Deciphering the Physalis floridana Double-Layered-Lantern1 Mutant Provides Insights into Functional Divergence of the GLOBOSA Duplicates within the Solanaceae

Glycosylphosphatidylinositol-anchor biosynthesis and glycosylphosphatidylinositol modification of proteins are central to coordinated plant development.Since their discovery (Low and Saltiel, 1988), glycosylphosphatidylinositol-anchored proteins (GPI-APs) have provoked intense interest as crucial regulators for growth, morphogenesis, reproduction, and disease pathogenesis in organisms ranging from yeast and trypanosomes to animals and plants. The lipid moiety, the glycosylphosphatidylinositol (GPI) anchor, is synthesized in the endoplasmic reticulum (ER); the protein component is cotranslationally inserted into the ER and posttranslationally modified by the addition of a GPI anchor (Kinoshita et al., 2013; Fig. 1). GPI-APs are then transported via the Golgi to the outer surface of the plasma membrane. The lipid anchor mediates stable attachment of these proteins to the cell surface, where some play important roles as signaling regulators from sphingolipid- and sterol-enriched membrane microdomains (Simons and Gerl, 2010). Some GPI-APs are released from the cell membrane by phosphatidylinositol-specific phospholipases to the extracellular matrix, where they might engage in processes such as cell adhesion and cell-cell communication. In Arabidopsis (Arabidopsis thaliana), there are about 250 predicted GPI-APs (Borner et al., 2003), a relatively large number compared with about 150 in mammals and 50 in the budding yeast (Saccharomyces cerevisiae). Important functions for plant GPI-APs have been elucidated through the study of individual proteins, such as the COBRA family in cell expansion and cell wall biosynthesis (Brady et al., 2007), ARABINOGALACTAN PROTEIN18 in megagametogenesis (Demesa-Arévalo and Vielle-Calzada, 2013), and LORELEI in the pollen tube-female gametophyte interaction (Capron et al., 2008; Tsukamoto et al., 2010; Duan et al., 2014). However, it is the studies of mutants defective in GPI biosynthesis that underscore the general importance of GPI-APs as a class: lacking the capacity to assemble the anchor is lethal.Open in a separate windowFigure 1.GPI anchor biosynthesis pathway. Ten or 11 stepwise modifications of phosphoinositide occur, starting from the synthesis of N-glucosamine-phosphoinositide on the cytoplasmic surface of the ER, followed by its flipping to the ER lumenal side for additional modifications, ending with the addition of the terminal ethanolamine phosphate. Proteins destined for GPI modification are synthesized with a C-terminal signature sequence recognized by the GPI transamidase (a five-protein-enzyme complex) that concomitantly cleaves the peptide at what is designated as the ω and ω+1 amino acids and attaches the GPI anchor in a transamination reaction (red arrows). The GPI-modified proteins are then sorted and transported via the Golgi apparatus to the cell membrane. The established biosynthetic proteins from Arabidopsis and their mammalian homologs are indicated; the galactosylation step appears to be plant specific. The diagram is modeled after figure 3 in Ellis et al. (2010), which also provided a complete list of potential plant orthologs of the human and yeast proteins in the pathway.The GPI anchor is synthesized by an elaborate biosynthetic pathway, starting on the cytoplasmic side of the ER and ending with a completely assembled core anchor on the lumenal surface of the ER (Fig. 1). Prior to their transport out of the ER, proteins destined for GPI modification are cleaved at a C-terminal signature sequence by a GPI transamidase complex that in two enzymatic steps concomitantly attaches a GPI anchor to the C terminus of processed proteins (Kinoshita, 2014). Most of the knowledge on GPI biosynthesis and GPI-AP modification is derived from studies in mammals and yeast, but the pathway is likely to be conserved in plants (Ellis et al., 2010). In a recent article in Plant Physiology, Dai et al. (2014) reported that a GPI anchor biosynthesis mutant, abnormal pollen tube guidance1 (atpg1), displays both embryo lethality and severely depressed male fertility. They determined that APTG1 is homologous to the yeast GPI10 and human PIG-B (for phosphatidylinositol glycan anchor biosynthesis) proteins, the last of three distinct mannosyltransferases that modify the precursor anchor (Fig. 1), and showed that APTG1 can functionally substitute for GPI10 in a conditionally lethal gpi10 yeast mutant. Previous studies have demonstrated that Arabidopsis SETH1 (a male fertility god in Egyptian mythology), SETH2, and PEANUT1 (PNT1), encoding homologs of mammalian PIG-C, PIG-A, and PIG-M (Fig. 1) and their corresponding yeast counterparts, respectively, are important for male fertility (Lalanne et al., 2004; Gillmor et al., 2005). In addition, loss of the first mannosyltransferase in the pathway in pnt1 results in early seedling lethality. pnt1 embryos are severely defective, displaying various cell division anomalies and exhibiting altered levels and ectopic deposition of cell wall polymers. The results reported by Dai et al. (2014), therefore, further demonstrate the conservation of the GPI biosynthesis pathway and the importance of GPI anchoring in plant development and reproduction.The aptg1 mutant was isolated in a search for mutants defective in pollen tube targeting of ovules (Fig. 2), an intriguing and crucial step in plant reproduction. A pollen tube is guided to an ovule by attractants, and upon reaching the target, the female gametophyte, the pollen tube ruptures, ejecting its cytoplasm and releasing sperm for fertilization (Dresselhaus and Franklin-Tong, 2013). aptg1 pollen tubes either fail to target ovules or undertake a more twisted pathway before entering an ovule. In an earlier study, Li et al. (2013) showed that a GPI-AP, COBRA-LIKE10 (COBL10), is required to maintain normal pollen tube growth rates and ovule targeting efficiency. In aptg1 pollen tubes, citrine fluorescent protein-COBL10 was absent from its normal apical membrane location while the citrine fluorescent signal in the cytoplasm was more intense, implying that the diminished presence of COBL10 on the apical membrane could be an underlying cause for the ovule-targeting phenotype. This observation also demonstrates that GPI anchoring is important for the subsequent sorting and transport of these proteins to their destined locations (Kinoshita et al., 2013) and consistent with a wholesale failure of GPI-APs to reach their functional locations as underlying lethality in GPI biosynthesis mutants.Open in a separate windowFigure 2.Pollen tube targeting of ovules in an Arabidopsis pistil. GUS-expressing pollen grains pollinated the pistil. Each blue dot represents discharged cytoplasm from a pollen tube that, in response to attractants, has successfully targeted the ovule and penetrated the female gametophyte and was induced to burst. The cytoplasmic discharge releases sperm for fertilization.While it is clear that major biological roles are played by GPI-APs, many questions remain. Most constituents of the plant GPI anchor biosynthetic pathway remain to be functionally established (Fig. 1). Much has been said about the membrane environments where GPI-APs are localized, but we are far from understanding the precise roles they play in assembling these domains and regulating their functional dynamics. Advances in high-resolution imaging at the cell surface and biochemical approaches to determine the constituents in these membrane microdomains (Simons and Gerl, 2010) should accelerate our understanding of the importance of GPI anchoring as a conserved strategy among eukaryotes to control a wide range of processes.     

Long-Term Evolution of Nucleotide-Binding Site-Leucine-Rich Repeat Genes: Understanding Gained from and beyond the Legume Family

Proper utilization of plant disease resistance genes requires a good understanding of their short- and long-term evolution. Here we present a comprehensive study of the long-term evolutionary history of nucleotide-binding site (NBS)-leucine-rich repeat (LRR) genes within and beyond the legume family. The small group of NBS-LRR genes with an amino-terminal RESISTANCE TO POWDERY MILDEW8 (RPW8)-like domain (referred to as RNL) was first revealed as a basal clade sister to both coiled-coil-NBS-LRR (CNL) and Toll/Interleukin1 receptor-NBS-LRR (TNL) clades. Using Arabidopsis (Arabidopsis thaliana) as an outgroup, this study explicitly recovered 31 ancestral NBS lineages (two RNL, 21 CNL, and eight TNL) that had existed in the rosid common ancestor and 119 ancestral lineages (nine RNL, 55 CNL, and 55 TNL) that had diverged in the legume common ancestor. It was shown that, during their evolution in the past 54 million years, approximately 94% (112 of 119) of the ancestral legume NBS lineages experienced deletions or significant expansions, while seven original lineages were maintained in a conservative manner. The NBS gene duplication pattern was further examined. The local tandem duplications dominated NBS gene gains in the total number of genes (more than 75%), which was not surprising. However, it was interesting from our study that ectopic duplications had created many novel NBS gene loci in individual legume genomes, which occurred at a significant frequency of 8% to 20% in different legume lineages. Finally, by surveying the legume microRNAs that can potentially regulate NBS genes, we found that the microRNA-NBS gene interaction also exhibited a gain-and-loss pattern during the legume evolution.To combat the constant challenges by pathogens, plants have evolved a sophisticated two-layered defense system, in which proteins encoded by disease RESISTANCE (R) genes serve to sense pathogen invasion signals and to elicit defense responses (Jones and Dangl, 2006; McDowell and Simon, 2006; Bent and Mackey, 2007). Over 140 R genes have been characterized from different flowering plants, which confer resistance to a large array of pathogens, including bacteria, fungi, oomycetes, viruses, and nematodes (Liu et al., 2007; Yang et al., 2013). Among these, about 80% belong to the NBS-LRR class, which encodes a central nucleotide-binding site (NBS) domain and a C-terminal leucine-rich repeat (LRR) domain. Based on whether their N termini are homologous to the Toll/Interleukin1 receptor (TIR), the angiosperm NBS-LRR genes are further divided into the TIR-NBS-LRR (TNL) subclass and the non-TIR-NBS-LRR (nTNL) subclass (Meyers et al., 1999; Bai et al., 2002; Cannon et al., 2002). The latter has been also called CC-NBS-LRR (CNL) subclass, since a coiled-coil (CC) structure is often detected at the N terminus (Meyers et al., 2003). Interestingly, a small group of nTNL genes have an N-terminal RPW8-like domain with a transmembrane region before the CC structure (Xiao et al., 2001). This group of RPW8-NBS-LRR (RNL) genes has been usually viewed as a specific lineage of CNLs (Bonardi et al., 2011; Collier et al., 2011); however, its real phylogenetic relationship with CNLs and TNLs requires further investigation.NBS-LRR genes have been surveyed in many sequenced genomes of flowering plants, including four monocots: rice (Oryza sativa), maize (Zea mays), sorghum (Sorghum bicolor), and Brachypodium distachyon; one basal eudicot: Nelumbo nucifera; two asterid species: potato (Solanum tuberosum) and tomato (Solanum lycopersicum); and 14 rosids: Vitis vinifera, Populus trichocarpa, Ricinus communis, Medicago truncatula, soybean (Glycine max), Lotus japonicus, Cucumis sativus, Cucumis melo, Citrullus lanatus, Gossypium raimondii, Carica papaya, Arabidopsis (Arabidopsis thaliana), Arabidopsis lyrata, and Brassica rapa (Bai et al., 2002; Meyers et al., 2003; Monosi et al., 2004; Zhou et al., 2004; Yang et al., 2006, 2008b; Ameline-Torregrosa et al., 2008; Mun et al., 2009; Porter et al., 2009; Chen et al., 2010; Li et al., 2010a, 2010b; Guo et al., 2011; Zhang et al., 2011; Jupe et al., 2012; Lozano et al., 2012; Luo et al., 2012; Tan and Wu, 2012; Andolfo et al., 2013; Jia et al., 2013; Lin et al., 2013; Wan et al., 2013; Wei et al., 2013; Wu et al., 2014). Variable numbers (from dozens to hundreds) of NBS-LRR genes were reported from these genomes, making one wonder: how did these genes evolve so variably during flowering plant speciation?Comparative genomic studies conducted in the available genome sequences of closely related species or subspecies revealed that a significant proportion of NBS-LRR genes are not shared. For example, 70 NBS-LRR genes between Arabidopsis and A. lyrata show the presence/absence of polymorphisms (Chen et al., 2010; Guo et al., 2011). Moreover, synteny analysis revealed that, among 363 NBS-LRR gene loci in indica (cv 93-11) and japonica (cv Nipponbare) rice, 124 loci exist in only one genome (Luo et al., 2012). Unequal crossover, homologous repair, and nonhomologous repair are the three ways that NBS-LRR gene deletions are caused in rice genomes (Luo et al., 2012).In many surveyed genomes, the majority of NBS-LRR genes are found in a clustered organization (physically close to each other), with the rest exhibited as singletons. Many clusters are homogenous, with only duplicated members occupying the same phylogenetic lineage, whereas heterogenous clusters comprise members from distantly related clades (Meyers et al., 2003). Leister (2004) defined three types of NBS gene duplications: local tandem, ectopic, and segmental duplications. Although a general agreement on the widespread occurrence of local tandem duplications can be reached by various genome survey studies, the relative importance of ectopic and segmental duplications has been seldom investigated since the earliest surveys of the Arabidopsis genome (Richly et al., 2002; Baumgarten et al., 2003; Meyers et al., 2003; McDowell and Simon, 2006).With more genomic data available in certain angiosperm families, NBS-LRR genes should be further investigated among phylogenetically distant species to fill the gaps in the understanding of their long-term evolutionary patterns. The legume family contains many economically important crop species, such as clover (Trifolium spp.), soybean, peanut (Arachis hypogaea), and common bean (Phaseolus vulgaris). Although these legumes are constantly threatened by various pathogens, only a few functional legume R genes have been characterized, and all of them belong to the NBS-LRR class (Ashfield et al., 2004; Hayes et al., 2004; Gao et al., 2005; Seo et al., 2006; Yang et al., 2008a; Meyer et al., 2009). Therefore, it would be interesting to investigate the NBS-LRR gene repertoire among different legume species. Here, we carried out a comprehensive analysis of NBS-LRR genes in four divergent legume genomes, M. truncatula, pigeon pea (Cajanus cajan), common bean, and soybean, which shared a common ancestor approximately 54 million years ago (MYA; Fig. 1; Lavin et al., 2005). Approximately 1,000 nTNL and 667 TNL subclass NBS-encoding genes were identified in our study. Their genomic distribution, organization modes, phylogenetic relationships, and syntenic patterns were examined to obtain insight into the long-term evolutionary patterns of NBS-LRR genes.Open in a separate windowFigure 1.The phylogenetic tree of four investigated legume species (M. truncatula, pigeon pea, common bean, and soybean). Two WGD events are indicated with triangles: one occurred approximately 59 MYA in the common ancestor of the four investigated legumes, and the other occurred approximately 13 MYA in the Glycine spp. lineage alone (Schmutz et al., 2010). The numbers at the branch nodes indicate divergence times (Lavin et al., 2005; Stefanovic et al., 2009). [See online article for color version of this figure.]     

Focus Issue on Calcium Signaling: Analyses of a Gravistimulation-Specific Ca2+ Signature in Arabidopsis using Parabolic Flights

Gravity is a critical environmental factor affecting the morphology and functions of organisms on the Earth. Plants sense changes in the gravity vector (gravistimulation) and regulate their growth direction accordingly. In Arabidopsis (Arabidopsis thaliana) seedlings, gravistimulation, achieved by rotating the specimens under the ambient 1g of the Earth, is known to induce a biphasic (transient and sustained) increase in cytoplasmic calcium concentration ([Ca2+]_c). However, the [Ca2+]_c increase genuinely caused by gravistimulation has not been identified because gravistimulation is generally accompanied by rotation of specimens on the ground (1g), adding an additional mechanical signal to the treatment. Here, we demonstrate a gravistimulation-specific Ca²⁺ response in Arabidopsis seedlings by separating rotation from gravistimulation by using the microgravity (less than 10⁻⁴g) conditions provided by parabolic flights. Gravistimulation without rotating the specimen caused a sustained [Ca2+]_c increase, which corresponds closely to the second sustained [Ca2+]_c increase observed in ground experiments. The [Ca2+]_c increases were analyzed under a variety of gravity intensities (e.g. 0.5g, 1.5g, or 2g) combined with rapid switching between hypergravity and microgravity, demonstrating that Arabidopsis seedlings possess a very rapid gravity-sensing mechanism linearly transducing a wide range of gravitational changes (0.5g–2g) into Ca²⁺ signals on a subsecond time scale.Calcium ion (Ca²⁺) functions as an intracellular second messenger in many signaling pathways in plants (White and Broadley, 2003; Hetherington and Brownlee, 2004; McAinsh and Pittman, 2009; Spalding and Harper, 2011). Endogenous and exogenous signals are spatiotemporally encoded by changing the free cytoplasmic concentration of Ca²⁺ ([Ca2+]_c), which in turn triggers [Ca2+]_c-dependent downstream signaling (Sanders et al., 2002; Dodd et al., 2010). A variety of [Ca2+]_c increases induced by diverse environmental and developmental stimuli are reported, such as phytohormones (Allen et al., 2000), temperature (Plieth et al., 1999; Dodd et al., 2006), and touch (Knight et al., 1991; Monshausen et al., 2009). The [Ca2+]_c increase couples each stimulus and appropriate physiological responses. In the Ca²⁺ signaling pathways, the stimulus-specific [Ca2+]_c pattern (e.g. amplitude and oscillation) provide the critical information for cellular signaling (Scrase-Field and Knight, 2003; Dodd et al., 2010). Therefore, identification of the stimulus-specific [Ca2+]_c signature is crucial for an understanding of the intracellular signaling pathways and physiological responses triggered by each stimulus, as shown in the case of cold acclimation (Knight et al., 1996; Knight and Knight, 2000).Plants often exhibit biphasic [Ca2+]_c increases in response to environmental stimuli. Thus, slow cooling causes a fast [Ca2+]_c transient followed by a second, extended [Ca2+]_c increase in Arabidopsis (Arabidopsis thaliana; Plieth et al., 1999; Knight and Knight, 2000). The Ca²⁺ channel blocker lanthanum (La³⁺) attenuated the fast transient but not the following increase (Knight and Knight, 2000), suggesting that these two [Ca2+]_c peaks have different origins. Similarly, hypoosmotic shock caused a biphasic [Ca2+]_c increase in tobacco (Nicotiana tabacum) suspension-culture cells (Takahashi et al., 1997; Cessna et al., 1998). The first [Ca2+]_c peak was inhibited by gadolinium (Gd³⁺), La³⁺, and the Ca²⁺ chelator EGTA (Takahashi et al., 1997; Cessna et al., 1998), whereas the second [Ca2+]_c increase was inhibited by the intracellular Ca²⁺ store-depleting agent caffeine but not by EGTA (Cessna et al., 1998). The amplitude of the first [Ca2+]_c peak affected the amplitude of the second increase and vice versa (Cessna et al., 1998). These results suggest that even though the two [Ca2+]_c peaks originate from different Ca²⁺ fluxes (e.g. Ca²⁺ influx through the plasma membrane and Ca²⁺ release from subcellular stores, respectively), they are closely interrelated, showing the importance of the kinetic and pharmacological analyses of these [Ca2+]_c increases.Changes in the gravity vector (gravistimulation) could work as crucial environmental stimuli in plants and are generally achieved by rotating the specimens (e.g. +180°) in ground experiments. Use of Arabidopsis seedlings expressing apoaequorin, a Ca²⁺-reporting photoprotein (Plieth and Trewavas, 2002; Toyota et al., 2008a), has revealed that gravistimulation induces a biphasic [Ca2+]_c increase that may be involved in the sensory pathway for gravity perception/response (Pickard, 2007; Toyota and Gilroy, 2013) and the intracellular distribution of auxin transporters (Benjamins et al., 2003; Zhang et al., 2011). These two Ca²⁺ changes have different characteristics. The first transient [Ca2+]_c increase depends on the rotational velocity but not angle, whereas the second sustained [Ca2+]_c increase depends on the rotational angle but not velocity. The first [Ca2+]_c transient was inhibited by Gd³⁺, La³⁺, and the Ca²⁺ chelator 1,2-bis(2-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid but not by ruthenium red (RR), whereas the second sustained [Ca2+]_c increase was inhibited by all these chemicals. These results suggest that the first transient and second sustained [Ca2+]_c increases are related to the rotational stimulation and the gravistimulation, respectively, and are mediated by distinct molecular mechanisms (Toyota et al., 2008a). However, it has not been demonstrated directly that the second sustained [Ca2+]_c increase is induced solely by gravistimulation; it could be influenced by other factors, such as an interaction with the first transient [Ca2+]_c increase (Cessna et al., 1998), vibration, and/or deformation of plants during the rotation.To elucidate the genuine Ca²⁺ signature in response to gravistimulation in plants, we separated rotation and gravistimulation under microgravity (μg; less than 10⁻⁴g) conditions provided by parabolic flight (PF). Using this approach, we were able to apply rotation and gravistimulation to plants separately (Fig. 1). When Arabidopsis seedlings were rotated +180° under μg conditions, the [Ca2+]_c response to the rotation was transient and almost totally attenuated in a few seconds. Gravistimulation (transition from μg to 1.5g) was then applied to these prerotated specimens at the terminating phase of the PF. This gravistimulation without simultaneous rotation induced a sustained [Ca2+]_c increase. The kinetic properties of this sustained [Ca2+]_c increase were examined under different gravity intensities (0.5g–2g) and sequences of gravity intensity changes (Fig. 2A). This analysis revealed that gravistimulation-specific Ca²⁺ response has an almost linear dependency on gravitational acceleration (0.5g–2g) and an extremely rapid responsiveness of less than 1 s.Open in a separate windowFigure 1.Diagram of the experimental procedures for applying separately rotation and gravistimulation to Arabidopsis seedlings. Rotatory stimulation (green arrow) was applied by rotating the seedlings 180° under μg conditions, and 1.5g 180° rotation gravistimulation (blue arrow) was applied to the prerotated seedlings after μg.Open in a separate windowFigure 2.Acceleration, temperature, humidity, and pressure in an aircraft during flight experiments. A, Accelerations along x, y, and z axes in the aircraft during PF. The direction of flight (FWD) and coordinates (x, y, and z) are indicated in the bottom graph. The inset shows an enlargement of the acceleration along the z axis (gravitational acceleration) during μg conditions lasting for approximately 20 s. B, Temperature, humidity, and pressure in the aircraft during PF. Shaded areas in graphs denote the μg condition.     

An Improved Simplified High-Sensitivity Quantification Method for Determining Brassinosteroids in Different Tissues of Rice and Arabidopsis

Quantification of brassinosteroids is essential and extremely important to study the molecular mechanisms of their physiological roles in plant growth and development. Herein, we present a simple, material and cost-saving high-performance method for determining endogenous brassinosteroids (BRs) in model plants. This new method enables simultaneous enrichment of a wide range of bioactive BRs such as brassinolide, castasterone, teasterone, and typhasterol with ion exchange solid-phase extraction and high-sensitivity quantitation of these BRs based on isotope dilution combined with internal standard approach. For routine analysis, the consumption of plant materials was reduced to one-twentieth of previously reported and the overall process could be completed within 1 day compared with previous 3 to 4 days. The strategy was validated by profiling BRs in different ecotypes and mutants of rice (Oryza sativa) and Arabidopsis (Arabidopsis thaliana), and the BR distributions in different model plants tissues were determined with the new method. The method allows plant physiologists to monitor the dynamics and distributions of BRs with 1 gram fresh weight of model plant tissues, which will speed up the process for the molecular mechanism research of BRs with these model plants in future work.Brassinosteroids (BRs) have been considered as the sixth class of endogenous plant hormones with wide occurrence across the plant kingdom (Bajguz and Tretyn, 2003). BRs play a key role in a variety of physiological processes, such as cell elongation, vascular differentiation, reproductive development, photomorphogenesis, stress tolerance, and so on (Hayat, 2010). Recently, it was found that BR deficiency could increase grain yield in rice (Oryza sativa) by more than 30%, which showed a food security-enhancing potential and guided new green revolution in the future (Sakamoto et al., 2006; Wu et al., 2008). Since BRs were first isolated and identified from rape (Brassica napus) pollen in 1970s (Mitchell et al., 1970; Grove et al., 1979), the natural occurrence of more than 60 BRs in a large quantity of plant species has been reported (Hayat, 2010). To date, research on the occurrence of BRs in different plants, physiological properties, and their action modes has made much progress (Fujioka and Yokota, 2003; Symons et al., 2008; Kim and Wang, 2010; Tang et al., 2010; Tong and Chu, 2012). However, so far, only limited information was obtained to understand the molecular mechanism of the physiological role of BRs. For example, although the biosynthetic pathway of C₂₈ BRs has been well established, the biosynthesis of C₂₇ and C₂₉ BRs remains unclear, and some intermediates on their biosynthetic pathways still need to be elucidated (Noguchi et al., 2000; Fujita et al., 2006). The plant physiology research of BRs is speeded up by employing BR mutants on biosynthesis and signaling pathways (Yamamuro et al., 2000; Hong et al., 2003; Kwon and Choe, 2005; Tanabe et al., 2005); however, a simple, high-sensitivity screening, detection, and quantification method for BR analysis is still a bottleneck technique for in-depth studying of the role of BRs during the life cycle of plants.In the past 20 years, most of the detection and identification processes of BRs could be described briefly as the following steps. The harvested plant materials were lyophilized and then ground to a fine powder, followed by the CH₃OH/CHCl₃ extraction. The concentrate was then partitioned with the CHCl₃/H₂O system three times. After that, the CHCl₃ fraction was subjected to a silica gel column for BR enrichment, and the collected BRs-containing fraction was purified with Sephadex LH-20 column and Sep-Pak Plus C18 cartridge in sequence. At last, the collected fractions were purified with preparative HPLC and then derivatized for analysis with gas chromatography-mass spectrometry (MS) under selected ion monitoring mode (Hong et al., 2005; Nomura et al., 2005; Kim et al., 2006). So far, this protocol has been proven to be workable in most cases and provided a great quantity of valuable data for plant physiological research (Hwang et al., 2006; Lee et al., 2010). However, at least more than 20 g of plant materials were consumed for quantifying/identifying BRs in one plant sample without replicates (Hong et al., 2005; Bancos et al., 2006; Kim et al., 2006), and it is difficult to collect sufficient plant tissues for BR measurement in some rare model plant mutants. In addition, the method involved multiple tedious and labor-intensive steps, which might result in poor recovery and low sensitivity, especially for some labile BR intermediates. The traditional method took one person about 3 to 4 d to treat one batch of samples. Most of the BR measurement experiments were performed without biological replicates using traditional methods due to the disadvantages mentioned above, which discounted the reliability of the results. Therefore, a simple, rapid, and sensitive analysis method for BRs is in urgent need, along with the development of BR research.Recently, several efforts were made to improve the BR determination (Svatos et al., 2004; Huo et al., 2012). The consumption of plant material was reduced to 2 g after modifying the BRs with a new derivatization reagent for further ultra-performance liquid chromatography (UPLC)-multiple reaction monitoring (MRM)-MS detection. However, the purification process consisting of deproteinization and multiple solid-phase extraction (SPE) steps was still quite tedious and couldn’t guarantee covering the four most important bioactive BRs, including brassinolide (BL), castasterone (CS), teasterone (TE), and typhasterol (TY; Fig. 1). In our previous study (Xin et al., 2013), we reported a simple, convenient, and high-sensitivity method for detection of endogenous BRs from real plant materials based on the dual role of specific boronate affinity. Although it was the first time to measure multiple BRs in subgram plant materials and the time duration of the method decreased to one-third of that previously reported, the synthesis of boronate affinity-functionalized magnetic nanoparticles made the method difficult to follow in biological laboratories for routine analysis.Open in a separate windowFigure 1.Chemical structure of four major bioactive BRs.BRs are neutral steroid compounds with a common four-ring cholestane skeleton and hydroxyl groups on A ring and/or the side chain linked to D ring. Especially, the vicinal diol moieties on C₂₂ and C₂₃ sites of BL, CS, TY, and TE allow these bioactive BRs to be derivatized with ionizable reagents for MS response enhancement. Considering the unique physicochemical properties of BRs, we herein developed a simplified high-sensitivity analytical method based on mixed-mode anion exchange (MAX)-cation exchange (MCX) solid phase extraction (SPE) purification, vicinal diol derivatization combined with UPLC-MRM3-MS detection for quantification of BL, CS, TE, and TY in model plants (Fig. 2). The performance of the method was demonstrated by determination of BRs in different tissues of both wild-type and mutant Arabidopsis (Arabidopsis thaliana) and rice.Open in a separate windowFigure 2.Simplified high-sensitivity protocol for quantitative analysis of BRs. IS, Internal standards. [See online article for color version of this figure.]     

Acyl Editing and Headgroup Exchange Are the Major Mechanisms That Direct Polyunsaturated Fatty Acid Flux into Triacylglycerols

Triacylglycerols (TAG) in seeds of Arabidopsis (Arabidopsis thaliana) and many plant species contain large amounts of polyunsaturated fatty acids (PUFA). These PUFA are synthesized on the membrane lipid phosphatidylcholine (PC). However, the exact mechanisms of how fatty acids enter PC and how they are removed from PC after being modified to participate in the TAG assembly are unclear, nor are the identities of the key enzymes/genes that control these fluxes known. By reverse genetics and metabolic labeling experiments, we demonstrate that two genes encoding the lysophosphatidylcholine acyltransferases LPCAT1 and LPCAT2 in Arabidopsis control the previously identified “acyl-editing” process, the main entry of fatty acids into PC. The lpcat1/lpcat2 mutant showed increased contents of very-long-chain fatty acids and decreased PUFA in TAG and the accumulation of small amounts of lysophosphatidylcholine in developing seeds revealed by [¹⁴C]acetate-labeling experiments. We also showed that mutations in LPCATs and the PC diacylglycerol cholinephosphotransferase in the reduced oleate desaturation1 (rod1)/lpcat1/lpcat2 mutant resulted in a drastic reduction of PUFA content in seed TAG, accumulating only one-third of the wild-type level. These results indicate that PC acyl editing and phosphocholine headgroup exchange between PC and diacylglycerols control the majority of acyl fluxes through PC to provide PUFA for TAG synthesis.Plant oils are an important natural resource to meet the increasing demands of food, feed, biofuel, and industrial applications (Lu et al., 2011; Snapp and Lu, 2012). The fatty acid composition in the triacylglycerols (TAG), especially the contents of polyunsaturated fatty acids (PUFA) or other specialized structures, such as hydroxy, epoxy, or conjugated groups, determines the properties and thus the uses of plant oils (Dyer and Mullen, 2008; Dyer et al., 2008; Pinzi et al., 2009; Riediger et al., 2009). To effectively modify seed oils tailored for different uses, it is necessary to understand the fundamental aspects of how plant fatty acids are synthesized and accumulated in seed oils.In developing oilseeds, fatty acids are synthesized in plastids and are exported into the cytosol mainly as oleic acid, 18:1 (carbon number:double bonds), and a small amount of palmitic acid (16:0) and stearic acid (18:0; Ohlrogge and Browse, 1995). Further modification of 18:1 occurs on the endoplasmic reticulum in two major pathways (Fig. 1): (1) the 18:1-CoA may be elongated into 20:1- to 22:1-CoA esters by a fatty acid elongase, FAE1 (Kunst et al., 1992); (2) the dominant flux of 18:1 in many oilseeds is to enter the membrane lipid phosphatidylcholine (PC; Shanklin and Cahoon, 1998; Bates and Browse, 2012), where they can be desaturated by the endoplasmic reticulum-localized fatty acid desaturases including the oleate desaturase, FAD2, and the linoleate desaturase, FAD3, to produce the polyunsaturated linoleic acid (18:2) and α-linolenic acid (18:3; Browse et al., 1993; Okuley et al., 1994). The PUFA may be removed from PC to enter the acyl-CoA pool, or PUFA-rich diacylglycerol (DAG) may be derived from PC by removal of the phosphocholine headgroup (Bates and Browse, 2012). The PUFA-rich TAG are then produced from de novo-synthesized DAG or PC-derived DAG (Bates and Browse, 2012) and PUFA-CoA by the acyl-CoA:diacylglycerol acyltransferases (DGAT; Hobbs et al., 1999; Zou et al., 1999). Alternatively, PUFA may be directly transferred from PC onto DAG to form TAG by an acyl-CoA-independent phospholipid:diacylglycerol acyltransferase (PDAT; Dahlqvist et al., 2000). Recent results demonstrated that DGAT and PDAT are responsible for the majority of TAG synthesized in Arabidopsis (Arabidopsis thaliana) seeds (Zhang et al., 2009).Open in a separate windowFigure 1.Reactions involved in the flux of fatty acids into TAG. De novo glycerolipid synthesis is shown in white arrows, acyl transfer reactions are indicated by dashed lines, and the movement of the lipid glycerol backbone through the pathway is shown in solid lines. Major reactions (in thick lines) controlling the flux of fatty acid from PC into TAG are as follows: LPC acylation reaction of acyl editing by LPCAT (A); PC deacylation reaction of acyl editing by the reverse action of LPCAT or phospholipase A (B); and the interconversion of DAG and PC by PDCT (C). Substrates are in boldface, enzymatic reactions are in italics. FAD, Fatty acid desaturase; FAS, fatty acid synthase; GPAT, acyl-CoA:G3P acyltransferase; LPA, lysophosphatidic acid; LPAT, acyl-CoA:LPA acyltransferase; PA, phosphatidic acid; PLC, phospholipase C; PLD, phospholipase D.The above TAG synthesis model highlights the importance of acyl fluxes through PC for PUFA enrichment in plant oils. However, the exact mechanisms of how fatty acids enter PC and how they are removed from PC after being modified to participate in the TAG assembly are unclear, nor are the identities of the enzymes/genes that control these fluxes known. The traditional view is that 18:1 enters PC through de novo glycerolipid synthesis (Fig. 1; Kennedy, 1961): the sequential acylation of glycerol-3-phosphate (G3P) at the sn-1 and sn-2 positions produces phosphatidic acid; subsequent removal of the phosphate group at the sn-3 position of phosphatidic acid by phosphatidic acid phosphatases (PAPs) produces de novo DAG; finally, PC is formed from DAG by a cytidine-5′-diphosphocholine:diacylglycerol cholinephosphotransferase (CPT; Slack et al., 1983; Goode and Dewey, 1999). However, metabolic labeling experiments in many different plant tissues by us and others (Williams et al., 2000; Bates et al., 2007, 2009; Bates and Browse, 2012; Tjellström et al., 2012) have demonstrated that the majority of newly synthesized fatty acids (e.g. 18:1) enter PC by a process termed “acyl editing” rather than by proceeding through de novo PC synthesis. Acyl editing is a deacylation-reacylation cycle of PC that exchanges the fatty acids on PC with fatty acids in the acyl-CoA pool (Fig. 1, A and B). Through acyl editing, newly synthesized 18:1 can be incorporated into PC for desaturation and PUFA can be released from PC to the acyl-CoA pool to be utilized for glycerolipid synthesis.Additionally, there is accumulating evidence that many plants utilize PC-derived DAG to synthesize TAG laden with PUFA (Bates and Browse, 2012). PC-derived DAG may be synthesized through the reverse reaction of the CPT (Slack et al., 1983, 1985) or by the phospholipases C and D (followed by PAP). However, our recent discovery indicates that the main PC-to-DAG conversion is catalyzed by a phosphatidylcholine:diacylglycerol cholinephosphotransferase (PDCT) through the phosphocholine headgroup exchange between PC and DAG (Fig. 1C; Lu et al., 2009; Hu et al., 2012). The PDCT is encoded by the REDUCED OLEATE DESATURATION1 (ROD1) gene (At3g15820) in Arabidopsis, which is responsible for about 40% of the flux of PUFA from PC through DAG into TAG synthesis (Lu et al., 2009). Acyl editing and PC-DAG interconversion through PDCT may work together to generate PUFA-rich TAG in oilseed plants (Bates and Browse, 2012).The enzymes/genes involved in the incorporation of 18:1 into PC through acyl editing are not known. However, stereochemical localization of newly synthesized fatty acid incorporation into PC predominantly at the sn-2 position (Bates et al., 2007, 2009; Tjellström et al., 2012) strongly suggest that the acyl editing cycle proceeds through the acylation of lysophosphatidylcholine (LPC) by acyl-CoA:lysophosphatidylcholine acyltransferases (LPCATs [Enzyme Commission 2.3.1.23]; Fig. 1A). High LPCAT activity has been detected in many different oilseed plants that accumulate large amounts of PUFA in TAG (Stymne and Stobart, 1987; Bates and Browse, 2012), suggesting the potential ubiquitous involvement of LPCAT in the generation of PUFA-rich TAG. Several possible pathways for the removal of acyl groups from PC to generate the lysophosphatidylcholine within the acyl editing cycle have been proposed. The acyl groups may be released from PC to enter the acyl-CoA pool via the reverse reactions of LPCATs (Stymne and Stobart, 1984) or by reactions of phospholipase A (Chen et al., 2011) followed by the acyl-CoA synthetases (Shockey et al., 2002). The main focus of this study was to identify the genes and enzymes involved in the incorporation of fatty acids into PC through acyl editing in Arabidopsis and to quantify the contribution of acyl editing and PDCT-based PC-DAG interconversion to controlling the flux of PUFA from PC into TAG. Herein, we demonstrate that mutants of two Arabidopsis genes encoding LPCATs (At1g12640 [LPCAT1] and At1g63050 [LPCAT2]) have reduced TAG PUFA content. Analysis of the acyl-editing cycle through metabolic labeling of developing seeds with [¹⁴C]acetate indicate that the lpcat1/lpcat2 double mutant was devoid of acyl editing-based incorporation of newly synthesized fatty acids into PC, indicating that these two genes are responsible for the acylation of LPC during acyl editing. Additionally, the triple mutant rod1/lpcat1/lpcat2 indicated that PDCT-based PC-DAG interconversion and acyl editing together provide two-thirds of the flux of PUFA from PC to TAG in Arabidopsis seeds.     

Association of the P6 Protein of Cauliflower mosaic virus with Plasmodesmata and Plasmodesmal Proteins

The P6 protein of Cauliflower mosaic virus (CaMV) is responsible for the formation of inclusion bodies (IBs), which are the sites for viral gene expression, replication, and virion assembly. Moreover, recent evidence indicates that ectopically expressed P6 inclusion-like bodies (I-LBs) move in association with actin microfilaments. Because CaMV virions accumulate preferentially in P6 IBs, we hypothesized that P6 IBs have a role in delivering CaMV virions to the plasmodesmata. We have determined that the P6 protein interacts with a C2 calcium-dependent membrane-targeting protein (designated Arabidopsis [Arabidopsis thaliana] Soybean Response to Cold [AtSRC2.2]) in a yeast (Saccharomyces cerevisiae) two-hybrid screen and have confirmed this interaction through coimmunoprecipitation and colocalization assays in the CaMV host Nicotiana benthamiana. An AtSRC2.2 protein fused to red fluorescent protein (RFP) was localized to the plasma membrane and specifically associated with plasmodesmata. The AtSRC2.2-RFP fusion also colocalized with two proteins previously shown to associate with plasmodesmata: the host protein Plasmodesmata-Localized Protein1 (PDLP1) and the CaMV movement protein (MP). Because P6 I-LBs colocalized with AtSRC2.2 and the P6 protein had previously been shown to interact with CaMV MP, we investigated whether P6 I-LBs might also be associated with plasmodesmata. We examined the colocalization of P6-RFP I-LBs with PDLP1-green fluorescent protein (GFP) and aniline blue (a stain for callose normally observed at plasmodesmata) and found that P6-RFP I-LBs were associated with each of these markers. Furthermore, P6-RFP coimmunoprecipitated with PDLP1-GFP. Our evidence that a portion of P6-GFP I-LBs associate with AtSRC2.2 and PDLP1 at plasmodesmata supports a model in which P6 IBs function to transfer CaMV virions directly to MP at the plasmodesmata.Through the years, numerous studies have focused on the characterization of viral replication sites within the cell, as well as how plant virus movement proteins (MPs) modify the plasmodesmata to facilitate cell-to-cell movement (for review, see Benitez-Alfonso et al., 2010; Laliberté and Sanfaçon, 2010; Niehl and Heinlein, 2011; Ueki and Citovsky, 2011; Verchot, 2012). It is accepted that plant virus replication is associated with host membranes, and at some point, the viral genomic nucleic acid must be transferred from the site of replication in the cell to the plasmodesmata. This step could involve transport from a distant site within the cell, or alternatively, it may be that replication is coupled with transport at the entrance of the plasmodesmata (Tilsner et al., 2013). However, even with the latter model, there is ample evidence that the viral proteins necessary for replication or cell-to-cell movement utilize intracellular trafficking pathways within the cell to become positioned at the plasmodesma. These pathways may involve microfilaments, microtubules, or specific endomembranes that participate in macromolecular transport pathways, or combinations of these elements (Harries et al., 2010; Schoelz et al., 2011; Patarroyo et al., 2012; Peña and Heinlein, 2012; Tilsner and Oparka 2012; Liu and Nelson, 2013).The P6 protein of Cauliflower mosaic virus (CaMV) is one viral protein that had not been considered to play a role in viral movement until recently. P6 is the most abundant protein component of the amorphous, electron-dense inclusion bodies (IBs) present during virus infection (Odell and Howell, 1980; Shockey et al., 1980). Ectopic expression of P6 in Nicotiana benthamiana leaves resulted in the formation of inclusion-like bodies (I-LBs) that were capable of intracellular movement along actin microfilaments. Furthermore, treatment of Nicotiana edwardsonii leaves with latrunculin B abolished the formation of CaMV local lesions, suggesting that intact microfilaments are required for CaMV infection (Harries et al., 2009a). A subsequent paper showed that P6 physically interacts with Chloroplast Unusual Positioning1 (CHUP1), a plant protein localized to the chloroplast outer membrane that contributes to movement of chloroplasts on microfilaments in response to changes in light intensity (Oikawa et al., 2003, 2008; Angel et al., 2013). The implication was that P6 might hijack CHUP1 to facilitate movement of the P6 IBs on microfilaments. Silencing of CHUP1 in N. edwardsonii, a host for CaMV, slowed the rate of local lesion formation, suggesting that CHUP1 contributes to intracellular movement of CaMV (Angel et al., 2013).In addition to its role in intracellular trafficking, the P6 protein has been shown to have at least four other distinct functions in the viral infection cycle. P6-containing IBs induced during virus infection are likely virion factories, as they are the primary site for CaMV protein synthesis, genome replication, and assembly of virions (Hohn and Fütterer, 1997). Second, P6 interacts with host ribosomes to facilitate reinitiation of translation of genes on the polycistronic 35S viral RNA, a process called translational transactivation (Bonneville et al., 1989; Park et al., 2001; Ryabova et al., 2002). The translational transactivator region of P6 (Fig. 1) defines the essential sequences required for translational transactivation (DeTapia et al., 1993). Third, P6 is an important pathogenicity determinant. P6 functions as an avirulence determinant in some solanaceous and cruciferous species (Daubert et al., 1984; Schoelz et al., 1986; Hapiak et al., 2008) and is a chlorosis symptom determinant in susceptible hosts (Daubert et al., 1984; Baughman et al., 1988; Goldberg et al., 1991; Cecchini et al., 1997). Finally, P6 has the capacity to compromise host defenses, as it is a suppressor of RNA silencing and cell death (Love et al., 2007; Haas et al., 2008), and it modulates signaling by salicylic acid, jasmonic acid, ethylene, and auxin (Geri et al., 2004; Love et al., 2012; Laird et al., 2013). Domain D1 of P6 has been shown to be necessary but not sufficient for suppression of silencing and salicylic acid-mediated defenses (Laird et al., 2013).Open in a separate windowFigure 1.CaMV and host constructs used for confocal microscopy or coimmunoprecipitation (co-IP). A, Structure of CaMV P6 and Arabidopsis (Arabidopsis thaliana) Soybean Response to Cold (AtSRC2.2) proteins. The functions of P6 domains D1 to D4 tested for interaction with AtSRC2.2 are indicated by the shaded boxes. The Mini TAV is the minimal region for the translational transactivation function. The NLSa sequence corresponds to the nuclear localization signal of influenza virus. The NLS sequence corresponds to the nuclear localization signal of human ribosomal protein L22. B, Structure of P6 (Angel et al., 2013), AtSRC2.2, PDLP (Thomas et al., 2008), and CaMV MP fusions developed for confocal microscopy and/or co-IP. aa, Amino acid.Because P6-containing IBs are the site for virion accumulation and they are capable of movement, they may be responsible for delivering virions to the CaMV MP located at the plasmodesmata (for review, see Schoelz et al., 2011). The vast majority of CaMV virions accumulate in association with P6-containing IBs. Furthermore, P6 physically interacts with the CaMV capsid and MP, as well as the two proteins necessary for aphid transmission, P2 and P3 (Himmelbach et al., 1996; Ryabova et al., 2002; Hapiak et al., 2008; Lutz et al., 2012). Recent studies have indicated that P6 IBs serve as a reservoir for virions, in which the virions may be rapidly transferred to P2 electron-lucent IBs for acquisition by aphids (Bak et al., 2013). It stands to reason that P6 IBs may also serve as a reservoir for CaMV virions to be transferred to the CaMV MP in the plasmodesmata.CaMV virions move from cell to cell through plasmodesmata modified into tubules through the function of its MP (Perbal et al., 1993; Kasteel et al., 1996). However, studies have suggested that CaMV virions do not appear to directly interact with the MP. Instead, the MP interacts with the CaMV P3 protein (also known as the virion-associated protein [VAP]), which forms a trimeric structure that is anchored into the virions (Leclerc et al., 1998; Leclerc et al., 2001). Electron microscopy studies have indicated that MP and VAP colocalize with virions only at the entrance to or within the plasmodesmata, and it has been suggested that the VAP/virion complex travels to the plasmodesmata independently from the MP (Stavolone et al., 2005). Consequently, there is a need for a second CaMV protein such as P6 to fulfill the role of delivery of virions to the plasmodesmata (Schoelz et al., 2011).Additional studies have shown that the CaMV MP is incorporated into vesicles and is trafficked on the endomembrane system to reach the plasmodesma (Carluccio et al., 2014). These authors suggest that the CaMV MP is recycled in a vesicular transport pathway between plasmodesmata and early endosome compartments. The CaMV MP interacts with µA-Adaptin (Carluccio et al., 2014) and Movement Protein-Interacting7 (Huang et al., 2001), two proteins shown to have a role in vesicular trafficking. Once the MP arrives at plasmodesmata, it interacts with the Plasmodesmata-Localized Protein (PDLP) proteins, which comprise a family of eight proteins associated with plasmodesmata (Amari et al., 2010). In addition to its interaction with CaMV MP, PDLP1 interacts with the 2B protein of Grapevine fan leaf virus (GFLV) at the base of tubules formed by the 2B protein. Furthermore, an Arabidopsis transfer DNA (T-DNA) mutant line in which three PDLP genes had been knocked out (pdlp1-pdlp2-pdlp3) responded to GFLV and CaMV inoculation with a delayed infection (Amari et al., 2010). This has led to the suggestion that the PDLPs might act as receptors for the MPs of the tubule-forming viruses such as GFLV and CaMV (Amari et al., 2010, 2011).To better understand the function of the P6 protein during CaMV intracellular movement, we have utilized a yeast (Saccharomyces cerevisiae) two-hybrid assay to identify host proteins that interact with CaMV P6. We show that P6 physically interacts with a C2-calcium-dependent protein (designated AtSRC2.2). AtSRC2.2 is a membrane-bound protein that is capable of forming punctate spots associated with plasmodesmata. The localization of AtSRC2.2 with plasmodesmata led to an analysis of interactions between P6 I-LBs, AtSRC2.2, PDLP1, and the CaMV MP and also revealed that a portion of P6 I-LBs are found adjacent to plasmodesmata. These results provide further evidence for a model in which P6 IBs are capable of delivery of virions to plasmodesmata for their transit to other host cells.     

Crystal Structures of Physcomitrella patens AOC1 and AOC2: Insights into the Enzyme Mechanism and Differences in Substrate Specificity

In plants, oxylipins regulate developmental processes and defense responses. The first specific step in the biosynthesis of the cyclopentanone class of oxylipins is catalyzed by allene oxide cyclase (AOC) that forms cis(+)-12-oxo-phytodienoic acid. The moss Physcomitrella patens has two AOCs (PpAOC1 and PpAOC2) with different substrate specificities for C₁₈- and C₂₀-derived substrates, respectively. To better understand AOC’s catalytic mechanism and to elucidate the structural properties that explain the differences in substrate specificity, we solved and analyzed the crystal structures of 36 monomers of both apo and ligand complexes of PpAOC1 and PpAOC2. From these data, we propose the following intermediates in AOC catalysis: (1) a resting state of the apo enzyme with a closed conformation, (2) a first shallow binding mode, followed by (3) a tight binding of the substrate accompanied by conformational changes in the binding pocket, and (4) initiation of the catalytic cycle by opening of the epoxide ring. As expected, the substrate dihydro analog cis-12,13S-epoxy-9Z,15Z-octadecadienoic acid did not cyclize in the presence of PpAOC1; however, when bound to the enzyme, it underwent isomerization into the corresponding trans-epoxide. By comparing complex structures of the C₁₈ substrate analog with in silico modeling of the C₂₀ substrate analog bound to the enzyme allowed us to identify three major molecular determinants responsible for the different substrate specificities (i.e. larger active site diameter, an elongated cavity of PpAOC2, and two nonidentical residues at the entrance of the active site).Oxylipins comprise a large family of oxidized fatty acids and metabolites thereof (Acosta and Farmer, 2010). They are abundant in mammals (Funk, 2001) and flowering plants (Creelman and Mulpuri, 2002). In addition, they have been found in fungi (Brodhun and Feussner, 2011) as well as in nonflowering plants like mosses and algae (Andreou et al., 2009). In plants, these lipids serve as signaling molecules regulating developmental processes and mediating defense reactions (Howe and Jander, 2008; Browse, 2009; Acosta and Farmer, 2010). The first committed step in oxylipin biosynthesis is the peroxidation of a polyunsaturated fatty acid containing a 1Z,4Z-pentadiene system by lipoxygenase (LOX) or the peroxidation at the C2 position of a fatty acid by α-dioxygenase. These reactions start the so-called LOX or oxylipin pathway (Feussner and Wasternack, 2002) and are followed by further enzymatic reactions in which the hydroperoxy fatty acid is converted to a set of different secondary products. In the case of LOX-derived hydroperoxy fatty acids, such conversions are mainly catalyzed by members of the cytochrome P450 subfamily Cyp74 (i.e. fatty acid hydroperoxide lyase, divinyl ether synthase, epoxy alcohol synthase, and allene oxide synthase [AOS]; Stumpe and Feussner, 2006; Lee et al., 2008). Additional conversions of the fatty acid hydroperoxide are catalyzed by other proteins, such as LOX or peroxygenase (Mosblech et al., 2009).Jasmonic acid (JA) biosynthesis is one specific branch of the oxylipin pathway. It may start with the release of α-linolenic acid [18:3(n-3)] from membrane lipids by a lipase (Schaller and Stintzi, 2009). This free fatty acid is subsequently oxidized by a 13-LOX to yield 13-hydroperoxy octadecatrienoic acid (13-HPOTE) and converted by the action of AOS into the unstable allene oxide 12,13S-epoxy-9Z,11E,15Z-octadecatrienoic acid (12,13-EOT; Fig. 1). 12,13-EOT is then cyclized by allene oxide cyclase (AOC) to the cyclopentenone derivative cis(+)-12-oxo-phytodienoic acid [cis(+)-OPDA]. In the absence of AOC, the epoxide is hydrolyzed into ketols and racemic 12-oxo-phytodienoic acid (OPDA). cis(+)-OPDA is the first cyclic and biologically active compound in that pathway (Dave and Graham, 2012). While the reactions leading from 18:3(n-3) to cis(+)-OPDA occur in the plastid, all further enzymatic steps resulting in the formation of JA are localized in the peroxisomes (Wasternack, 2007). Here, cis(+)-OPDA is reduced in a NADPH-dependent reaction by cis(+)-OPDA reductase isoform 3 to 3-oxo-2(2′Z-pentenyl)-cyclopentane-1-octanoic acid. This step is followed by activation of the carboxyl group and three steps of β-oxidation and finally leads to the formation of (+)-7-iso-JA (Dave and Graham, 2012).Open in a separate windowFigure 1.Overview of the enzymatic steps in JA biosynthesis with molecular focus (box) on the reaction catalyzed by AOC. JA biosynthesis may start with the release of 18:3(n-3) or roughanic acid from a lipid. Next, the fatty acid is oxidized by a 13-LOX, yielding the 13-hydroperoxy derivative. This serves as a substrate for a subsequent conversion catalyzed by AOS and AOC, yielding the cyclopentenone derivatives cis(+)-OPDA and cis(+)-dinorOPDA, respectively, via an unstable allene oxide. Cyclization of the allene oxide seems to be initiated by one particular Glu residue in the active site of AOC that leads to an opening of the epoxy ring, conformational changes, and a concerted pericyclic ring closure (details are explained in the text). After reduction of the cyclopentenone by cis(+)-OPDA reductase isoform 3 (OPR3), the octanoic or hexanoic side chain is shortened by β-oxidation cycles.The conversion of 13-HPOTE into cis(+)-OPDA was first observed using a flaxseed (Linum usitatissimum) acetone powder preparation and was suggested to take place via a hypothetical epoxide intermediate (Vick et al., 1980). Later studies unequivocally demonstrated that 12,13-EOT (Hamberg, 1987; Brash et al., 1988), an allene oxide formed from 13-HPOTE by AOS (Song and Brash, 1991; Song et al., 1993), serves as substrate for the cyclization reaction catalyzed by AOC (Hamberg and Fahlstadius, 1990). The enzyme was purified (Ziegler et al., 1997), characterized with regard to the substrate specificity (Ziegler et al., 1999), and cloned and recombinantly expressed (Ziegler et al., 2000; Stenzel et al., 2003). In 2006, the crystal structure of an AOC from Arabidopsis (Arabidopsis thaliana; AtAOC2) was solved (Hofmann et al., 2006), and the reaction mechanism as well as the subcellular localization were studied (Schaller et al., 2008). The enzyme crystallized as a homotrimer, with each subunit forming an eight-stranded antiparallel β-barrel harboring a hydrophobic cavity in which the active site of the enzyme is located. While the exterior loops showed a high degree of flexibility, the central part of the enzyme was very rigid, and no induced-fit mechanism could be observed upon binding of a substrate analog (Hofmann et al., 2006). Based on the structure of AtAOC2 in complex with vernolic acid [cis(+/−)-12,13-epoxy-9Z-octadecenoic acid (12,13-EOM)] as an inert substrate analog, the following reaction mechanism has been proposed (Fig. 1, box): the allene oxide substrate binds with its fatty acid backbone deep in the barrel, where it interacts with hydrophobic amino acid residues, while the polar carboxy head group is located on the exterior of the cavity. One particular Glu residue (Glu-23 in AtAOC2) pointing to the Δ^15Z-double bond of the substrate may induce a partial charge separation that leads to a delocalization of the π-electron system, thereby facilitating opening of the epoxide ring. The oxyanion thus formed is stabilized via polar interactions with a catalytic, structurally conserved water molecule that is positioned in the polar cavity of the enzyme formed by two Asn residues (Asn-25 and Asn-53 in AtAOC2, respectively), one Ser (Ser-31 in AtAOC2), and one Pro (Pro-32 in AtAOC2). The ring closure that leads to the formation of the cyclopentenone derivative is achieved by a conformational reorganization of the C10-C11 substrate bond from the trans- to the cis-geometry. Due to steric limitations in the active site, this rotation may be accompanied by a cis/trans-isomerization of the C8-C9 substrate bond. Since the enzyme dictates the stereochemistry of the final ring closure, the released product is exclusively the (+)-enantiomer, cis(+)-OPDA (Schaller et al., 2008). Notably, this reaction competes with the spontaneous decomposition of the allene oxide substrate that leads to the formation of racemic OPDA as well as α-ketols and γ-ketols. This hints toward a low-energy barrier of the cyclization reaction and suggests that AOC does not need much of a catalytic functionality in terms of lowering this barrier (Schaller and Stintzi, 2009). It has been proposed that the enzymatic cyclization reaction is achieved according to the rules of Hoffmann and Woodward (1970) via a concerted pericyclic ring closure while spontaneous cyclization proceeds through a dipolar ring closure (Grechkin et al., 2002). The facts that the allene oxide formed by AOS has a very short half-life in aqueous solution and that natural OPDA is found in its enantiopure cis(+)-configuration suggest that AOS and AOC are coupled. However, no physical interaction of both enzymes may be necessary to form cis(+)-OPDA in vitro (Zerbe et al., 2007).Recently, it was shown that the moss Physcomitrella patens harbors and metabolizes not only C₁₈ but also C₂₀ polyunsaturated fatty acids to form oxylipins (Fig. 2; Stumpe et al., 2010). In particular, it was shown that (12S)-hydroperoxy eicosatetraenoic acid (12-HPETE) is endogenously formed by a bifunctional LOX as the major hydroperoxy fatty acid of arachidonic acid [20:4(n-6)] (Wichard et al., 2004). 12-HPETE serves as a substrate for further conversions either leading to the formation of C₈- and C₉-volatiles (e.g. octenals, octenols, and nonenals) or the cyclopentenone derivative 11-oxo prostatrienoic acid (11-OPTA; Stumpe et al., 2010). Whereas the volatiles are formed by at least two bifunctional LOXs with an additional hydroperoxide lyase activity (Wichard et al., 2004; Senger et al., 2005; Anterola et al., 2009) or by a Cyp74-derived hydroperoxide lyase (Stumpe et al., 2006), 11-OPTA is formed in analogy to the octadecanoids by one particular AOC, PpAOC2, via the allene oxide intermediate formed by PpAOS (Bandara et al., 2009). In contrast, PpAOC1 does not accept the 12-HPETE-derived C₂₀-allene oxide and thus converts only the 13-HPOTE-derived allene oxide.Open in a separate windowFigure 2.AOS/AOC pathways in P. patens. 13-HPOTE is converted by PpAOS to 12,13-EOT, which may either hydrolyze in the absence of PpAOC1 or PpAOC1 to ketols and racemic OPDA or, in the presence of PpAOC1 and PpAOC2, cyclize to cis(+)-OPDA. 12-HPETE is converted by PpAOS to 11,12-EET, which again may either hydrolyze in the absence of PpAOC2 to ketols and racemic OPDA or, in the presence of PpAOC2, cyclize to 11-OPTA.In this study, the crystal structures of PpAOC1 and PpAOC2 were solved. Data were also obtained for mutated forms of PpAOC1 and for PpAOC1 and PpAOC2 in complex with the allene oxide stable analog 12,13-EOD. In this way, detailed information about the allene oxide-to-cyclopentenone conversions promoted by the two AOCs was obtained.     

Tryptophan-Independent Indole-3-Acetic Acid Synthesis: Critical Evaluation of the Evidence

Evidence for Trp-independent IAA synthesis is critically reevaluated in the light of tryptophan synthase proteome data, local IAA synthesis and Trp, indole-3-pyruvate, and IAA turnover.Trp-independent synthesis of indole-3-acetic acid (IAA) was proposed back in the early 1990s based on observations from Trp auxotrophs in maize (Zea mays; Wright et al., 1991) and Arabidopsis (Arabidopsis thaliana; Normanly et al., 1993). Recently, Wang et al. (2015) published new data suggesting that a cytosolic indole synthase (INS) may catalyze the first step separating the Trp-dependent and Trp-independent pathways in Arabidopsis. If this is the case, it would be a major breakthrough; however, in this article, I critically evaluate both recent and older evidence for the Trp-independent route and suggest that the INS is more likely to participate in Trp-dependent IAA production.The original work supporting Trp-independent IAA production was carried out prior to the availability of genome/proteome data and before the discovery that the final step of Trp-dependent IAA synthesis is carried out by a large number of YUCCA homologs operating in a highly localized manner (Zhao, 2008). I argue that experimental data supporting the Trp-independent route needs to be reconsidered in light of complete proteome data. Further, the evidence from feeding labeled compounds should be critically evaluated in light of recent data on the highly localized nature of IAA synthesis as well as older quantitative data on Trp, indole-3-pyruvic acid (IPA), and IAA turnover from my own laboratory (Cooney and Nonhebel, 1991). I conclude that evidence for the Trp-independent route is at best equivocal, and that it is not a conserved source of IAA in angiosperms.Figure 1 shows the major Trp-dependent route for IAA production whereby Trp, produced by the concerted action of Trp synthase α- and β-subunits, is converted to IAA in a further two steps catalyzed by Trp aminotransferase and the flavin monooxygenases commonly known as YUCCA (Mashiguchi et al., 2011; Won et al., 2011). This is compared with the Trp-independent route in which IAA may be produced from free indole by an unknown route (Ouyang et al., 2000; Wang et al., 2015).Open in a separate windowFigure 1.Outline of the major pathway for Trp-dependent IAA synthesis and the proposed Trp-independent route. The role proposed for Trp synthase beta (TSB) homologs discussed in the present paper is also shown. For clarity, reactions are simplified to show only the major compounds relevant to IAA synthesis.The Trp-independent route was originally based on data from Trp auxotrophs that have mutations in genes encoding either the α- or β-subunits of Trp synthase. The α-subunit catalyzes the removal of the side chain from indole-3-glycerol phosphate, passing the indole product directly to the β-subunit where the Trp side chain is created from a Ser substrate (Pan et al., 1997). In plants, this is a chloroplast-localized enzyme. Elevated levels of IAA have been reported in Trp auxotrophs of both maize and Arabidopsis. However, the trp3-1 and trp2-1 mutants of Arabidopsis, deficient in the α- and β-subunits, respectively, only showed an increase in total IAA measured following conjugate hydrolysis. No difference in free IAA levels was found (Normanly et al., 1993). The orange pericarp (orp) maize mutant was reported to have 50 times more IAA than the wild type (Wright et al., 1991). However, this was also total IAA; no data on free IAA were published. Work by Müller and Weiler (2000) indicated that IAA measured following conjugate hydrolysis could have originated via the degradation of indole-3-glycerol phosphate that accumulates in trp3-1 mutants. Further doubt regarding the accuracy of IAA measurements following conjugate hydrolysis has recently been published. Yu et al. (2015) have shown that conjugate hydrolysis treatment substantially overestimates the actual conjugated IAA due to degradation of glucobrassicin and proteins. In addition, neither report (Wright et al., 1991; Normanly et al., 1993) described a high auxin phenotype for the Trp auxotrophs. This contrasts with the superroot1 (sur1) and sur2 mutants, where the accumulation of indole intermediates resulted in a high level of free IAA as well as a high auxin phenotype (Boerjan et al., 1995; Delarue et al., 1998). It is therefore doubtful that Trp auxotrophs actually accumulate more IAA than the wild-type plants.In addition, proteome data have revealed new homologs of TSB in both Arabidopsis and maize that may contribute to Trp production in TSB mutants; these have not been considered in arguments supporting Trp-independent IAA synthesis. Maize orp has mutations in two TSB genes, resulting in a seedling lethal phenotype with high levels of accumulated indole. However, proteome sequence information now indicates that maize has three TSB genes. Plants and bacteria have divergent forms of TSB, type 1 and type 2 (Xie et al., 2001); the major TSB genes responsible for Trp synthase activity in maize and Arabidopsis are type 1. The third maize TSB gene, maize locus ID GRMZM2G054465, is a member of the TSB type 2 group. Its product is reported not to interact directly with a Trp synthase alpha (TSA) subunit but has experimentally demonstrated catalytic activity converting indole and Ser to Trp (Yin et al., 2010). This type 2 TSB may allow orp plants to make sufficient Trp for IAA production from the accumulated free indole.When the original work on trp2 mutants of Arabidopsis was carried out, two TSB genes were known (Last et al., 1991). As the trp2 plants were deficient only in TSB1, they were able to make sufficient Trp to survive under low-light conditions. Full proteome data now indicate that Arabidopsis has four TSB-like genes; in addition to TSB1 and TSB2, there is a third type 1 TSB gene, Arabidopsis locus ID AT5G28237. The product of this gene has not been experimentally characterized. The fourth gene, AT5G38530, encodes a type 2 TSB with demonstrated catalytic activity similar to ZmTSB type 2 mentioned above (Yin et al., 2010). Thus, the trp2 plants may also make enough Trp for IAA production. It is even possible that one of the minor forms of TSB has a specific role in IAA production. Type 2 TSBs are conserved throughout the plant kingdom, and the biological role for this protein is not known (Xie et al., 2001).A phylogenetic analysis of type 1 TSBs is shown in Figure 2. This indicates that the product of AT5G28237 belongs to a eudicot-conserved TSB type 1-like clade, divergent from that containing major experimentally characterized TSBs. A multiple sequence alignment (not shown) reveals that members of this divergent clade have a shortened N terminus with respect to the major chloroplast-localized TSB proteins. A localization prediction carried out in CELLO (Yu et al., 2006) suggests a cytosolic location for these proteins. Examination of EST databases indicates that the genes encoding these proteins are expressed. It is possible that the product of AT5G28237 could interact with the cytosolic INS studied by Wang et al. (2015), or separately with its indole product, to produce Trp that is further converted to IAA.Open in a separate windowFigure 2.Phylogeny of TSB type 1 homologs from Oryza sativa (LOC_Os), Sorghum bicolor (Sobic), Z. mays (GRMZM), Arabidopsis (AT), Brassica rapa (Brara), Solanum lycopersicum (Solyc), Populus trichocarpa (Potri), and Physcomitrella patens (Phpat). Protein sequences were downloaded from Phytozome v10.2 (Goodstein et al., 2012). The phylogenetic analysis was conducted in MEGA6 (http://megasoftware.net; Tamura et al., 2013) with multiple sequence alignment by MUSCLE (Edgar, 2004) and evolutionary history inferred using the neighbor-joining method (Saitou and Nei, 1987). The optimal tree is shown; the percentage of replicate trees in which the associated sequences clustered together in the bootstrap test (500 replicates) is shown next to the branches (Felsenstein, 1985). The tree is drawn to scale; the scale bar indicates the number of amino acid substitutions per site. It is rooted with type 1 TSBs from the moss P. patens.The second major line of evidence for Trp-independent IAA synthesis comes from isotopic labeling experiments. Wright et al. (1991) observed greater incorporation of ²H into IAA than Trp in orp seedlings grown on ²H₂O. Normanly et al. (1993) reported higher enrichment of ¹⁵N in IAA than Trp in trp2-1 mutants of Arabidopsis grown on ¹⁵N anthranilate; very poor incorporation of deuterium from ²H-Trp into IAA was reported in the trp2-1 plants. A number of similar reports relating to other plants have been published showing differences in the incorporation of label from Trp into IAA depending on experimental tissue and environmental conditions (e.g. Michalczuk et al., 1992; Rapparini et al., 2002; Sztein et al., 2002). This evidence has been persuasive; however, it assumes a single pool of Trp to which ¹⁵N anthranilate and ²H-Trp contribute and from which IAA is made. If Trp is made at different rates in different parts of the plant, and/or exogenous ²H-Trp does not equilibrate with newly synthesized Trp, then the ratio of ¹⁵N to ²H in Trp will vary in different plant organs/tissues/cells. Trp turnover and thus incorporation of label from ¹⁵N anthranilate are likely to differ substantially throughout the plant, with the highest rates of labeling occurring in cells with high rates of protein synthesis. This would not be a problem for the experiment if IAA is made at equal rates in different parts of the plant, but we know it is not. The Trp aminotransferase/YUCCA pathway of IAA synthesis elegantly shown to be responsible for the bulk of IAA synthesis (Mashiguchi et al., 2011; Won et al., 2011) appears to be locally controlled in Arabidopsis via 11 different YUCCA-encoding genes that have highly localized expression (Zhao, 2008). Adding to the complication is the need for ¹⁵N anthranilate and ²H-Trp to move into and through the plant to regions of Trp and IAA synthesis, respectively. This is likely to occur at different rates due to differing transporter requirements.Data from my own laboratory (Cooney and Nonhebel, 1991) is particularly relevant to this discussion. We monitored incorporation of ²H from deuterated water into IAA and Trp in tomato (S. lycopersicum) shoots. Unlike the other studies, we also measured the incorporation of label into IPA. Our data showed that IPA became labeled at a rate consistent with this compound acting as the major/sole precursor of IAA. Crucially, the proportion of labeled Trp was lower than ²H-IPA. Our interpretation of these data was that IPA and IAA were produced from newly synthesized Trp, and that Trp was not uniformly labeled throughout the shoot. At the time, we suggested different subcellular pools of Trp; this may be the case, but in light of new knowledge of localized IAA synthesis, it is most likely that substantial differences in Trp and IAA turnover in different cells/tissues may be the reason for these observations.The arguments above cast doubt on the existence of the Trp-independent route; however, a recent publication by Wang et al. (2015) claims to provide new evidence for its importance. They present the interesting finding that Arabidopsis plants with a null mutation in INS, a cytosolic TSA homolog previously shown to have indole-3-glycerol phosphate lyase (IGL) activity (Zhang et al., 2008), had reduced levels of IAA. The mutation particularly affected early embryo development. I suggest that the INS may make a contribution to IAA synthesis, but the only specific evidence that it does so via a Trp-independent route is the observation that the ins-1 mutation has an additive effect with the weakly ethylene insensitive8-1 Trp aminotransferase mutation. This evidence is indicative rather than conclusive. The possibility that INS may act in concert with a minor TSB homolog, as suggested in Figure 1, needs to be considered.In addition, Wang et al. (2015) focus on Arabidopsis alone. If INS has a key role in IAA synthesis, then evolutionary theory predicts a conserved protein with wide taxonomic distribution. On the contrary, an exhaustive BLAST search (Altschul et al., 1997) of diverse taxa in Phytozome v10.2 (Goodstein et al., 2012) and GenBank (Benson et al., 2013) revealed that INS orthologs with cytosolic prediction and shortened N terminus occur only in members of the Brassicaceae (Eutrema salsugineum, Arabis alpina, Camelina sativa, Capsella rubella, Brassica napus, Boechera stricta, Arabidopsis lyrata, B. rapa) and in Tarenaya hassleriana from the Brassicaceae sister family, the Cleomaceae. The phylogenetic tree in Figure 3 shows relationships between INS and TSA homologs from several plant species and indicates the separate clade of cytosolic INS homologs in the Brassicaceae. In this diagram, the sequence most closely related to INS from another group is that from tomato. This protein is the only TSA found in tomato and has an unambiguous chloroplast signal peptide. P. trichocarpa and M. truncatula as well as other eudicots outside the Brassicaeae and Cleomaceae also lack cytosolic TSA homologs. Furthermore, INS and its orthologs are phylogenetically distinct from the other experimentally characterized indole-3-glycerol phosphate lyases benzoxazin1 and IGL (Frey et al., 2000) and their orthologs. The latter are restricted to the grasses where they are involved in the production of cyclic hydroxamic acid defense compounds (Frey et al., 2000). The grasses also have an additional separate clade of cytosolic TSA homologs, although work by Kriechbaumer et al. (2008) did not detect any catalytic activity for the product of GRMZM2G046191_T01. The phylogeny of INS and its orthologs would suggest the major role of these proteins may be the production of lineage-specific metabolites such as the indole-derived defense compounds produced in grasses; any role in IAA synthesis may be incidental and restricted to the Brassicaeae and Cleomaceae.Open in a separate windowFigure 3.Phylogeny of TSA homologs from O. sativa (LOC_Os), S. bicolor (Sobic), Z. mays (GRMZM), Arabidopsis (AT), A. lyrata (Alyrata), B. rapa (Brara), S. lycopersicum (Solyc), Medicago truncatula (Medtr), P. trichocarpa (Potri), and P. patens (Phpat). Protein sequences were downloaded from Phytozome v10.2 (Goodstein et al., 2012). The phylogenetic analysis was conducted in MEGA6 (Tamura et al., 2013) with multiple sequence alignment by MUSCLE (Edgar, 2004) and evolutionary history inferred using the neighbor-joining method (Saitou and Nei, 1987). The rooted optimal tree is shown; the percentage of replicate trees in which the associated sequences clustered together in the bootstrap test (500 replicates) is shown next to the branches (Felsenstein, 1985). The tree is drawn to scale; the scale bar indicates the number of amino acid substitutions per site. It is rooted with the TSA ortholog from the moss P. patens.In conclusion, I contend that experimental data relating to IAA synthesis in Arabidopsis, including that suggesting the involvement of a cytosolic INS, can be explained by the Trp-dependent IAA synthesis pathway. I show that INS and its orthologs are not found outside the Brassicaceae and a closely related sister clade; any alternative IAA synthesis pathway in which they may be involved is likely to have similar limited taxonomic occurrence. Furthermore, Arabidopsis and its relatives contain two additional TSB homologs that could convert free indole into Trp. Curiously, both of these proteins have a wider taxonomic distribution. A priority for further experimental work should be testing the involvement of minor TSB homologs in IAA synthesis, including the highly conserved type 2 TSBs as well as a eudicot-specific clade of possibly cytosolic type 1 TSBs. Work would also have to establish whether free indole exists in plants other than the Brassicaceae and the grasses. Finally, I argue that isotope-labeling experiments do not provide strong support for the Trp-independent route, as IAA production is highly localized. Previously published data from my laboratory clearly show that the main Trp-dependent IAA precursor IPA becomes more highly labeled from ²H₂O than Trp, even though the latter is produced from Trp in a single reaction. Thus, it cannot be argued that differences in isotope enrichment between Trp and IAA demonstrate the existence of a Trp-independent route.     

A New LxxxA Motif in the Transmembrane Helix3 of Maize Aquaporins Belonging to the Plasma Membrane Intrinsic Protein PIP2 Group Is Required for Their Trafficking to the Plasma Membrane

Aquaporins play important roles in maintaining plant water status under challenging environments. The regulation of aquaporin density in cell membranes is essential to control transcellular water flows. This work focuses on the maize (Zea mays) plasma membrane intrinsic protein (ZmPIP) aquaporin subfamily, which is divided into two sequence-related groups (ZmPIP1s and ZmPIP2s). When expressed alone in mesophyll protoplasts, ZmPIP2s are efficiently targeted to the plasma membrane, whereas ZmPIP1s are retained in the endoplasmic reticulum (ER). A protein domain-swapping approach was utilized to demonstrate that the transmembrane domain3 (TM3), together with the previously identified N-terminal ER export diacidic motif, account for the differential localization of these proteins. In addition to protoplasts, leaf epidermal cells transiently transformed by biolistic particle delivery were used to confirm and refine these results. By generating artificial proteins consisting of a single transmembrane domain, we demonstrated that the TM3 of ZmPIP1;2 or ZmPIP2;5 discriminates between ER and plasma membrane localization, respectively. More specifically, a new LxxxA motif in the TM3 of ZmPIP2;5, which is highly conserved in plant PIP2s, was shown to regulate its anterograde routing along the secretory pathway, particularly its export from the ER.Aquaporins are of major importance to plant physiology, being essential for the regulation of transcellular water movement during growth and development (Maurel et al., 2008; Gomes et al., 2009; Heinen et al., 2009; Prado and Maurel, 2013; Chaumont and Tyerman, 2014). Aquaporins are small membrane proteins consisting of six transmembrane (TM) domains connected by five loops (A–E), and N and C termini facing the cytosol (Fig. 1A). They assemble as homotetramers and/or heterotetramers in the membrane, with each monomer acting as an independent water channel (Murata et al., 2000; Fetter et al., 2004; Gomes et al., 2009). Aquaporins form a highly divergent protein family in plants (Chaumont et al., 2001; Johanson et al., 2001), and this work focuses on the maize (Zea mays) plasma membrane intrinsic protein (ZmPIP) family (Chaumont et al., 2001). The regulation of the subcellular localization of these proteins is a key process controlling their density in the plasma membrane (PM) and, hence, their physiological roles (Hachez et al., 2013).Open in a separate windowFigure 1.Swapping TM3 of ZmPIP2;5 with that of ZmPIP1;2 retains the protein in intracellular structures. A, Cartoons representing the chimeric proteins composed of ZmPIP2;5, in which each TM has been replaced by the corresponding TM from ZmPIP1;2. All proteins are drawn with the cytosolic domains facing down. ZmPIP2;5 and ZmPIP1;2 portions are shown in black and white, respectively. All chimeras were fused to the C terminus of mYFP, which is not displayed for clarity purposes. B, Confocal microscopy images of maize mesophyll protoplasts transiently coexpressing mYFP-tagged ZmPIP2;5-PIP1;2 TM chimeric proteins (green) and the ER marker mCFP:HDEL (cyan). FM4-64 was added as a PM marker (red). Arrowheads in image 13 indicate accumulation of the protein in punctate structures that are not labeled by mCFP:HDEL. The localization patterns of the proteins of interest are representative of a total of at least 22 cells from three independent experiments. C, Confocal microscopy images of a maize mesophyll protoplast transiently expressing mYFP:ZmPIP2;5-TM3_PIP1;2 (green) and ST:mCFP (magenta). Arrowheads indicate colocalization in Golgi stacks. The images are representative of a total of 17 cells from two independent experiments. Bar = 5 µm.PIP aquaporins cluster in two groups (PIP1s and PIP2s), which are highly conserved across species (Kammerloher et al., 1994; Chaumont et al., 2000, 2001; Johanson et al., 2001; Anderberg et al., 2012). We previously showed that the maize PIP1 and PIP2 isoforms exhibit different water channel activities when expressed in Xenopus laevis oocytes, with only PIP2s increasing the membrane water permeability coefficient (P_f; Chaumont et al., 2000). However, when ZmPIP1 and ZmPIP2 are coexpressed, the isoforms physically interact to modify their stability and trafficking to the oocyte membrane, and synergistically increase the oocyte P_f (Fetter et al., 2004). Similar synergistic interactions between PIP1s and PIP2s have been reported in numerous plant species (Temmei et al., 2005; Mahdieh et al., 2008; Vandeleur et al., 2009; Bellati et al., 2010; Ayadi et al., 2011; Horie et al., 2011; Yaneff et al., 2014).PIPs were originally thought to be exclusively localized in the PM and were named accordingly (Kammerloher et al., 1994). However, recent experiments have shown that not all PIPs are located to the PM under all conditions, and that regulation of PIP subcellular localization is a highly dynamic process involving protein interactions (Boursiac et al., 2005, 2008; Zelazny et al., 2007, 2009; Uehlein et al., 2008; Besserer et al., 2012; Luu et al., 2012). When expressed singly in maize leaf mesophyll protoplasts, fluorescently tagged ZmPIP1s and ZmPIP2s differ in their subcellular localization. ZmPIP1s are retained in the endoplasmic reticulum (ER), whereas ZmPIP2s are targeted to the PM (Zelazny et al., 2007). However, upon coexpression, ZmPIP1s are relocalized from the ER to the PM, where they perfectly colocalize with ZmPIP2s. This relocalization results from their physical interaction as demonstrated by Förster resonance energy transfer/fluorescence lifetime imaging microscopy and immunoprecipitation experiments (Zelazny et al., 2007). These results indicate that ZmPIP2s, but not ZmPIP1s, possess signals that allow them to be delivered to the PM, and that hetero-oligomerization is required for ZmPIP1 trafficking to the PM. Interestingly, a diacidic motif (DxE, Asp-any amino acid-Glu) located in the N terminus of ZmPIP2;4, ZmPIP2;5, and Arabidopsis (Arabidopsis thaliana) AtPIP2;1 was shown to be required to exit the ER (Zelazny et al., 2009; Sorieul et al., 2011). Diacidic motifs interact with Secretory protein24, which is thought to be the main cargo-selection protein of the Coat proteinII complex that mediates vesicle formation at ER export sites (Miller et al., 2003). However, not all PM-localized PIP2s contain a diacidic ER export signal (Zelazny et al., 2009). In addition, swapping the N-terminal region of ER-retained ZmPIP1;2 with that of PM-localized ZmPIP2;5, which contains the functional diacidic motif, is not sufficient to trigger ER export of the protein (Zelazny et al., 2009). This result suggests that other export signals might be present in PIP2s and/or ER retention signals might be present in PIP1s elsewhere than in the N terminus.To identify new signals regulating ZmPIP1 and ZmPIP2 protein trafficking along the secretory pathway, we used a protein domain swapping-based approach and identified the TM3 as an important region that discriminates between ER-retained ZmPIP1;2 and PM-localized ZmPIP2;5. Specific mutations in the TM3 region of ZmPIP2;5 allowed the identification of a new ZmPIP2-conserved LxxxA motif, which regulates its export from the ER.     

The Presence of Fucogalactoxyloglucan and Its Synthesis in Rice Indicates Conserved Functional Importance in Plants

The predominant structure of the hemicellulose xyloglucan (XyG) found in the cell walls of dicots is a fucogalactoXyG with an XXXG core motif, whereas in the Poaceae (grasses and cereals), the structure of XyG is less xylosylated (XXGG_n core motif) and lacks fucosyl residues. However, specialized tissues of rice (Oryza sativa) also contain fucogalactoXyG. Orthologous genes of the fucogalactoXyG biosynthetic machinery of Arabidopsis (Arabidopsis thaliana) are present in the rice genome. Expression of these rice genes, including fucosyl-, galactosyl-, and acetyltransferases, in the corresponding Arabidopsis mutants confirmed their activity and substrate specificity, indicating that plants in the Poaceae family have the ability to synthesize fucogalactoXyG in vivo. The data presented here provide support for a functional conservation of XyG structure in higher plants.The plant cell wall protects and structurally supports plant cells. The wall consists of a variety of polymers, including polysaccharides, the polyphenol lignin, and glycoproteins. One of the major polysaccharides present in the primary walls (i.e. walls of growing cells) in dicots is xyloglucan (XyG), which consists of a β-1,4-glucan backbone with xylosyl substituents. XyG binds noncovalently to cellulose microfibrils and thereby, is thought to act as a spacer molecule, hindering cellulose microfibrils to aggregate (Carpita and Gibeaut, 1993; Pauly et al., 1999a; Bootten et al., 2004; Cosgrove, 2005; Hayashi and Kaida, 2011; Park and Cosgrove, 2012).The side-chain substitutions on XyG can be structurally diverse depending on plant species, tissue type, and developmental stage of the tissue (Pauly et al., 2001; Hoffman et al., 2005; Peña et al., 2008; Hsieh and Harris, 2009, 2012; Lampugnani et al., 2013; Schultink et al., 2014). A one-letter code nomenclature has been established to specify the XyG side-chain substitutions (Fry et al., 1993; Tuomivaara et al., 20145). According to this nomenclature, an unsubstituted glucosyl residue is indicated by a G, whereas a glucosyl residue substituted with a xylosyl moiety is shown as an X. In most dicots, the xylosyl residue can be further substituted with a galactosyl residue (L), which in turn, can be further decorated with a fucosyl residue (F) and/or an acetyl group (F/L). In some species, the xylosyl residue can be substituted with an arabinosyl moiety (S), and the backbone glucosyl residue can be O-acetylated (G; Jia et al., 2003; Hoffman et al., 2005).Numerous genes have been identified in Arabidopsis (Arabidopsis thaliana) that are involved in fucogalactoXyG biosynthesis (Fig. 1; Pauly et al., 2013; Schultink et al., 2014). The glucan backbone is thought to be synthesized by cellulose synthase-like C (CSLC) family proteins, such as AtCSLC4, as shown by in vitro activity data (Cocuron et al., 2007). Several xylosyltransferases (XXTs) from glycosyl transferase family 34 (GT34) are thought to be responsible for XyG xylosylation. Five of these XXTs in Arabidopsis seem to have XXT activity on XyG in vitro (Faik et al., 2002; Zabotina et al., 2008; Vuttipongchaikij et al., 2012; Mansoori et al., 2015). MURUS3 (MUR3) represents a galactosyltransferase that transfers galactosyl moieties specifically to xylosyl residues adjacent to an unsubstituted glucosyl residue on an XXXG unit, converting it to XXLG, whereas Xyloglucan L-side chain galactosyl Transferase2 (XLT2) was identified as another galactosyltransferase transferring a galactosyl moiety specifically to the second xylosyl residue, resulting in XLXG (Madson et al., 2003; Jensen et al., 2012). Both MUR3 and XLT2 belong to GT47 (Li et al., 2004). MUR2/FUCOSYLTRANSFERASE1 (FUT1) from GT37 was found to harbor fucosyltransferase activity, transferring Fuc from GDP-Fuc to a galactosyl residue adjacent to the unsubstituted glucosyl residue (i.e. onto XXLG but not onto XLXG; Perrin et al., 1999; Vanzin et al., 2002). O-acetylation of the galactosyl residue is mediated by Altered Xyloglucan4 (AXY4) and AXY4L, both of which belong to the Trichome Birefringence-Like (TBL) protein family (Bischoff et al., 2010; Gille et al., 2011; Gille and Pauly, 2012).Open in a separate windowFigure 1.Schematic structures of two types of XyGs and known biosynthetic proteins in Arabidopsis (Hsieh and Harris, 2009; Pauly et al., 2013). The corresponding one-letter code for XyG is shown below the pictograms (Fry et al., 1993; Tuomivaara et al., 2015).XyG found throughout land plants exhibits structural diversity with respect to side-chain substitution patterns (Schultink et al., 2014). Most dicots, such as Arabidopsis, and the noncommelinoid monocots possess a fucogalactoXyG of the XXXG-type XyG structure as shown in Figure 1. However, plant species in the Solanaceae and Poaceae as well as the moss Physcomitrella patens contain a different XyG structure with a reduced level of xylosylation, resulting in an XXGG_n core motif (York et al., 1996; Kato et al., 2004; Gibeaut et al., 2005; Jia et al., 2005; Peña et al., 2008; Hsieh and Harris, 2009). In addition, the glucan backbone can be O-acetylated in plants of Solanaceae and Poaceae families (Gibeaut et al., 2005; Jia et al., 2005). XyG from Solanaceae with an XXGG core motif can be further arabinosylated and/or galactosylated (Jia et al., 2005). No XyGs with an XXGG_n motif backbone have been reported to be fucosylated.The function of structural diversity of XyG substitutions, such as fucosylation and/or altered xylosylation pattern, remains enigmatic. Removing the terminal fucosyl or acetyl moieties in the corresponding Arabidopsis mutants does not lead to any change in plant growth and development (Vanzin et al., 2002; Gille et al., 2011). However, removing galactosyl residues as well as fucosyl and acetyl moieties in the Arabidopsis xlt2 mur3.1 double mutant results in a dwarfed plant (Jensen et al., 2012; Kong et al., 2015). Replacing the galactosyl moiety with an arabinofuranosyl residue by, for example, expressing a tomato (Solanum lycopersicum) arabinosyltransferase in the Arabidopsis xlt2 mur3.1 mutant rescues the growth phenotype and restores wall biomechanics, indicating that galactosylation and arabinosylation in XyG have an equivalent function (Schultink et al., 2013). Recently, fucosylated XyG structures were found in the pollen tubes of tobacco (Nicotiana alata) and tomato, indicating that fucogalactoXyG is likely also present in other Solanaceae plants, albeit restricted to specific tissues (Lampugnani et al., 2013; Dardelle et al., 2015). Although there is circumstantial evidence that fucogalactoXyG is present in cell suspension cultures of rice (Oryza sativa) and cell suspension cultures of fescue (Festuca arundinaceae; McDougall and Fry, 1994; Peña et al., 2008), fucogalactoXyG has not been found in any physiologically relevant plant tissues of members of the Poaceae (Kato et al., 1982; Watanabe et al., 1984; Gibeaut et al., 2005; Hsieh and Harris, 2009; Brennan and Harris, 2011). Here, we provide chemical and genetic evidence that fucogalactoXyG is, indeed, present in plant tissues of a grass (rice) and prove that the rice genome harbors the genes that could be part of the synthetic machinery necessary to produce fucogalactoXyG.     

The Arabidopsis Chloroplast Stromal N-Terminome: Complexities of Amino-Terminal Protein Maturation and Stability

Protein amino (N) termini are prone to modifications and are major determinants of protein stability in bacteria, eukaryotes, and perhaps also in chloroplasts. Most chloroplast proteins undergo N-terminal maturation, but this is poorly understood due to insufficient experimental information. Consequently, N termini of mature chloroplast proteins cannot be accurately predicted. This motivated an extensive characterization of chloroplast protein N termini in Arabidopsis (Arabidopsis thaliana) using terminal amine isotopic labeling of substrates and mass spectrometry, generating nearly 14,000 tandem mass spectrometry spectra matching to protein N termini. Many nucleus-encoded plastid proteins accumulated with two or three different N termini; we evaluated the significance of these different proteoforms. Alanine, valine, threonine (often in N-α-acetylated form), and serine were by far the most observed N-terminal residues, even after normalization for their frequency in the plastid proteome, while other residues were absent or highly underrepresented. Plastid-encoded proteins showed a comparable distribution of N-terminal residues, but with a higher frequency of methionine. Infrequent residues (e.g. isoleucine, arginine, cysteine, proline, aspartate, and glutamate) were observed for several abundant proteins (e.g. heat shock proteins 70 and 90, Rubisco large subunit, and ferredoxin-glutamate synthase), likely reflecting functional regulation through their N termini. In contrast, the thylakoid lumenal proteome showed a wide diversity of N-terminal residues, including those typically associated with instability (aspartate, glutamate, leucine, and phenylalanine). We propose that, after cleavage of the chloroplast transit peptide by stromal processing peptidase, additional processing by unidentified peptidases occurs to avoid unstable or otherwise unfavorable N-terminal residues. The possibility of a chloroplast N-end rule is discussed.Following synthesis, most proteins undergo various N-terminal (Nt) protein modifications, including removal of the Nt Met and signal peptide, N-terminal α-acetylation (NAA), ubiquitination, and acylations. These Nt modifications play an important role in the regulation of cellular functions. The N terminus of proteins has also been shown to be a major determinant of protein stability in bacteria (Varshavsky, 2011), eukaryotes (Graciet et al., 2009), mitochondria, and perhaps in plastids/chloroplasts (Apel et al., 2010; Nishimura et al., 2013; van Wijk, 2015). The role of the N terminus in protein stability is conceptualized in the N-end rule, which states that certain amino acids, when exposed at the N terminus of a protein, act as triggers for degradation (Bachmair et al., 1986; Dougan et al., 2012; Tasaki et al., 2012; Gibbs et al., 2014).Most of the approximately 3,000 plastid proteins are nucleus encoded (n-encoded) and are targeted to the plastid through an Nt chloroplast transit peptide (cTP). After import, the cTP is cleaved by the stromal processing peptidase (SPP; Richter and Lamppa, 1998; Trösch and Jarvis, 2011). The consensus site of cTP cleavage by SPP is only loosely defined, and the rules, mechanisms, and enzymes for possible subsequent processing, stabilization, and other posttranslational modifications (PTMs) are not well characterized (for discussion, see van Wijk, 2015). The exact N terminus is unknown for many chloroplast proteins and cannot be accurately predicted, because SPP specificity is not sufficiently understood (Emanuelsson et al., 2000; Zybailov et al., 2008) and probably also because additional Nt processing occurs for many chloroplast proteins (Fig. 1A). The approximately 85 plastid-encoded (p-encoded) proteins typically first undergo cotranslational Nt deformylation, followed by N-terminal Met excision (NME; Giglione et al., 2009; Fig. 1B); both these PTMs are required for normal plastid/chloroplast development and protein stability (Dirk et al., 2001, 2002; Giglione et al., 2003; Meinnel et al., 2006). Both n-encoded and p-encoded proteins can undergo NAA inside the plastid (Zybailov et al., 2008; Fig. 1). Postulated functions of NAA in eukaryotes include the mediation of protein location, assembly, and stability (Jones and O’Connor, 2011; Starheim et al., 2012; Hoshiyasu et al., 2013; Xu et al., 2015), thereby affecting a variety of processes, including drought tolerance in Arabidopsis (Arabidopsis thaliana; Linster et al., 2015).Open in a separate windowFigure 1.Conceptual illustration of Nt maturation of n-encoded and p-encoded proteins. Ac, Acetylated; MAP, Met amino peptidase; NAT, N-acetyltransferase; N-term, N-terminal; PDF, peptide deformylase. A, Nt maturation of n-encoded plastid proteins including removal of cTP by SPP and potential subsequent Nt modifications. B, Nt maturation of p-encoded proteins. *, The removal depends on the penultimate residue, generally following the N-terminal Met Excision (NME) rule; **, N-terminal acetylation typically occurs only for selected residues; “Results”).Typical proteomics work flows generally yield only partial coverage of protein sequences, and it is often difficult to know which peptides represent the true N termini (Nti) or C termini. Systematic identification of Nti or C termini requires specific labeling and enrichment strategies, such as combined fractional diagonal chromatography, developed by Gevaert and colleagues (Staes et al., 2011), and terminal amine isotopic labeling of substrates (TAILS), developed by the group of Overall (Kleifeld et al., 2011; Lange and Overall, 2013). These strategies allow the identification of different Nt proteoforms and were recently also applied to plants (Tsiatsiani et al., 2013; Carrie et al., 2015; Kohler et al., 2015; Zhang et al., 2015) and diatoms (Huesgen et al., 2013). We previously reported on Nti of chloroplast proteins based on tandem mass spectrometry (MS/MS) analysis, but because no Nt enrichment/labeling technique was used, only those that underwent NAA could be considered bona fide Nti (Zybailov et al., 2008). Nt Edman degradation sequencing was systematically carried out for thylakoid lumen proteins (Peltier et al., 2000, 2002) but not for stromal proteins or chloroplast membrane proteins with their Nti exposed to the stroma. The Nti of thylakoid lumen proteins are mostly generated by lumenal peptidases (Hsu et al., 2011; Midorikawa et al., 2014), and the thylakoid lumen contains a different set of peptidases than the stroma; hence, rules for Nt maturation and stability are likely different than those for stroma-exposed proteins.The objective of this study was to systematically determine the Nti of stroma-exposed chloroplast proteins of Arabidopsis (the N-terminome) and to provide a baseline for understanding Nt protein maturation and protein stability in the chloroplast stroma. To that end, we applied the TAILS technique and determined the Nti of approximately 250 chloroplast proteins by mass spectrometry (MS). We observed that many n-encoded plastid proteins accumulated with two or even three different Nt residues, in many cases both with and without NAA. The extent of accumulation of different Nt proteoforms is surprising and will be discussed. The p-encoded proteins generally showed very similar Nt residues as compared with the n-encoded proteins, with the exception of Met. Our data show that small, apolar, or hydroxylated residues (Ala, Val, Ser, and Thr) are the most frequent Nt residues of stromal proteins, whereas other residues are strictly avoided or are only present for very specific proteins likely to aid in their function. Chloroplast protein degradation products were also detected, with enrichment for peptides generated by cleavage between Arg and Thr residues. We present testable hypotheses for understanding Nt processing and maturation, stability, and a possible N-end rule in chloroplast stroma.