuticle have been established as alternatives (Tanaka et al., 2004; Kurdyukov et al., 2006a; Bessire et al., 2007). Toluidine blue (TB) is a dye that binds to anions, such as those present in pectin, while calcofluor white binds to polymeric α- or β-glucans of the cell wall (i.e., callose and cellulose, respectively). Thus, for these water-soluble dyes to stain the leaf, they must be able to cross the lipophilic cuticular barrier and bind to the polar cell wall. The uptake of TB can also be determined semi-quantitatively in relation to the chlorophyll content of the plant using a spectrometric assay (Xing et al., 2013). Similarly, herbicide penetration assays can be used to characterize mutants with an altered/defective cuticle, with lower concentrations of herbicide causing tissue damage in the mutants but not the wt (Bessire et al., 2007; Bessire et al., 2011).

The application of these assays for testing the diffusion barrier properties of the leaf cuticle led to the identification of mutants defective in cutin formation and the subsequent identification of many genes involved in this process (Kurdyukov et al., 2006a; Bessire et al., 2007; Bird et al., 2007; Bessire et al., 2011). Some of these mutants have a lesion in both wax and cutin deposition (Bird et al., 2007), but none of the mutants have a wax-specific phenotype. This may be due to the fact that the applied assays detect only strongly permeable cuticles. It remains to be seen whether the same assays can also be used to identify minor changes in cuticle permeability by altering assay conditions (e.g., through the use of long incubation periods). Chemical characterization of the cutin mutants identified to date did not reveal a clear relationship between cutin/wax amount and cuticle permeability. Even plants that have significantly more wax and cutin than wt plants may exhibit increased cuticle permeability, indicating that the organization of cutin and wax is crucial in establishing a functional barrier at the leaf/atmosphere interface. However, a co-regulated increase of wax and cutin may indeed improve cuticular properties, as demonstrated by the cer9 Arabidopsis mutant (see above) (Lu et al., 2012).

The role of the cuticle in biotic interactions

Since the first report describing Arabidopsis as a viral host in 1981 (Balazs and Lebeurier, 1981), significant advances have been made in our understanding of the mechanisms by which this model plant defends itself against a plethora of pathogens (Nishimura and Dangl, 2010). In the past two decades, the use of Arabidopsis mutants has contributed enormously to the advancement of our knowledge of the molecular basis of plant resistance. Reviews on this topic can be found in other chapters of TAB (Katagiri et al., 2002; Knepper and Day, 2008; Compagnon et al., 2009). Because the plant cuticle is a specialization of the extracellular matrix of epidermal cells in aerial tissues, any alteration of cell wall metabolism in these cells could affect the cuticle and vice versa. The role of the cell wall in pathogen interactions has been recently reviewed (Huckelhoven, 2007; Cantu et al., 2008), so this topic will not be addressed in this section. Here, we will discuss the complex relationship between plant-pathogen interactions and cuticle biology.

The role of the cuticle in plant development

The role of the cuticle in post-embryonic development

Organs of aerial parts of the plant are generated from the shoot apical meristem and the floral apical meristem. Although adjacent organs maintain tight contact with each other, they remain separate and do not fuse. The fiddlehead (fdh) mutant and a number of other mutants have been identified because of striking organ fusions that occur among the rosette leaves or the floral organs (Lolle et al., 1992; Lolle et al., 1998; Wellesen et al., 2001). Organ fusions and the rapid leaching of chlorophyll were well phenocopied by transgenic Arabidopsis plants expressing a fungal cutinase, supporting the hypothesis that the fusion phenotypes are associated with a defective cuticle (Figure 10) (Sieber et al., 2000). Although FDH has been identified as member of the KCS family based on sequence similarity, the biochemical role of this protein in cutin formation has not yet been elucidated (Yephremov et al., 1999; Pruitt et al., 2000; Efremova et al., 2004). Similar to the fdh mutant, a link between cutin deposition and organ fusions has been identified in a number of other mutants that display a similar phenotype, including the lcr, bdg, cer10, ace/hth, and abgc11 mutants (Yephremov et al., 1999; Pruitt et al., 2000; Wellesen et al., 2001 ; Kurdyukov et al., 2006a; Kurdyukov et al., 2006b; Bird et al., 2007; Panikashvili et al., 2007; Panikashvili et al., 2009; Voisin et al., 2009; Panikashvili et al., 2010; Panikashvili et al., 2011; Shi et al., 2011). Based on observed alterations in cutin amount and composition, a role for the mutated genes in cutin biosynthesis was established (see above). Because organ fusions have been described in mutants with a block in different steps of the cutin biosynthetic pathway, namely in cutin monomer synthesis (i.e., lacs2, ace/hth, and lcr) (Wellesen et al., 2001; Kurdyukov et al., 2006b; Bessire et al., 2007), transport (i.e., pec1, abcg11 and abcg13) (Bird et al., 2007; Panikashvili et al., 2007; Bessire et al., 2011; Panikashvili et al., 2011), and extracellular polyester formation (i.e., bdg) (Kurdyukov et al., 2006a), as well as in CUTE plants that degrade their cutin (Sieber et al., 2000), these developmental changes may be triggered by an increased permeability of the cuticle during the early development of the organ undergoing fusion. However, no direct correlation between cuticle permeability and aberrations in epidermal cell development could be detected in mature plants (Yephremov and Schreiber, 2005).

In most organ fusion mutants, ultrastructural changes in the cuticle could be identified in zones that were not fused (see above). In the zones of organ fusion, the cell wall polysaccharides of epidermal cells were joined, as the cuticle was partially disrupted or entirely missing, such as in the bdg and lacs2 mutants (Figure 11A, 11B)(Sieber et al., 2000; Kurdyukov et al., 2006a; Bessire et al., 2007; Voisin et al., 2009; Bessire et al., 2011). Only the cuticle of fdh was not visibly altered, although the cuticle permeability of this mutant was even higher than that of other cuticle mutants displaying organ fusions (Figure 11C) (Voisin et al., 2009).

Figure 10.

Visual phenotype of a cutinase-expressing transgenic Arabidopsis plant.

Inflorescence of a cutinase-expressing transgenic Arabidopsis plant with multiple organ fusion events between a series of flowers of different ages and between flower organs. I: inflorescence, mF: mature flower, sF: senescing flower. The points of cell contact are hidden.

The cells in a fusion zone retain their epidermal identity as they continue the differentiation process and form guard cells in the fusion zone. Although stomata are present, they are usually closed and are not likely to be physiologically functional (Sieber et al., 2000) (Figure 11D). However, a number of changes in epidermal patterning were observed, such as an altered shape of the epidermal pavement cells of the rosette leaves, as in the dcr mutant, or alterations in the number and shape of trichomes, as in the fdh, lcr, yore-yore/wax2/cer3 and dcr mutants (Wellesen et al., 2001; Kurata et al., 2003; Yephremov and Schreiber, 2005; Kurdyukov et al., 2006a; Panikashvili et al., 2009). Thus, feedback regulation loops may control both the formation of a functional cuticle and epidermal differentiation (Yephremov et al., 1999; Wellesen et al., 2001; Kurdyukov et al., 2006a; Panikashvili et al., 2009; Panikashvili et al., 2010; Shi et al., 2011). The Arabidopsis cuticle mutants with the most pronounced phenotypes, such as CUTE-expressing plants, AtMYB41-over-expressing plants, and abcg11 and bgd mutants, also exhibit dwarfism, indicating a pleiotropic influence of impaired cuticle formation on plant development (Sieber et al., 2000; Kurdyukov et al., 2006a; Bird et al., 2007; Panikashvili et al., 2007; Cominelli et al., 2008).

A comparison of the transcriptomes of expanding rosette leaves in the bdg, lcr and fdh mutants (Voisin et al., 2009) identified a set of approximately 90 genes that are upregulated in all three genotypes. Among these genes, a number of genes are involved in cutin and wax biosynthesis, indicating that cell walls and cuticles are remodeled and defenses against abiotic stresses and pathogens are activated in organ fusion mutants. A regulator of organ fusions has been identified from in silico studies of the transcriptomes of organ fusion mutants (Voisin et al., 2009). A mutation in SERRATE (SE) (At2g27100) suppresses the formation of organ fusions in se 1cr and se bdg double mutants. SERRATE (SE) (At2g27100) encodes a zinc finger protein that functions in RNA splicing and the processing of pre-miRNAs to mature miRNAs, linking the regulation of RNA processing to cuticle formation (Prigge and Wagner, 2001; Grigg et al., 2005; Lobbes et al., 2006; Yang et al., 2006). However, the exact function of SE in cuticle formation has not been established (Figure 9).

Organ fusion always occurs during organ formation, when the epidermis of a newly developing organ is in tight contact with the epidermis of other organs (Sieber et al., 2000). Once the leaves or flowers have grown out, they are no longer fusion competent. However, if organs that are tightly fused are exposed to mechanical stress during development, the point of fusion may break, forming a wound site (Sieber et al., 2000; Wellesen et al., 2001). The formation of cutin is not only under developmental control but also under environmental control. Therefore, the type and severity of organ fusions may depend on the plant environment, which explains why the extent of organ fusion often varies when a single genotype (e.g. the lacs-1lacs2 double mutant) is grown under different conditions (Lu et al., 2009; Weng et al., 2010).

Figure 11.

Organ fusions between epidermal cells of different organs in cutin mutants.

Organ fusions between epidermal cell layers of the leaves (A-C) and between inflorescence stems (D-E). Visual appearence of the organ fusion zone in the Arabidopsis mutants fdh (A) (picture by Jean Mac Donald-Petetot, University Lausanne ), bdg (B) (from (Kurdyukov et al., 2006a) and lacs2–3 (C) (from Bessire et al., 2007), as well as in transgenic plants expressing a fungal cutinase (D, E) (from Sieber et al. 2000). Scale bars (A–E): 800nm.

The role of root diffusion barriers in water and nutrient uptake

The organization of the apoplastic barrier in the shoots of Arabidopsis and other herbaceous plants without extensive secondary growth is fairly simple, with the cuticle covering the surface of all organs and forming ledges surrounding the stomata. In contrast, the structure of the apoplastic transport barriers in the roots is more complex. Roots are designed for taking up water and dissolved nutrients, but unfavorable or toxic compounds must be filtered and restricted from entering the plant (Steudle and Peterson, 1998). In addition, harmful microorganisms, especially pathogens, which occur in the soil at a much higher density than in air, must be prevented from colonizing the living plant tissues. To solve this problem, Casparian strips and suberin lamellae are deposited in the apoplast of the root in its primary developmental state, chemically modifying the primary cell wall (Schreiber, 2010). In Arabidopsis, these structures are found in the endodermis (Franke et al., 2005), while in other species, Casparian strips and suberin lamellae are also found in the hypodermis (Wilson and Peterson, 1983). In the secondary developmental state of the Arabidopsis root, one to two suberized cells cover the outer root surface (Franke et al., 2005).

Water and nutrient uptake are thought to be highest in younger root sections of Arabidopsis. Thus, the endodermis, with its cell wall modifications, should be a major barrier that allows the plant to separate interior and exterior compartments. Asymmetric localization of membrane bound transporters (influx and efflux transporters) in the endodermis supports this idea (Alassimone et al., 2010). Dyes that act as apoplastic tracers have been used for microscopic investigations of sites of water and solute uptake and for localization of apoplastic transport barriers (Bayliss et al., 1996; Aloni et al., 1998). In Arabidopsis, it was demonstrated that the radial movement of the dye in the cell wall was restricted at the outer side of the endodermis (Alassimone et al., 2010). This finding provides evidence that water and ions cannot diffuse radially in the cell wall space across the endodermis. However, it must be noted that tracer dyes are fairly large molecules with molecular weights of several hundred Daltons (e.g., propidium iodine has a molecular weight of 668 Da). Water and dissolved nutrients are much smaller (10–100 Da); thus, an endodermal barrier may not efficiently inhibit radial movement of these molecules.

The radial flow of water and solutes can be measured in roots using the pressure probe technique (Steudle et al., 1993). Three pathways represent routes of radial water and nutrient uptake into roots: (i) the apoplastic pathway, (ii) the symplastic pathway, and (iii) the transcellular pathway, which represents a mixture of the former 2 pathways (Steudle, 1994). Whole root systems of Arabidopsis have been mounted onto the pressure probe, and overall water flow (i.e., water uptake via all three pathways) was measured by applying a hydrostatic pressure gradient between the surrounding medium and the root xylem vessels (Ranathunge and Schreiber, 2011). The determined hydraulic hydrostatic conductivity was 3.7 x 10-8 m s^-1 MPa^-1. This value is not significantly different from those measured for rice (Ranathunge et al., 2003). Two to ten times higher values were reported for maize (Zimmermann and Steudle, 1998). In parallel, osmotic pressure gradients can be used for measuring the symplastic flow of water. Thus, from hydraulic conductivities, which are obtained with hydrostatic and osmotic pressure gradients, the extent to which the apoplastic pathway contributes to the overall water uptake of the root can be estimated. Because the hydraulic conductivity of the Arabidopsis root did not differ with the application of different pressure gradients (i.e., hydraulic vs. osmotic), water uptake in Arabidopsis primarily occurs via the symplastic pathway (Ranathunge and Schreiber, 2011).

Mutants characterized by a 60% decrease (e.g., cyp86a1/horst) (Höfer et al., 2008) or a 100% increase (e.g., esb1) in the amount of root suberin (Baxter et al., 2009) were also investigated using the pressure probe. Hydraulic conductivities in the presence of a hydrostatic pressure gradient significantly increased (by a factor of three) in roots with reduced suberin amounts, while they were unchanged in the presence of an osmotic pressure gradient. Thus, the apoplastic barrier was less effective towards radial water flow in the mutant with reduced suberization. However, in comparison to wt, no differences in hydraulic conductivities were observed with esb1 mutants, which have increased root suberization. Interestingly, esb1 was previously characterized as having a significant reduction in calcium, nickel and iron content in the shoot. This shoot phenotype might represent an indirect measure of reduced ion translocation through the root, but it may also be a secondary effect of physiological adaptations that affect ion homeostasis. A recently identified mutant in sphingolipid metabolism, which also shows suberin phenotypes and changes in the leaf ionome, supports the former hypothesis (Chao et al., 2011). In conclusion, the relationship between the amount of the aliphatic biopolymer suberin in roots and radial hydraulic conductivity is not simple. The molecular arrangement of suberin in the cell wall and the amount of other cell wall polymers, such as lignin, should also be considered as potential factors affecting radial hydraulic conductivity in the Arabidopsis root.