Overexpression of Flavodoxin in Bacteroids Induces Changes in Antioxidant Metabolism Leading to Delayed Senescence and Starch Accumulation in Alfalfa Root Nodules

Sinorhizobium meliloti cells were engineered to overexpress Anabaena variabilis flavodoxin, a protein that is involved in the response to oxidative stress. Nodule natural senescence was characterized in alfalfa (Medicago sativa) plants nodulated by the flavodoxin-overexpressing rhizobia or the corresponding control bacteria. The decline of nitrogenase activity and the nodule structural and ultrastructural alterations that are associated with nodule senescence were significantly delayed in flavodoxin-expressing nodules. Substantial changes in nodule antioxidant metabolism, involving antioxidant enzymes and ascorbate-glutathione cycle enzymes and metabolites, were detected in flavodoxin-containing nodules. Lipid peroxidation was also significantly lower in flavodoxin-expressing nodules than in control nodules. The observed amelioration of the oxidative balance suggests that the delay in nodule senescence was most likely due to a role of the protein in reactive oxygen species detoxification. Flavodoxin overexpression also led to high starch accumulation in nodules, without reduction of the nitrogen-fixing activity.Symbiotic nodules have a limited functional life that varies among different legume species. Nodule senescence is the sequence of structural, molecular, biochemical, and physiological events taking place in the process that a mature and functional nodule undergoes leading to the loss of the nitrogen-fixing activity and culminating in cell death of symbiotic tissue (Swaraj and Bishnoi, 1996; Puppo et al., 2005; Van de Velde et al., 2006).Various models have been proposed to explain the mechanisms that trigger the process of natural or stress-induced nodule senescence. However, it is generally accepted that a senescence-inducing signal from the plant causes a decrease in antioxidant levels and thus an increase in reactive oxygen species (ROS) up to a point of no return. Numerous studies have shown that ROS and antioxidant systems are involved in natural (Lucas et al., 1998; Evans et al., 1999; Hernández-Jiménez et al., 2002; Puppo et al., 2005) as well as induced (Dalton et al., 1993; Becana et al., 2000; Hernández-Jiménez et al., 2002; Matamoros et al., 2003) nodule senescence. Nitrogen fixation is very sensitive to ROS, and nitrogenase activity drastically decreases during nodule senescence (Dalton et al., 1986).Antioxidant systems that protect cells from oxidative damage have been described in symbiotic nodules (Dalton et al., 1986, 1993; Evans et al., 1999; Becana et al., 2000; Matamoros et al., 2003; Puppo et al., 2005). These include the enzymes superoxide dismutase (SOD), catalase, and peroxidase. Another enzymatic system associated with ROS detoxification is the ascorbate-glutathione pathway, which includes ascorbate peroxidase (APX), dehydroascorbate reductase (DHAR), monodehydroascorbate reductase (MDHAR), and glutathione reductase (GR; Dalton et al., 1986, 1992; Noctor and Foyer 1998; Becana et al., 2000). Ascorbate and reduced glutathione (GSH) in this pathway can also scavenge superoxide and hydrogen peroxide.During nodule senescence, several ultrastructural alterations in the nodule tissues and cells have been observed (Lucas et al., 1998; Hernández-Jiménez et al., 2002; Puppo et al., 2005, and refs. therein; Van de Velde et al., 2006). Cytosol becomes electron dense, altered vesicles proliferate, and eventually the cytosol undergoes lysis. The number of peroxisomes increases, mitochondria form complex elongated structures, and symbiosomes change in size and shape and fuse during natural and induced senescence of nodules (Hernández-Jiménez et al., 2002). Damage of the symbiosome membrane is also detected (Puppo et al., 2005; Van de Velde et al., 2006).A strategy of delayed nodule senescence could lead to increased nitrogen fixation and legume productivity. Delayed nodule senescence together with enhanced sustainability under field conditions are among the key aims of legume improvement programs (Puppo et al., 2005). An interesting approach proposed to achieve delayed senescence is to induce nodulation in legumes using rhizobial strains with modified redox capacity (Zahran, 2001).The protein flavodoxin contains a FMN group acting as a redox center transferring electrons at low potentials (Pueyo et al., 1991; Pueyo and Gómez-Moreno, 1991). The FMN cofactor of flavodoxin can exist in three different redox states: oxidized, one-electron-reduced semiquinone, and two-electron-reduced hydroquinone. This property confers high versatility to flavodoxins in electron transport systems (Simondsen and Tollin, 1980; McIver et al., 1998). To date, flavodoxin has not been described in plants, as flavodoxin-encoding genes were lost during the transition of algae to plants (Zurbriggen et al., 2007) and, consequently, no homologs have been identified in the sequenced genome of Arabidopsis (Arabidopsis thaliana; Arabidopsis Genome Initiative, 2000). Flavodoxin is present as a constitutive or inducible protein in different microorganisms (Klugkist et al., 1986). In the nitrogen-fixing cyanobacterium Anabaena variabilis PCC 7119, flavodoxin is expressed under conditions of limited iron availability, replacing ferredoxin in the photosynthetic electron transport from PSI to NADP⁺ and in nitrogenase reduction (Sandmann et al., 1990). Reversible electron transfer from flavodoxin to NADP⁺ is catalyzed by ferredoxin NADP⁺ reductase in different pathways of oxidative metabolism (Arakaki et al., 1997). In its reduced state, flavodoxin might be able to react with ROS and revert to its original redox state in the presence of an appropriate electron source. This could probably occur without the associated molecular damage that metallic complexes in catalases or SODs suffer (Keyer et al., 1995). The presence of flavodoxin has not been documented to date in the symbiotic bacterium Sinorhizobium meliloti. In Escherichia coli, however, flavodoxin induction is linked to the oxidative stress-responsive regulon soxRS (Zheng et al., 1999). It has been suggested that flavodoxin and ferredoxin (flavodoxin) NADP⁺ reductase might be induced and have a role in reestablishing the cell redox balance under oxidative stress conditions (Liochev et al., 1994). The properties of flavodoxin suggest that its presence in the cell may have a facilitating effect on ROS detoxification. In fact, an increase in the amount of flavodoxin has been observed in some bacterial species subjected to oxidative stress (Zheng et al., 1999; Yousef et al., 2003; Singh et al., 2004), and transgenic tobacco (Nicotiana tabacum) plants expressing flavodoxin in chloroplasts show enhanced tolerance to a broad range of stresses related to oxidative damage (Tognetti et al., 2006, 2007a, 2007b).In this work, Sinorhizobium meliloti was transformed with the A. variabilis flavodoxin gene and used to nodulate alfalfa (Medicago sativa) plants. The effects of flavodoxin expression on nodulation dynamics, on nodule development and senescence processes, and on nitrogen-fixing activity were analyzed. Mechanistic insights suggesting putative roles for flavodoxin in protection from ROS and the induced delay of nodule senescence are likewise discussed.     

Differential Impact of Lipoxygenase 2 and Jasmonates on Natural and Stress-Induced Senescence in Arabidopsis

Jasmonic acid and related oxylipins are controversially discussed to be involved in regulating the initiation and progression of leaf senescence. To this end, we analyzed profiles of free and esterified oxylipins during natural senescence and upon induction of senescence-like phenotypes by dark treatment and flotation on sorbitol in Arabidopsis (Arabidopsis thaliana). Jasmonic acid and free 12-oxo-phytodienoic acid increased during all three processes, with the strongest increase of jasmonic acid after dark treatment. Arabidopside content only increased considerably in response to sorbitol treatment. Monogalactosyldiacylglycerols and digalactosyldiacylglycerols decreased during these treatments and aging. Lipoxygenase 2-RNA interference (RNAi) plants were generated, which constitutively produce jasmonic acid and 12-oxo-phytodienoic acid but do not exhibit accumulation during natural senescence or upon stress treatment. Chlorophyll loss during aging and upon dark incubation was not altered, suggesting that these oxylipins are not involved in these processes. In contrast, lipoxygenase 2-RNAi lines and the allene oxid synthase-deficient mutant dde2 were less sensitive to sorbitol than the wild type, indicating that oxylipins contribute to the response to sorbitol stress.Senescence is an important, highly regulated process at the end of development. Senescence is characterized by breakdown of organelles and molecules, export and transport of these nutrients to other organs/parts of the organism, and finally programmed cell death of the senescing organ.The process of senescence has been intensively studied in leaves, and morphological as well as molecular changes in senescing leaves have been described. Yellowing as a consequence of chlorophyll and chloroplast degradation is the most obvious process during natural leaf senescence. In addition, gene expression changes dramatically during senescence. Some senescence-associated genes (SAG, SEN) have been reported that are induced during this process, and several of the encoded proteins function in macromolecule degradation, detoxification and defense metabolism, or signal transduction (Gepstein et al., 2003). Based on the degradation of chloroplasts and macromolecules, leaf metabolism changes from carbon assimilation to catabolism (Lim et al., 2007).The initiation and progression of senescence is regulated by endogenous as well as exogenous factors. Among the endogenous factors, the developmental status of the organ and of the whole plant (e.g. age and progress in flowering and seed production) has a great impact on the process of senescence. Different stress factors such as pathogen attack, drought, osmotic stress, heat, cold, ozone, UV light, and shading can induce or accelerate senescence (Quirino et al., 2000). Phytohormones are very important regulators that integrate information about the developmental status and the environmental factors. Cytokinins are antagonistic signals and delay senescence. Endogenous levels of cytokinins decrease during senescence, and exogenous application and transgenic approaches, enhancing endogenous levels of these compounds, lead to delayed senescence (Gan and Amasino, 1995). In contrast, the gaseous phytohormone ethylene is known to induce and accelerate senescence (John et al., 1995). There are also several indications that abscisic acid modulates senescence (van der Graaff et al., 2006). The roles of other phytohormones/signaling compounds such as auxin, salicylic acid, and jasmonates are less clear (Lim et al., 2007).Jasmonates are oxylipin signaling molecules derived from linolenic acid. The term jasmonates comprises 12-oxo-phytodienoic acid (OPDA), jasmonic acid (JA), and derivatives such as the methyl ester and amino acid conjugates of JA. One of the first biological activities described for these compounds was the promotion of senescence in oat (Avena sativa) leaves by methyl jasmonate (MeJa) isolated from Artemisia absinthium (Ueda and Kato, 1980). Later on, the induction of senescence-like phenotypes by exogenous application of MeJa was also found in other plant species (Ueda and Kato, 1980; Weidhase et al., 1987a; He et al., 2002). On the molecular level, this senescence-promoting effect of MeJa is accompanied by chlorophyll loss and decreases in Rubisco and photosynthesis (Weidhase et al., 1987a, 1987b). In addition, expression of some senescence-up-regulated genes is also responsive to JA; examples are SEN1, SEN4, SEN5, SAG12, SAG14, and SAG15 (Park et al., 1998; Schenk et al., 2000; He et al., 2002). Due to the results described above, jasmonates have been described for decades as compounds with senescence-promoting activities, while the function of these compounds in natural senescence in planta was critically discussed (Parthier, 1990; Sembdner and Parthier, 1993; Creelman and Mullet, 1997; Wasternack, 2007; Balbi and Devoto, 2008; Reinbothe et al., 2009). Additional indications for a role of jasmonates in regulating senescence are the transient up-regulation of expression of some enzymes involved in JA biosynthesis, such as allene oxide synthase (AOS) and OPDA reductase 3 (OPR3), and the increase in JA levels during natural senescence (He et al., 2002; van der Graaff et al., 2006). Furthermore, alterations in natural and induced senescence have been reported for some mutants with defects in the JA pathway. The mutant coi1, which is impaired in JA signaling, exhibited delayed chlorophyll loss upon dark incubation of detached leaves (Castillo and Leon, 2008). Plants with reduced expression of the 3-ketoacyl-CoA-thiolase KAT2, which is involved in β-oxidation and JA production, showed delayed yellowing during natural senescence and upon dark incubation of detached leaves (Castillo and Leon, 2008).However, there are also several reports that cast doubt on an important function of JA in senescence. For most mutants in JA biosynthesis or signaling, no differences in natural senescence are apparent (He et al., 2002; Schommer et al., 2008). In addition, mutants defective in the expression of AOS or OPR3 do not show altered senescence-like phenotypes upon dark treatment (Schommer et al., 2008; Kunz et al., 2009). It has to be taken into consideration that the knockout in these mutants has pleiotrophic effects during whole plant development. For example, the leaves of plants with reduced expression of the lipase DGL or of OPR3 are larger (Hyun et al., 2008). In addition, several knockout mutants defective in JA biosynthesis or signaling do not produce fertile flowers (Feys et al., 1994; McConn and Browse, 1996; Sanders et al., 2000; Stintzi and Browse, 2000; Ishiguro et al., 2001; von Malek et al., 2002). These changes in development might affect other developmental processes such as senescence.To investigate the function of jasmonates in senescence in more detail, we compared the oxylipin profile of wild-type leaves during natural senescence and upon stress induction of senescence-like phenotypes. The analysis of lipoxygenase 2 (LOX2)-RNA interference (RNAi) plants, which produce low basal levels of oxylipins but are impaired in the accumulation of OPDA and JA during senescence or in response to stress, indicates that 13-LOX products are not necessary for natural senescence or dark-induced chlorophyll loss but are involved in the response to sorbitol.     

Autophagy Plays a Role in Chloroplast Degradation during Senescence in Individually Darkened Leaves

Chloroplasts contain approximately 80% of total leaf nitrogen and represent a major source of recycled nitrogen during leaf senescence. While bulk degradation of the cytosol and organelles in plants is mediated by autophagy, its role in chloroplast catabolism is largely unknown. We investigated the effects of autophagy disruption on the number and size of chloroplasts during senescence. When leaves were individually darkened, senescence was promoted similarly in both wild-type Arabidopsis (Arabidopsis thaliana) and in an autophagy-defective mutant, atg4a4b-1. The number and size of chloroplasts decreased in darkened leaves of wild type, while the number remained constant and the size decrease was suppressed in atg4a4b-1. When leaves of transgenic plants expressing stroma-targeted DsRed were individually darkened, a large accumulation of fluorescence in the vacuolar lumen was observed. Chloroplasts exhibiting chlorophyll fluorescence, as well as Rubisco-containing bodies, were also observed in the vacuole. No accumulation of stroma-targeted DsRed, chloroplasts, or Rubisco-containing bodies was observed in the vacuoles of the autophagy-defective mutant. We have succeeded in demonstrating chloroplast autophagy in living cells and provide direct evidence of chloroplast transportation into the vacuole.Chloroplasts contain 75% to 80% of total leaf nitrogen mainly as proteins (Makino and Osmond, 1991). During leaf senescence, chloroplast proteins are gradually degraded as a major source of nitrogen for new growth (Wittenbach, 1978; Friedrich and Huffaker, 1980; Mae et al., 1984), correlating with a decline in photosynthetic activity, while chloroplasts gradually shrink and transform into gerontoplasts, characterized by the disintegration of the thylakoid membranes and accumulation of plastoglobuli (for a recent review, see Krupinska, 2006). Concomitantly, a decline in the cellular population of chloroplasts is also evident in many cases, for example, during natural (Kura-Hotta et al., 1990; Inada et al., 1998), dark-induced (Wittenbach et al., 1982), and nutrient-limited senescence (Mae et al., 1984; Ono et al., 1995), suggesting the existence of a whole chloroplast degradation system. Some electron microscopic studies have shown whole chloroplasts in the central vacuole, which is rich in lytic hydrolases (Wittenbach et al., 1982; Minamikawa et al., 2001). However, there is no direct evidence of chloroplasts moving into the vacuole in living cells and the mechanism of transport is not yet understood (Hörtensteiner and Feller, 2002; Krupinska, 2006).The most abundant chloroplast protein is Rubisco (EC 4.1.1.39), comprising approximately 50% of the soluble protein (Wittenbach, 1978). The amount of Rubisco decreases rapidly in the early phase of leaf senescence, although more slowly in the later phase (Friedrich and Huffaker, 1980; Mae et al., 1984). In contrast, the chloroplast number remains relatively constant, making it impossible to explain Rubisco loss solely by whole chloroplast degradation. However, the mechanism of intrachloroplastic Rubisco degradation is still unknown (for review, see Feller et al., 2008). Using immunoelectron microscopy, we previously demonstrated in naturally senescing wheat (Triticum aestivum) leaves that Rubisco is released from chloroplasts into the cytoplasm and transported to the vacuole for subsequent degradation in small spherical bodies, named Rubisco-containing bodies (RCBs; Chiba et al., 2003). Similar chloroplast-derived structures were also subsequently confirmed in senescent leaves of soybean (Glycine max) and/or Arabidopsis (Arabidopsis thaliana) by electron microscopy (Otegui et al., 2005), and recently in tobacco (Nicotiana tabacum) leaves by immunoelectron microscopy, although the authors gave them a different name, Rubisco vesicular bodies (Prins et al., 2008). RCBs have double membranes, which seem to be derived from the chloroplast envelope; thus, the RCB-mediated degradation of stromal proteins represents a potential mechanism for chloroplast shrinkage during senescence. We recently demonstrated that Rubisco and stroma-targeted fluorescent proteins can be mobilized to the vacuole by ATG-dependent autophagy via RCBs, using leaves treated with concanamycin A, a vacuolar H⁺-ATPase inhibitor (Ishida et al., 2008). To investigate further, we wished to observe chloroplast autophagy and degradation directly in living cells to determine whether autophagy is responsible for chloroplast shrinkage and whether it is involved in the vacuolar degradation of whole chloroplasts during leaf senescence.Autophagy is known to be a major system for the bulk degradation of intracellular proteins and organelles in the vacuole in yeast and plants, or the lysosome in animals (for detailed mechanisms, see reviews by Ohsumi, 2001; Levine and Klionsky, 2004; Thompson and Vierstra, 2005; Bassham et al., 2006). In those systems, a portion of the cytoplasm, including entire organelles, is engulfed in membrane-bound vesicles and delivered to the vacuole/lysosome. A recent genome-wide search confirmed that Arabidopsis has many genes homologous to the yeast autophagy genes (ATGs; Doelling et al., 2002; Hanaoka et al., 2002; for detailed functions of ATGs, see the reviews noted above). Using knockout mutants of ATGs and a monitoring system with an autophagy marker, GFP-ATG8, numerous studies have demonstrated the presence of the autophagy system in plants and its importance in several biological processes (Yoshimoto et al., 2004; Liu et al., 2005; Suzuki et al., 2005; Thompson et al., 2005; Xiong et al., 2005, 2007; Fujiki et al., 2007; Phillips et al., 2008). These articles suggest that autophagy plays an important role in nutrient recycling during senescence, especially in nutrient-starved plants. The atg mutants exhibited an accelerated loss of some chloroplast proteins, but not all, under nutrient-starved conditions and during senescence, suggesting that autophagy is not the sole mechanism for the degradation of chloroplast proteins; other, as yet unidentified systems must be responsible for the degradation of chloroplast contents when the ATG system is compromised (Levine and Klionsky, 2004; Bassham et al., 2006). However, it still remains likely that autophagy is responsible for the vacuolar degradation of chloroplasts in wild-type plants.Prolonged observation is generally required to follow leaf senescence events in naturally aging leaves and senescence-associated processes tend to become chaotic over time. To observe chloroplast degradation over a short period, and to draw clear conclusions, a suitable experimental model of leaf senescence is required. Weaver and Amasino (2001) reported that senescence is rapidly induced in individually darkened leaves (IDLs) of Arabidopsis, but retarded in plants subjected to full darkness. In addition, Keech et al. (2007) observed a significant decrease of both the number and size of chloroplasts in IDLs within 6 d.In this study, using IDLs as a senescence model, we aimed to investigate the involvement of autophagy in chloroplast degradation. We show direct evidence for the transport of whole chloroplasts and RCBs to the vacuole by autophagy.     

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.     

Nontransgenic Genome Modification in Plant Cells

Zinc finger nucleases (ZFNs) are a powerful tool for genome editing in eukaryotic cells. ZFNs have been used for targeted mutagenesis in model and crop species. In animal and human cells, transient ZFN expression is often achieved by direct gene transfer into the target cells. Stable transformation, however, is the preferred method for gene expression in plant species, and ZFN-expressing transgenic plants have been used for recovery of mutants that are likely to be classified as transgenic due to the use of direct gene-transfer methods into the target cells. Here we present an alternative, nontransgenic approach for ZFN delivery and production of mutant plants using a novel Tobacco rattle virus (TRV)-based expression system for indirect transient delivery of ZFNs into a variety of tissues and cells of intact plants. TRV systemically infected its hosts and virus ZFN-mediated targeted mutagenesis could be clearly observed in newly developed infected tissues as measured by activation of a mutated reporter transgene in tobacco (Nicotiana tabacum) and petunia (Petunia hybrida) plants. The ability of TRV to move to developing buds and regenerating tissues enabled recovery of mutated tobacco and petunia plants. Sequence analysis and transmission of the mutations to the next generation confirmed the stability of the ZFN-induced genetic changes. Because TRV is an RNA virus that can infect a wide range of plant species, it provides a viable alternative to the production of ZFN-mediated mutants while avoiding the use of direct plant-transformation methods.Methods for genome editing in plant cells have fallen behind the remarkable progress made in whole-genome sequencing projects. The availability of reliable and efficient methods for genome editing would foster gene discovery and functional gene analyses in model plants and the introduction of novel traits in agriculturally important species (Puchta, 2002; Hanin and Paszkowski, 2003; Reiss, 2003; Porteus, 2009). Genome editing in various species is typically achieved by integrating foreign DNA molecules into the target genome by homologous recombination (HR). Genome editing by HR is routine in yeast (Saccharomyces cerevisiae) cells (Scherer and Davis, 1979) and has been adapted for other species, including Drosophila, human cell lines, various fungal species, and mouse embryonic stem cells (Baribault and Kemler, 1989; Venken and Bellen, 2005; Porteus, 2007; Hall et al., 2009; Laible and Alonso-González, 2009; Tenzen et al., 2009). In plants, however, foreign DNA molecules, which are typically delivered by direct gene-transfer methods (e.g. Agrobacterium and microbombardment of plasmid DNA), often integrate into the target cell genome via nonhomologous end joining (NHEJ) and not HR (Ray and Langer, 2002; Britt and May, 2003).Various methods have been developed to indentify and select for rare site-specific foreign DNA integration events or to enhance the rate of HR-mediated DNA integration in plant cells. Novel T-DNA molecules designed to support strong positive- and negative-selection schemes (e.g. Thykjaer et al., 1997; Terada et al., 2002), altering the plant DNA-repair machinery by expressing yeast chromatin remodeling protein (Shaked et al., 2005), and PCR screening of large numbers of transgenic plants (Kempin et al., 1997; Hanin et al., 2001) are just a few of the experimental approaches used to achieve HR-mediated gene targeting in plant species. While successful, these approaches, and others, have resulted in only a limited number of reports describing the successful implementation of HR-mediated gene targeting of native and transgenic sequences in plant cells (for review, see Puchta, 2002; Hanin and Paszkowski, 2003; Reiss, 2003; Porteus, 2009; Weinthal et al., 2010).HR-mediated gene targeting can potentially be enhanced by the induction of genomic double-strand breaks (DSBs). In their pioneering studies, Puchta et al. (1993, 1996) showed that DSB induction by the naturally occurring rare-cutting restriction enzyme I-SceI leads to enhanced HR-mediated DNA repair in plants. Expression of I-SceI and another rare-cutting restriction enzyme (I-CeuI) also led to efficient NHEJ-mediated site-specific mutagenesis and integration of foreign DNA molecules in plants (Salomon and Puchta, 1998; Chilton and Que, 2003; Tzfira et al., 2003). Naturally occurring rare-cutting restriction enzymes thus hold great promise as a tool for genome editing in plant cells (Carroll, 2004; Pâques and Duchateau, 2007). However, their wide application is hindered by the tedious and next to impossible reengineering of such enzymes for novel DNA-target specificities (Pâques and Duchateau, 2007).A viable alternative to the use of rare-cutting restriction enzymes is the zinc finger nucleases (ZFNs), which have been used for genome editing in a wide range of eukaryotic species, including plants (e.g. Bibikova et al., 2001; Porteus and Baltimore, 2003; Lloyd et al., 2005; Urnov et al., 2005; Wright et al., 2005; Beumer et al., 2006; Moehle et al., 2007; Santiago et al., 2008; Shukla et al., 2009; Tovkach et al., 2009; Townsend et al., 2009; Osakabe et al., 2010; Petolino et al., 2010; Zhang et al., 2010). Here too, ZFNs have been used to enhance DNA integration via HR (e.g. Shukla et al., 2009; Townsend et al., 2009) and as an efficient tool for the induction of site-specific mutagenesis (e.g. Lloyd et al., 2005; Zhang et al., 2010) in plant species. The latter is more efficient and simpler to implement in plants as it does not require codelivery of both ZFN-expressing and donor DNA molecules and it relies on NHEJ—the dominant DNA-repair machinery in most plant species (Ray and Langer, 2002; Britt and May, 2003).ZFNs are artificial restriction enzymes composed of a fusion between an artificial Cys₂His₂ zinc-finger protein DNA-binding domain and the cleavage domain of the FokI endonuclease. The DNA-binding domain of ZFNs can be engineered to recognize a variety of DNA sequences (for review, see Durai et al., 2005; Porteus and Carroll, 2005; Carroll et al., 2006). The FokI endonuclease domain functions as a dimer, and digestion of the target DNA requires proper alignment of two ZFN monomers at the target site (Durai et al., 2005; Porteus and Carroll, 2005; Carroll et al., 2006). Efficient and coordinated expression of both monomers is thus required for the production of DSBs in living cells. Transient ZFN expression, by direct gene delivery, is the method of choice for targeted mutagenesis in human and animal cells (e.g. Urnov et al., 2005; Beumer et al., 2006; Meng et al., 2008). Among the different methods used for high and efficient transient ZFN delivery in animal and human cell lines are plasmid injection (Morton et al., 2006; Foley et al., 2009), direct plasmid transfer (Urnov et al., 2005), the use of integrase-defective lentiviral vectors (Lombardo et al., 2007), and mRNA injection (Takasu et al., 2010).In plant species, however, efficient and strong gene expression is often achieved by stable gene transformation. Both transient and stable ZFN expression have been used in gene-targeting experiments in plants (Lloyd et al., 2005; Wright et al., 2005; Maeder et al., 2008; Cai et al., 2009; de Pater et al., 2009; Shukla et al., 2009; Tovkach et al., 2009; Townsend et al., 2009; Osakabe et al., 2010; Petolino et al., 2010; Zhang et al., 2010). In all cases, direct gene-transformation methods, using polyethylene glycol, silicon carbide whiskers, or Agrobacterium, were deployed. Thus, while mutant plants and tissues could be recovered, potentially without any detectable traces of foreign DNA, such plants were generated using a transgenic approach and are therefore still likely to be classified as transgenic. Furthermore, the recovery of mutants in many cases is also dependent on the ability to regenerate plants from protoplasts, a procedure that has only been successfully applied in a limited number of plant species. Therefore, while ZFN technology is a powerful tool for site-specific mutagenesis, its wider implementation for plant improvement may be somewhat limited, both by its restriction to certain plant species and by legislative restrictions imposed on transgenic plants.Here we describe an alternative to direct gene transfer for ZFN delivery and for the production of mutated plants. Our approach is based on the use of a novel Tobacco rattle virus (TRV)-based expression system, which is capable of systemically infecting its host and spreading into a variety of tissues and cells of intact plants, including developing buds and regenerating tissues. We traced the indirect ZFN delivery in infected plants by activation of a mutated reporter gene and we demonstrate that this approach can be used to recover mutated plants.     

Patterns of Metabolite Changes Identified from Large-Scale Gene Perturbations in Arabidopsis Using a Genome-Scale Metabolic Network

Metabolomics enables quantitative evaluation of metabolic changes caused by genetic or environmental perturbations. However, little is known about how perturbing a single gene changes the metabolic system as a whole and which network and functional properties are involved in this response. To answer this question, we investigated the metabolite profiles from 136 mutants with single gene perturbations of functionally diverse Arabidopsis (Arabidopsis thaliana) genes. Fewer than 10 metabolites were changed significantly relative to the wild type in most of the mutants, indicating that the metabolic network was robust to perturbations of single metabolic genes. These changed metabolites were closer to each other in a genome-scale metabolic network than expected by chance, supporting the notion that the genetic perturbations changed the network more locally than globally. Surprisingly, the changed metabolites were close to the perturbed reactions in only 30% of the mutants of the well-characterized genes. To determine the factors that contributed to the distance between the observed metabolic changes and the perturbation site in the network, we examined nine network and functional properties of the perturbed genes. Only the isozyme number affected the distance between the perturbed reactions and changed metabolites. This study revealed patterns of metabolic changes from large-scale gene perturbations and relationships between characteristics of the perturbed genes and metabolic changes.Rational and quantitative assessment of metabolic changes in response to genetic modification (GM) is an open question and in need of innovative solutions. Nontargeted metabolite profiling can detect thousands of compounds, but it is not easy to understand the significance of the changed metabolites in the biochemical and biological context of the organism. To better assess the changes in metabolites from nontargeted metabolomics studies, it is important to examine the changed metabolites in the context of the genome-scale metabolic network of the organism.Metabolomics is a technique that aims to quantify all the metabolites in a biological system (Nikolau and Wurtele, 2007; Nicholson and Lindon, 2008; Roessner and Bowne, 2009). It has been used widely in studies ranging from disease diagnosis (Holmes et al., 2008; DeBerardinis and Thompson, 2012) and drug discovery (Cascante et al., 2002; Kell, 2006) to metabolic reconstruction (Feist et al., 2009; Kim et al., 2012) and metabolic engineering (Keasling, 2010; Lee et al., 2011). Metabolomic studies have demonstrated the possibility of identifying gene functions from changes in the relative concentrations of metabolites (metabotypes or metabolic signatures; Ebbels et al., 2004) in various species including yeast (Saccharomyces cerevisiae; Raamsdonk et al., 2001; Allen et al., 2003), Arabidopsis (Arabidopsis thaliana; Brotman et al., 2011), tomato (Solanum lycopersicum; Schauer et al., 2006), and maize (Zea mays; Riedelsheimer et al., 2012). Metabolomics has also been used to better understand how plants interact with their environments (Field and Lake, 2011), including their responses to biotic and abiotic stresses (Dixon et al., 2006; Arbona et al., 2013), and to predict important agronomic traits (Riedelsheimer et al., 2012). Metabolite profiling has been performed on many plant species, including angiosperms such as Arabidopsis, poplar (Populus trichocarpa), and Catharanthus roseus (Sumner et al., 2003; Rischer et al., 2006), basal land plants such as Selaginella moellendorffii and Physcomitrella patens (Erxleben et al., 2012; Yobi et al., 2012), and Chlamydomonas reinhardtii (Fernie et al., 2012; Davis et al., 2013). With the availability of whole genome sequences of various species, metabolomics has the potential to become a useful tool for elucidating the functions of genes using large-scale systematic analyses (Fiehn et al., 2000; Saito and Matsuda, 2010; Hur et al., 2013).Although metabolomics data have the potential for identifying the roles of genes that are associated with metabolic phenotypes, the biochemical mechanisms that link functions of genes with metabolic phenotypes are still poorly characterized. For example, we do not yet know the principles behind how perturbing the expression of a single gene changes the metabolic system as a whole. Large-scale metabolomics data have provided useful resources for linking phenotypes to genotypes (Fiehn et al., 2000; Roessner et al., 2001; Tikunov et al., 2005; Schauer et al., 2006; Lu et al., 2011; Fukushima et al., 2014). For example, Lu et al. (2011) compared morphological and metabolic phenotypes from more than 5,000 Arabidopsis chloroplast mutants using gas chromatography (GC)- and liquid chromatography (LC)-mass spectrometry (MS). Fukushima et al. (2014) generated metabolite profiles from various characterized and uncharacterized mutant plants and clustered the mutants with similar metabolic phenotypes by conducting multidimensional scaling with quantified metabolic phenotypes. Nonetheless, representation and analysis of such a large amount of data remains a challenge for scientific discovery (Lu et al., 2011). In addition, these studies do not examine the topological and functional characteristics of metabolic changes in the context of a genome-scale metabolic network. To understand the relationship between genotype and metabolic phenotype, we need to investigate the metabolic changes caused by perturbing the expression of a gene in a genome-scale metabolic network perspective, because metabolic pathways are not independent biochemical factories but are components of a complex network (Berg et al., 2002; Merico et al., 2009).Much progress has been made in the last 2 decades to represent metabolism at a genome scale (Terzer et al., 2009). The advances in genome sequencing and emerging fields such as biocuration and bioinformatics enabled the representation of genome-scale metabolic network reconstructions for model organisms (Bassel et al., 2012). Genome-scale metabolic models have been built and applied broadly from microbes to plants. The first step toward modeling a genome-scale metabolism in a plant species started with developing a genome-scale metabolic pathway database for Arabidopsis (AraCyc; Mueller et al., 2003) from reference pathway databases (Kanehisa and Goto, 2000; Karp et al., 2002; Zhang et al., 2010). Genome-scale metabolic pathway databases have been built for several plant species (Mueller et al., 2005; Zhang et al., 2005, 2010; Urbanczyk-Wochniak and Sumner, 2007; May et al., 2009; Dharmawardhana et al., 2013; Monaco et al., 2013, 2014; Van Moerkercke et al., 2013; Chae et al., 2014; Jung et al., 2014). Efforts have been made to develop predictive genome-scale metabolic models using enzyme kinetics and stoichiometric flux-balance approaches (Sweetlove et al., 2008). de Oliveira Dal’Molin et al. (2010) developed a genome-scale metabolic model for Arabidopsis and successfully validated the model by predicting the classical photorespiratory cycle as well as known key differences between redox metabolism in photosynthetic and nonphotosynthetic plant cells. Other genome-scale models have been developed for Arabidopsis (Poolman et al., 2009; Radrich et al., 2010; Mintz-Oron et al., 2012), C. reinhardtii (Chang et al., 2011; Dal’Molin et al., 2011), maize (Dal’Molin et al., 2010; Saha et al., 2011), sorghum (Sorghum bicolor; Dal’Molin et al., 2010), and sugarcane (Saccharum officinarum; Dal’Molin et al., 2010). These predictive models have the potential to be applied broadly in fields such as metabolic engineering, drug target discovery, identification of gene function, study of evolutionary processes, risk assessment of genetically modified crops, and interpretations of mutant phenotypes (Feist and Palsson, 2008; Ricroch et al., 2011).Here, we interrogate the metabotypes caused by 136 single gene perturbations of Arabidopsis by analyzing the relative concentration changes of 1,348 chemically identified metabolites using a reconstructed genome-scale metabolic network. We examine the characteristics of the changed metabolites (the metabolites whose relative concentrations were significantly different in mutants relative to the wild type) in the metabolic network to uncover biological and topological consequences of the perturbed genes.     

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.     

Integrating Image-Based Phenomics and Association Analysis to Dissect the Genetic Architecture of Temporal Salinity Responses in Rice

Salinity affects a significant portion of arable land and is particularly detrimental for irrigated agriculture, which provides one-third of the global food supply. Rice (Oryza sativa), the most important food crop, is salt sensitive. The genetic resources for salt tolerance in rice germplasm exist but are underutilized due to the difficulty in capturing the dynamic nature of physiological responses to salt stress. The genetic basis of these physiological responses is predicted to be polygenic. In an effort to address this challenge, we generated temporal imaging data from 378 diverse rice genotypes across 14 d of 90 mm NaCl stress and developed a statistical model to assess the genetic architecture of dynamic salinity-induced growth responses in rice germplasm. A genomic region on chromosome 3 was strongly associated with the early growth response and was captured using visible range imaging. Fluorescence imaging identified four genomic regions linked to salinity-induced fluorescence responses. A region on chromosome 1 regulates both the fluorescence shift indicative of the longer term ionic stress and the early growth rate decline during salinity stress. We present, to our knowledge, a new approach to capture the dynamic plant responses to its environment and elucidate the genetic basis of these responses using a longitudinal genome-wide association model.Nearly one-third of the 54 million ha of the highly saline soils in the world are located in South and Southeast Asia. Rice (Oryza sativa), which is the primary source of calories and protein for these two regions, is very sensitive to salinity stress, with even moderate salinity levels known to decrease yields by 50% (Zeng et al., 2002). Projected sea level rise due to climate change is expected to increase saltwater ingress in coastal rice-growing regions of South and Southeast Asia. Therefore, development of salt-tolerant rice cultivars is essential to maintain rice productivity in the salinity-affected regions globally.Salt tolerance, defined as the ability to maintain growth and productivity in saline conditions, is a complex polygenic trait that may be influenced by distinct physiological mechanisms (Munns et al., 1982; Munns and Termaat, 1986; Cheeseman, 1988; Munns and Tester, 2008; Horie et al., 2012; for a comprehensive review of genes involved in salinity tolerance in rice, see Negrão et al., 2011) At the cellular level, plants respond to saline conditions in two phases, namely an osmotic (shoot ion independent) and an ionic stress phase, which can occur in an overlapping manner with varying intensity during the course of salinity stress (Munns and Termaat, 1986; Munns, 2002; Munns and James, 2003; Munns and Tester, 2008; Horie et al., 2012). During the osmotic stress phase, which occurs soon after the onset of salinity, the reduction in external osmotic potential disrupts water uptake and impedes cell expansion, which, at the whole plant level, leads to reduced growth rate (Matsuda and Riazi, 1981; Munns and Passioura, 1984; Rawson and Munns, 1984; Azaizeh and Steudle, 1991; Fricke and Peters, 2002; Fricke, 2004; Boursiac et al., 2005). As salinity stress persists over several days and weeks, sodium ions (Na⁺) accumulate to toxic levels, resulting in cell death and precocious leaf senescence (Lutts and Bouharmont, 1996; Munns, 2002; Munns and James, 2003; Ghanem et al., 2008). This is typically observed during the ionic phase of the salinity response (Munns, 2002; Munns and James, 2003; Munns and Tester, 2008). Plants possess distinct mechanisms to adapt to these osmotic and ionic stresses that are controlled by a suite of genes (Apse et al., 1999; Carvajal et al., 1999; Halfter et al., 2000; Ishitani et al., 2000; Shi et al., 2000; Zeng and Shannon, 2000; Rus et al., 2001; Berthomieu et al., 2003; Martínez-Ballesta et al., 2003; Boursiac et al., 2005, 2008; Ren et al., 2005; Huang et al., 2006; Davenport et al., 2007; Obata et al., 2007; Székely et al., 2008; Horie et al., 2011; Rivandi et al., 2011; Asano et al., 2012; Munns et al., 2012; Latz et al., 2013; Schmidt et al., 2013; Campo et al., 2014; Choi et al., 2014; Liu et al., 2014). The genetic basis of temporal adaptive responses to salinity stress remains to be explored in rice and other crops. This is primarily due to challenges in capturing the dynamic physiological responses to salinity for a large number of genotypes in a nondestructive manner. Manual phenotyping to detect incremental changes in growth rate during the osmotic stress and slight shifts in leaf color due to ionic stress is difficult to quantify for a large number of genotypes.In rice, at least one major quantitative trait loci (QTL; saltol) for salinity tolerance has been characterized based on end point measurements of biomass, senescence/injury, and Na⁺ and K⁺ concentrations (Bonilla et al., 2002; Lin et al., 2004; Thomson et al., 2010). SHOOT K⁺ CONTENT1 (SKC1) is the causative gene underlying the saltol region. SKC1 encodes a Na⁺-selective high-affinity potassium transporter that regulates Na⁺/K⁺ homeostasis during salinity stress (Ren et al., 2005). High levels of Na⁺ displace cellular K⁺, an essential element for several enzymatic reactions and physiological processes (Gierth and Mäser, 2007). The ability to maintain cellular K⁺ levels during salinity through the action of Na⁺-selective potassium transporters or Na⁺/H⁺ antiporters is a well-characterized tolerance mechanism in cereals including rice (Ren et al., 2005; Sunarpi et al., 2005; Huang et al., 2006; Møller et al., 2009; Mian et al., 2011; Munns et al., 2012).Numerous studies have utilized conventional linkage mapping to identify QTL for morphological and physiological responses to salinity in rice using discrete end point measurements (Bonilla et al., 2002; Lin et al., 2004; Ren et al., 2005; Negrão et al., 2011; Wang et al., 2012). However, the physiological adaptation to saline conditions is a complex continuous process that develops over time. While some accessions will exhibit similar end point phenotypic values, the genetic and physiological mechanisms giving rise to the similar phenotypes may be very different and the growth trajectories throughout the experiment may be distinct. Although single time point studies have yielded important information regarding the genetic basis of salinity tolerance, such approaches are too simple to reveal the genetic architecture of stress adaptation. With the advent of high-throughput image-based phenotyping platforms, it is now feasible to quantify dynamic responses during the stress treatment for a large number of genotypes (Berger et al., 2010; Golzarian et al., 2011; Chen et al., 2014; Honsdorf et al., 2014).Image-based phenotyping has been combined with genome-wide association studies (GWAS) and linkage mapping to examine the genetic basis of complex developmental processes (Busemeyer et al., 2013; Moore et al., 2013; Topp et al., 2013; Slovak et al., 2014; Würschum et al., 2014; Yang et al., 2014; Bac-Molenaar et al., 2015). Moreover, the introduction of the time axis provides a better understanding of the physiological processes underlying complex stress and developmental responses compared with single end point measurements (Zhang et al., 2012; Moore et al., 2013; Brown et al., 2014; Chen et al., 2014; Slovak et al., 2014; Bac-Molenaar et al., 2015). However, to date, no studies have implemented an association mapping approach using image-derived phenotypes to address the genetic basis of dynamic stress responses in plants. Image-based phenotyping offers several advantages over conventional phenotyping: (1) quantitative measurements can be recorded over discrete time points to capture morphological and physiological responses in a nondestructive manner, and (2) the use of various types of spectral imaging address phenotypes that are not detectable to the human eye such as chlorophyll fluorescence and leaf water content. Integrating dynamic phenotypic data and association mapping has the potential to query genetic diversity across hundreds of accessions for complex traits and provides much higher resolution compared with conventional linkage mapping. Here, we explored the dynamic growth and chlorophyll responses to salinity of a diverse set of rice accessions using high-throughput visible and fluorescence imaging. To assess the genetic basis of plant growth in saline conditions, a logistic model was used to describe the temporal growth responses and was incorporated into the statistical framework necessary for association mapping. Coupled with temporal fluorescence imaging, we present, to our knowledge, new insights into the genetic architecture of osmotic and ionic responses during salinity stress in rice.