Leaf Senescence Signaling: The Ca2+-Conducting Arabidopsis Cyclic Nucleotide Gated Channel2 Acts through Nitric Oxide to Repress Senescence Programming

Ca²⁺ and nitric oxide (NO) are essential components involved in plant senescence signaling cascades. In other signaling pathways, NO generation can be dependent on cytosolic Ca²⁺. The Arabidopsis (Arabidopsis thaliana) mutant dnd1 lacks a plasma membrane-localized cation channel (CNGC2). We recently demonstrated that this channel affects plant response to pathogens through a signaling cascade involving Ca²⁺ modulation of NO generation; the pathogen response phenotype of dnd1 can be complemented by application of a NO donor. At present, the interrelationship between Ca²⁺ and NO generation in plant cells during leaf senescence remains unclear. Here, we use dnd1 plants to present genetic evidence consistent with the hypothesis that Ca²⁺ uptake and NO production play pivotal roles in plant leaf senescence. Leaf Ca²⁺ accumulation is reduced in dnd1 leaves compared to the wild type. Early senescence-associated phenotypes (such as loss of chlorophyll, expression level of senescence-associated genes, H₂O₂ generation, lipid peroxidation, tissue necrosis, and increased salicylic acid levels) were more prominent in dnd1 leaves compared to the wild type. Application of a Ca²⁺ channel blocker hastened senescence of detached wild-type leaves maintained in the dark, increasing the rate of chlorophyll loss, expression of a senescence-associated gene, and lipid peroxidation. Pharmacological manipulation of Ca²⁺ signaling provides evidence consistent with genetic studies of the relationship between Ca²⁺ signaling and senescence with the dnd1 mutant. Basal levels of NO in dnd1 leaf tissue were lower than that in leaves of wild-type plants. Application of a NO donor effectively rescues many dnd1 senescence-related phenotypes. Our work demonstrates that the CNGC2 channel is involved in Ca²⁺ uptake during plant development beyond its role in pathogen defense response signaling. Work presented here suggests that this function of CNGC2 may impact downstream basal NO production in addition to its role (also linked to NO signaling) in pathogen defense responses and that this NO generation acts as a negative regulator during plant leaf senescence signaling.Senescence can be considered as the final stage of a plant’s development. During this process, nutrients will be reallocated from older to younger parts of the plant, such as developing leaves and seeds. Leaf senescence has been characterized as a type of programmed cell death (PCD; Gan and Amasino, 1997; Quirino et al., 2000; Lim et al., 2003). During senescence, organelles such as chloroplasts will break down first. Biochemical changes will also occur in the peroxisome during this process. When the chloroplast disassembles, it is easily observed as a loss of chlorophyll. Mitochondria, the source of energy for cells, will be the last cell organelles to undergo changes during the senescence process (Quirino et al., 2000). At the same time, other catabolic events (e.g. protein and lipid breakdown, etc.) are occurring (Quirino et al., 2000). Hormones may also contribute to this process (Gepstein, 2004). From this information we can infer that leaf senescence is regulated by many signals.Darkness treatment can induce senescence in detached leaves (Poovaiah and Leopold, 1973; Chou and Kao, 1992; Weaver and Amasino, 2001; Chrost et al., 2004; Guo and Crawford, 2005; Ülker et al., 2007). Ca²⁺ can delay the senescence of detached leaves (Poovaiah and Leopold, 1973) and leaf senescence induced by methyl jasmonate (Chou and Kao, 1992); the molecular events that mediate this effect of Ca²⁺ are not well characterized at present.Nitric oxide (NO) is a critical signaling molecule involved in many plant physiological processes. Recently, published evidence supports NO acting as a negative regulator during leaf senescence (Guo and Crawford, 2005; Mishina et al., 2007). Abolishing NO generation in either loss-of-function mutants (Guo and Crawford, 2005) or transgenic Arabidopsis (Arabidopsis thaliana) plants expressing NO degrading dioxygenase (NOD; Mishina et al., 2007) leads to an early senescence phenotype in these plants compared to the wild type. Corpas et al. (2004) showed that endogenous NO is mainly accumulated in vascular tissues of pea (Pisum sativum) leaves. This accumulation is significantly reduced in senescing leaves (Corpas et al., 2004). Corpas et al. (2004) also provided evidence that NO synthase (NOS)-like activity (i.e. generation of NO from l-Arg) is greatly reduced in senescing leaves. Plant NOS activity is regulated by Ca²⁺/calmodulin (CaM; Delledonne et al., 1998; Corpas et al., 2004, 2009; del Río et al., 2004; Valderrama et al., 2007; Ma et al., 2008). These studies suggest a link between Ca²⁺ and NO that could be operating during senescence.In animal cells, all three NOS isoforms require Ca²⁺/CaM as a cofactor (Nathan and Xie, 1994; Stuehr, 1999; Alderton et al., 2001). Notably, animal NOS contains a CaM binding domain (Stuehr, 1999). It is unclear whether Ca²⁺/CaM can directly modulate plant NOS or if Ca²⁺/CaM impacts plant leaf development/senescence through (either direct or indirect) effects on NO generation. However, recent studies from our lab suggest that Ca²⁺/CaM acts as an activator of NOS activity in plant innate immune response signaling (Ali et al., 2007; Ma et al., 2008).Although Arabidopsis NO ASSOCIATED PROTEIN1 (AtNOA1; formerly named AtNOS1) was thought to encode a NOS enzyme, no NOS-encoding gene has yet been identified in plants (Guo et al., 2003; Crawford et al., 2006; Zemojtel et al., 2006). However, the AtNOA1 loss-of-function mutant does display reduced levels of NO generation, and several groups have used the NO donor sodium nitroprusside (SNP) to reverse some low-NO related phenotypes in Atnoa1 plants (Guo et al., 2003; Bright et al., 2006; Zhao et al., 2007). Importantly, plant endogenous NO deficiency (Guo and Crawford, 2005; Mishina et al., 2007) or abscisic acid/methyl jasmonate (Hung and Kao, 2003, 2004) induced early senescence can be successfully rescued by application of exogenous NO. Addition of NO donor can delay GA-elicited PCD in barley (Hordeum vulgare) aleurone layers as well (Beligni et al., 2002).It has been suggested that salicylic acid (SA), a critical pathogen defense metabolite, can be increased in natural (Morris et al., 2000; Mishina et al., 2007) and transgenic NOD-induced senescent Arabidopsis leaves (Mishina et al., 2007). Pathogenesis related gene1 (PR1) expression is up-regulated in transgenic Arabidopsis expressing NOD (Mishina et al., 2007) and in leaves of an early senescence mutant (Ülker et al., 2007).Plant cyclic nucleotide gated channels (CNGCs) have been proposed as candidates to conduct extracellular Ca²⁺ into the cytosol (Sunkar et al., 2000; Talke et al., 2003; Lemtiri-Chlieh and Berkowitz, 2004; Ali et al., 2007; Demidchik and Maathuis, 2007; Frietsch et al., 2007; Kaplan et al., 2007; Ma and Berkowitz, 2007; Urquhart et al., 2007; Ma et al., 2009a, 2009b). Arabidopsis “defense, no death” (dnd1) mutant plants have a null mutation in the gene encoding the plasma membrane-localized Ca²⁺-conducting CNGC2 channel. This mutant also displays no hypersensitive response to infection by some pathogens (Clough et al., 2000; Ali et al., 2007). In addition to involvement in pathogen-mediated Ca²⁺ signaling, CNGC2 has been suggested to participate in the process of leaf development/senescence (Köhler et al., 2001). dnd1 mutant plants have high levels of SA and expression of PR1 (Yu et al., 1998), and spontaneous necrotic lesions appear conditionally in dnd1 leaves (Clough et al., 2000; Jirage et al., 2001). Endogenous H₂O₂ levels in dnd1 mutants are increased from wild-type levels (Mateo et al., 2006). Reactive oxygen species molecules, such as H₂O₂, are critical to the PCD/senescence processes of plants (Navabpour et al., 2003; Overmyer et al., 2003; Hung and Kao, 2004; Guo and Crawford, 2005; Zimmermann et al., 2006). Here, we use the dnd1 mutant to evaluate the relationship between leaf Ca²⁺ uptake during plant growth and leaf senescence. Our results identify NO, as affected by leaf Ca²⁺ level, to be an important negative regulator of leaf senescence initiation. Ca²⁺-mediated NO production during leaf development could control senescence-associated gene (SAG) expression and the production of molecules (such as SA and H₂O₂) that act as signals during the initiation of leaf senescence programs.     

Characterization of the Possible Roles for B Class MADS Box Genes in Regulation of Perianth Formation in Orchid

To investigate sepal/petal/lip formation in Oncidium Gower Ramsey, three paleoAPETALA3 genes, O. Gower Ramsey MADS box gene5 (OMADS5; clade 1), OMADS3 (clade 2), and OMADS9 (clade 3), and one PISTILLATA gene, OMADS8, were characterized. The OMADS8 and OMADS3 mRNAs were expressed in all four floral organs as well as in vegetative leaves. The OMADS9 mRNA was only strongly detected in petals and lips. The mRNA for OMADS5 was only strongly detected in sepals and petals and was significantly down-regulated in lip-like petals and lip-like sepals of peloric mutant flowers. This result revealed a possible negative role for OMADS5 in regulating lip formation. Yeast two-hybrid analysis indicated that OMADS5 formed homodimers and heterodimers with OMADS3 and OMADS9. OMADS8 only formed heterodimers with OMADS3, whereas OMADS3 and OMADS9 formed homodimers and heterodimers with each other. We proposed that sepal/petal/lip formation needs the presence of OMADS3/8 and/or OMADS9. The determination of the final organ identity for the sepal/petal/lip likely depended on the presence or absence of OMADS5. The presence of OMADS5 caused short sepal/petal formation. When OMADS5 was absent, cells could proliferate, resulting in the possible formation of large lips and the conversion of the sepal/petal into lips in peloric mutants. Further analysis indicated that only ectopic expression of OMADS8 but not OMADS5/9 caused the conversion of the sepal into an expanded petal-like structure in transgenic Arabidopsis (Arabidopsis thaliana) plants.The ABCDE model predicts the formation of any flower organ by the interaction of five classes of homeotic genes in plants (Yanofsky et al., 1990; Jack et al., 1992; Mandel et al., 1992; Goto and Meyerowitz, 1994; Jofuku et al., 1994; Pelaz et al., 2000, 2001; Theißen and Saedler, 2001; Pinyopich et al., 2003; Ditta et al., 2004; Jack, 2004). The A class genes control sepal formation. The A, B, and E class genes work together to regulate petal formation. The B, C, and E class genes control stamen formation. The C and E class genes work to regulate carpel formation, whereas the D class gene is involved in ovule development. MADS box genes seem to have a central role in flower development, because most ABCDE genes encode MADS box proteins (Coen and Meyerowitz, 1991; Weigel and Meyerowitz, 1994; Purugganan et al., 1995; Rounsley et al., 1995; Theißen and Saedler, 1995; Theißen et al., 2000; Theißen, 2001).The function of B group genes, such as APETALA3 (AP3) and PISTILLATA (PI), has been thought to have a major role in specifying petal and stamen development (Jack et al., 1992; Goto and Meyerowitz, 1994; Krizek and Meyerowitz, 1996; Kramer et al., 1998; Hernandez-Hernandez et al., 2007; Kanno et al., 2007; Whipple et al., 2007; Irish, 2009). In Arabidopsis (Arabidopsis thaliana), mutation in AP3 or PI caused identical phenotypes of second whorl petal conversion into a sepal structure and third flower whorl stamen into a carpel structure (Bowman et al., 1989; Jack et al., 1992; Goto and Meyerowitz, 1994). Similar homeotic conversions for petal and stamen were observed in the mutants of the AP3 and PI orthologs from a number of core eudicots such as Antirrhinum majus, Petunia hybrida, Gerbera hybrida, Solanum lycopersicum, and Nicotiana benthamiana (Sommer et al., 1990; Tröbner et al., 1992; Angenent et al., 1993; van der Krol et al., 1993; Yu et al., 1999; Liu et al., 2004; Vandenbussche et al., 2004; de Martino et al., 2006), from basal eudicot species such as Papaver somniferum and Aquilegia vulgaris (Drea et al., 2007; Kramer et al., 2007), as well as from monocot species such as Zea mays and Oryza sativa (Ambrose et al., 2000; Nagasawa et al., 2003; Prasad and Vijayraghavan, 2003; Yadav et al., 2007; Yao et al., 2008). This indicated that the function of the B class genes AP3 and PI is highly conserved during evolution.It has been thought that B group genes may have arisen from an ancestral gene through multiple gene duplication events (Doyle, 1994; Theißen et al., 1996, 2000; Purugganan, 1997; Kramer et al., 1998; Kramer and Irish, 1999; Lamb and Irish, 2003; Kim et al., 2004; Stellari et al., 2004; Zahn et al., 2005; Hernandez-Hernandez et al., 2007). In the gymnosperms, there was a single putative B class lineage that duplicated to generate the paleoAP3 and PI lineages in angiosperms (Kramer et al., 1998; Theißen et al., 2000; Irish, 2009). The paleoAP3 lineage is composed of AP3 orthologs identified in lower eudicots, magnolid dicots, and monocots (Kramer et al., 1998). Genes in this lineage contain the conserved paleoAP3- and PI-derived motifs in the C-terminal end of the proteins, which have been thought to be characteristics of the B class ancestral gene (Kramer et al., 1998; Tzeng and Yang, 2001; Hsu and Yang, 2002). The PI lineage is composed of PI orthologs that contain a highly conserved PI motif identified in most plant species (Kramer et al., 1998). Subsequently, there was a second duplication at the base of the core eudicots that produced the euAP3 and TM6 lineages, which have been subject to substantial sequence changes in eudicots during evolution (Kramer et al., 1998; Kramer and Irish, 1999). The paleoAP3 motif in the C-terminal end of the proteins was retained in the TM6 lineage and replaced by a conserved euAP3 motif in the euAP3 lineage of most eudicot species (Kramer et al., 1998). In addition, many lineage-specific duplications for paleoAP3 lineage have occurred in plants such as orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009), Ranunculaceae, and Ranunculales (Kramer et al., 2003; Di Stilio et al., 2005; Shan et al., 2006; Kramer, 2009).Unlike the A or C class MADS box proteins, which form homodimers that regulate flower development, the ability of B class proteins to form homodimers has only been reported in gymnosperms and in the paleoAP3 and PI lineages of some monocots. For example, LMADS1 of the lily Lilium longiflorum (Tzeng and Yang, 2001), OMADS3 of the orchid Oncidium Gower Ramsey (Hsu and Yang, 2002), and PeMADS4 of the orchid Phalaenopsis equestris (Tsai et al., 2004) in the paleoAP3 lineage, LRGLOA and LRGLOB of the lily Lilium regale (Winter et al., 2002), TGGLO of the tulip Tulipa gesneriana (Kanno et al., 2003), and PeMADS6 of the orchid P. equestris (Tsai et al., 2005) in the PI lineage, and GGM2 of the gymnosperm Gnetum gnemon (Winter et al., 1999) were able to form homodimers that regulate flower development. Proteins in the euAP3 lineage and in most paleoAP3 lineages were not able to form homodimers and had to interact with PI to form heterodimers in order to regulate petal and stamen development in various plant species (Schwarz-Sommer et al., 1992; Tröbner et al., 1992; Riechmann et al., 1996; Moon et al., 1999; Winter et al., 2002; Kanno et al., 2003; Vandenbussche et al., 2004; Yao et al., 2008). In addition to forming dimers, AP3 and PI were able to interact with other MADS box proteins, such as SEPALLATA1 (SEP1), SEP2, and SEP3, to regulate petal and stamen development (Pelaz et al., 2000; Honma and Goto, 2001; Theißen and Saedler, 2001; Castillejo et al., 2005).Orchids are among the most important plants in the flower market around the world, and research on MADS box genes has been reported for several species of orchids during the past few years (Lu et al., 1993, 2007; Yu and Goh, 2000; Hsu and Yang, 2002; Yu et al., 2002; Hsu et al., 2003; Tsai et al., 2004, 2008; Xu et al., 2006; Guo et al., 2007; Kim et al., 2007; Chang et al., 2009). Unlike the flowers in eudicots, the nearly identical shape of the sepals and petals as well as the production of a unique lip in orchid flowers make them a very special plant species for the study of flower development. Four clades (1–4) of genes in the paleoAP3 lineage have been identified in several orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009). Several works have described the possible interactions among these four clades of paleoAP3 genes and one PI gene that are involved in regulating the differentiation and formation of the sepal/petal/lip of orchids (Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009). However, the exact mechanism that involves the orchid B class genes remains unclear and needs to be clarified by more experimental investigations.O. Gower Ramsey is a popular orchid with important economic value in cut flower markets. Only a few studies have been reported on the role of MADS box genes in regulating flower formation in this plant species (Hsu and Yang, 2002; Hsu et al., 2003; Chang et al., 2009). An AP3-like MADS gene that regulates both floral formation and initiation in transgenic Arabidopsis has been reported (Hsu and Yang, 2002). In addition, four AP1/AGAMOUS-LIKE9 (AGL9)-like MADS box genes have been characterized that show novel expression patterns and cause different effects on floral transition and formation in Arabidopsis (Hsu et al., 2003; Chang et al., 2009). Compared with other orchids, the production of a large and well-expanded lip and five small identical sepals/petals makes O. Gower Ramsey a special case for the study of the diverse functions of B class MADS box genes during evolution. Therefore, the isolation of more B class MADS box genes and further study of their roles in the regulation of perianth (sepal/petal/lip) formation during O. Gower Ramsey flower development are necessary. In addition to the clade 2 paleoAP3 gene OMADS3, which was previously characterized in our laboratory (Hsu and Yang, 2002), three more B class MADS box genes, OMADS5, OMADS8, and OMADS9, were characterized from O. Gower Ramsey in this study. Based on the different expression patterns and the protein interactions among these four orchid B class genes, we propose that the presence of OMADS3/8 and/or OMADS9 is required for sepal/petal/lip formation. Further sepal and petal formation at least requires the additional presence of OMADS5, whereas large lip formation was seen when OMADS5 expression was absent. Our results provide a new finding and information pertaining to the roles for orchid B class MADS box genes in the regulation of sepal/petal/lip formation.     

Wettability,Polarity, and Water Absorption of Holm Oak Leaves: Effect of Leaf Side and Age

Plant trichomes play important protective functions and may have a major influence on leaf surface wettability. With the aim of gaining insight into trichome structure, composition, and function in relation to water-plant surface interactions, we analyzed the adaxial and abaxial leaf surface of holm oak (Quercus ilex) as a model. By measuring the leaf water potential 24 h after the deposition of water drops onto abaxial and adaxial surfaces, evidence for water penetration through the upper leaf side was gained in young and mature leaves. The structure and chemical composition of the abaxial (always present) and adaxial (occurring only in young leaves) trichomes were analyzed by various microscopic and analytical procedures. The adaxial surfaces were wettable and had a high degree of water drop adhesion in contrast to the highly unwettable and water-repellent abaxial holm oak leaf sides. The surface free energy and solubility parameter decreased with leaf age, with higher values determined for the adaxial sides. All holm oak leaf trichomes were covered with a cuticle. The abaxial trichomes were composed of 8% soluble waxes, 49% cutin, and 43% polysaccharides. For the adaxial side, it is concluded that trichomes and the scars after trichome shedding contribute to water uptake, while the abaxial leaf side is highly hydrophobic due to its high degree of pubescence and different trichome structure, composition, and density. Results are interpreted in terms of water-plant surface interactions, plant surface physical chemistry, and plant ecophysiology.Plant surfaces have an important protecting function against multiple biotic and abiotic stress factors (Riederer, 2006). They may, for example, limit the attack of insects (Eigenbrode and Jetter, 2002) or pathogenic fungi (Gniwotta et al., 2005; Łaźniewska et al., 2012), avoid damage caused by high intensities of UV and visible radiation (Reicosky and Hanover, 1978; Karabourniotis and Bormann, 1999), help to regulate leaf temperature (Ehleringer and Björkman, 1978; Ripley et al., 1999), and chiefly prevent plant organs from dehydration (Riederer and Schreiber, 2001).The epidermis of plants has been found to have a major degree of physical and chemical variability and may often contain specialized cells such as trichomes or stomata (Roth-Nebelsick et al., 2009; Javelle et al., 2011). Most aerial organs are covered with an extracellular and generally lipid-rich layer named the cuticle, which is typically composed of waxes embedded in (intracuticular waxes) or deposited on (epicuticular waxes) a biopolymer matrix of cutin, forming a network of cross-esterified hydroxy C₁₆ and/or C₁₈ fatty acids, and/or cutan, with variable amounts of polysaccharides and phenolics (Domínguez et al., 2011; Yeats and Rose, 2013). Different nano- and/or microscale levels of plant surface sculpturing have been observed by scanning electron microscopy (SEM), generally in relation to the topography of epicuticular waxes, cuticular folds, and epidermal cells (Koch and Barthlott, 2009). Such surface features together with their chemical composition (Khayet and Fernández, 2012) may lead to a high degree of roughness and hydrophobicity (Koch and Barthlott, 2009; Konrad et al., 2012). The interactions of plant surfaces with water have been addressed in some investigations (Brewer et al., 1991; Brewer and Smith, 1997; Pandey and Nagar, 2003; Hanba et al., 2004; Dietz et al., 2007; Holder, 2007a, 2007b; Fernández et al., 2011, 2014; Roth-Nebelsick et al., 2012; Wen et al., 2012; Urrego-Pereira et al., 2013) and are a topic of growing interest for plant ecophysiology (Helliker and Griffiths, 2007; Aryal and Neuner, 2010; Limm and Dawson, 2010; Kim and Lee, 2011; Berry and Smith, 2012; Berry et al., 2013; Rosado and Holder, 2013; Helliker, 2014). On the other hand, the mechanisms of foliar uptake of water and solutes by plant surfaces are still not fully understood (Fernández and Eichert, 2009; Burkhardt and Hunsche, 2013), but they may play an important ecophysiological role (Limm et al., 2009; Johnstone and Dawson, 2010; Adamec, 2013; Berry et al., 2014).The importance of trichomes and pubescent layers on water drop-plant surface interactions and on the subsequent potential water uptake into the organs has been analyzed in some investigations (Fahn, 1986; Brewer et al., 1991; Grammatikopoulos and Manetas, 1994; Brewer and Smith, 1997; Pierce et al., 2001; Kenzo et al., 2008; Fernández et al., 2011, 2014; Burrows et al., 2013). Trichomes are unicellular or multicellular and glandular or nonglandular appendages, which originate from epidermal cells only and develop outwards on the surface of plant organs (Werker, 2000). Nonglandular trichomes are categorized according to their morphology and exhibit a major variability in size, morphology, and function. On the other hand, glandular trichomes are classified by the secretory materials they excrete, accumulate, or absorb (Johnson, 1975; Werker, 2000; Wagner et al., 2004). Trichomes can be often found in xeromorphic leaves and in young organs (Fahn, 1986; Karabourniotis et al., 1995). The occurrence of protecting leaf trichomes has been also reported for Mediterranean species such as holm oak (Quercus ilex; Karabourniotis et al., 1995, 1998; Morales et al., 2002; Karioti et al., 2011; Camarero et al., 2012). There is limited information about the nature of the surface of trichomes, but they are also covered with a cuticle similarly to other epidermal cell types (Fernández et al., 2011, 2014).In this study and using holm oak as a model, we assessed, for the first time, the leaf surface-water relations of the abaxial (always pubescent) versus the adaxial (only pubescent in developing leaves and for a few months) surface, including their capacity to absorb surface-deposited water drops. Based on membrane science methodologies (Fernández et al., 2011; Khayet and Fernández, 2012) and following a new integrative approach, the chemical, physical, and anatomical properties of holm oak leaf surfaces and trichomes were analyzed, with the aim of addressing the following questions. Are young and mature adaxial and abaxial leaf surfaces capable of absorbing water deposited as drops on to the surfaces? Are young and mature abaxial and adaxial leaf surfaces similar in relation to their wettability, hydrophobicity, polarity, work of adhesion (W_a) for water, solubility parameter (δ), and surface free energy (γ)? What is the physical and chemical nature of the adaxial versus the abaxial trichomes, chiefly in relation to young leaves?     

Elucidating the Role of Transport Processes in Leaf Glucosinolate Distribution

In Arabidopsis (Arabidopsis thaliana), a strategy to defend its leaves against herbivores is to accumulate glucosinolates along the midrib and at the margin. Although it is generally assumed that glucosinolates are synthesized along the vasculature in an Arabidopsis leaf, thereby suggesting that the margin accumulation is established through transport, little is known about these transport processes. Here, we show through leaf apoplastic fluid analysis and glucosinolate feeding experiments that two glucosinolate transporters, GTR1 and GTR2, essential for long-distance transport of glucosinolates in Arabidopsis, also play key roles in glucosinolate allocation within a mature leaf by effectively importing apoplastically localized glucosinolates into appropriate cells. Detection of glucosinolates in root xylem sap unambiguously shows that this transport route is involved in root-to-shoot glucosinolate allocation. Detailed leaf dissections show that in the absence of GTR1 and GTR2 transport activity, glucosinolates accumulate predominantly in leaf margins and leaf tips. Furthermore, we show that glucosinolates accumulate in the leaf abaxial epidermis in a GTR-independent manner. Based on our results, we propose a model for how glucosinolates accumulate in the leaf margin and epidermis, which includes symplasmic movement through plasmodesmata, coupled with the activity of putative vacuolar glucosinolate importers in these peripheral cell layers.Feeding behavior of herbivorous insects and distribution of defense compounds in plants have been suggested to be a result of an arms race between plants and insects that has spanned millions of years (Ehrlich and Raven, 1964). Whether insects adapted first to plants or the other way around is an ongoing debate in this research field (Schoonhoven et al., 2005; Ali and Agrawal, 2012). Leaf margin accumulation of defense compounds has been demonstrated in various plant species (Gutterman and Chauser-Volfson, 2000; Chauser-Volfson et al., 2002; Kester et al., 2002; Cooney et al., 2012). In the model plant Arabidopsis (Arabidopsis thaliana), higher concentration of glucosinolates, which constitute a major part of the chemical defense system in this plant (Kliebenstein et al., 2001a; Halkier and Gershenzon, 2006), was found at the leaf midrib and margins compared with the leaf lamina (Shroff et al., 2008; Sønderby et al., 2010). This nonuniform leaf distribution of glucosinolates appeared to explain the feeding pattern of a generalist herbivore (Helicoverpa armigera), as it avoided feeding at the leaf margin and midrib (Shroff et al., 2008). A similar feeding pattern on Arabidopsis was observed for a different generalist herbivore, Spodoptera littoralis (Schweizer et al., 2013). Interestingly, S. littoralis was shown to favor feeding from Arabidopsis leaf margins in glucosinolate-deficient mutants (Schweizer et al., 2013), which could indicate an inherent preference for margin feeding and that Arabidopsis adapted to such behavior by accumulating defense compounds here. A damaged leaf margin may be more critical for leaf stability than damage to inner leaf parts (Shroff et al., 2008), further motivating protection of this tissue. The margin-feeding preference of S. littoralis might be explained by better nutritional value of the leaf margin cells (Schweizer et al., 2013), which has been shown to consist of specialized elongated cell files (Koroleva et al., 2010; Nakata and Okada, 2013).Other distribution patterns have been reported for glucosinolates in an Arabidopsis leaf. A study investigating spatiotemporal metabolic shifts during senescence in Arabidopsis reported that fully expanded mature leaves exhibited a glucosinolate gradient from base to tip, with highest level of glucosinolates at the leaf base (Watanabe et al., 2013). In contrast to the horizontal plane, less has been reported on distribution of glucosinolates in the vertical plane of a leaf. A localization study of cyanogenic glucosides, defense molecules related to glucosinolates (Halkier and Gershenzon, 2006), determined that these compounds primarily were located in the epidermis of sorghum (Sorghum bicolor; Kojima et al., 1979). Whereas epidermis-derived trichomes in Arabidopsis were recently demonstrated to contain glucosinolates and to express glucosinolate biosynthetic genes (Frerigmann et al., 2012), no studies have investigated glucosinolates in the epidermal cell layer.Based on promoter-GUS studies, biosynthesis of glucosinolates in leaves of Arabidopsis has been associated with primarily major and minor veins in leaves and silique walls (Mikkelsen et al., 2000; Reintanz et al., 2001; Tantikanjana et al., 2001; Chen et al., 2003; Grubb et al., 2004; Schuster et al., 2006; Gigolashvili et al., 2007; Li et al., 2011; Redovniković et al., 2012). The discrepancy between vasculature-associated glucosinolate biosynthesis and margin accumulation of glucosinolates suggests that transport processes must be involved in establishing the distribution pattern of glucosinolates within a leaf.Plant transport systems include the apoplastic xylem, the symplastic phloem, and plasmodesmata. Xylem transport is mainly driven by an upward pull generated by transpiration from aerial plant organs, thereby directing transport to sites of rapid evaporation (such as leaf margins; Sattelmacher, 2001). Phloem flow is facilitated by an osmosis-regulated hydrostatic pressure difference between source and sink tissue, primarily generated by Suc bulk flow (Lucas et al., 2013). Plasmodesmata are intercellular channels that establish symplasmic pathways between neighboring cells, and most cell types in a plant are symplastically connected via plasmodesmata (Roberts and Oparka, 2003). Translocation of small molecules in these channels is driven by diffusion and is regulated developmentally as well as spatially to form symplastically connected domains (Roberts and Oparka, 2003; Christensen et al., 2009). To what extent any of these transport processes are involved in establishing specific distribution patterns of glucosinolates within leaves is not known.Recently, two plasma membrane-localized, glucosinolate-specific importers, GLUCOSINOLATE TRANSPORTER1 (GTR1) and GTR2, were identified in Arabidopsis (Nour-Eldin et al., 2012). In leaf, their expression patterns were shown to be in leaf veins (GTR1 and GTR2) and surrounding mesophyll cells (GTR1; Nour-Eldin et al., 2012). Absence of aliphatic and indole glucosinolates in seeds of the gtr1gtr2 double knockout (dKO) mutant (gtr1gtr2 dKO) demonstrated that these transporters are essential for long-distance glucosinolate transport to the seeds and indicates a role in phloem loading (Nour-Eldin et al., 2012). Another study investigating long-distance transport of glucosinolates in the 3-week-old wild type and gtr1gtr2 dKO indicated that GTR1 and GTR2 were involved in bidirectional transport of aliphatic glucosinolates between root and shoot via both phloem and xylem pathways (Andersen et al., 2013). The authors suggested a role for GTR1 and GTR2 in the retention of long-chained aliphatic glucosinolates in roots by removing the compounds from the xylem (Andersen et al., 2013).Identification of the glucosinolate transporters GTR1 and GTR2 has provided a molecular tool to investigate the role of transport processes in establishing leaf glucosinolate distribution. In this study, we have performed a detailed spatial investigation of the distribution of an exogenously fed glucosinolate (sinigrin) and endogenous glucosinolates within mature wild-type and gtr1gtr2 dKO Arabidopsis leaves, achieved by collecting and analyzing leaf parts at the horizontal (y axis: petiole, base, and tip; x axis: midrib, lamina, and margin) as well as at the vertical leaf plane (z axis: abaxial epidermis). Furthermore, we analyze wild-type and gtr1gtr2 dKO root xylem sap and leaf apoplastic fluids for glucosinolates. Based on our results, we propose a model where GTR1 and GTR2 import glucosinolates from the apoplast to the symplast and where the glucosinolate distribution pattern within an Arabidopsis leaf is established via symplasmic movement of glucosinolates through plasmodesmata, coupled with the activity of putative vacuolar glucosinolate importers in peripheral cell layers.