Why have chloroplasts developed a unique motility system?

Organelle movement in plants is dependent on actin filaments with most of the organelles being transported along the actin cables by class XI myosins. Although chloroplast movement is also actin filament-dependent, a potential role of myosin motors in this process is poorly understood. Interestingly, chloroplasts can move in any direction and change the direction within short time periods, suggesting that chloroplasts use the newly formed actin filaments rather than preexisting actin cables. Furthermore, the data on myosin gene knockouts and knockdowns in Arabidopsis and tobacco do not support myosins'' XI role in chloroplast movement. Our recent studies revealed that chloroplast movement and positioning are mediated by the short actin filaments localized at chloroplast periphery (cp-actin filaments) rather than cytoplasmic actin cables. The accumulation of cp-actin filaments depends on kinesin-like proteins, KAC1 and KAC2, as well as on a chloroplast outer membrane protein CHUP1. We propose that plants evolved a myosin XI-independent mechanism of the actin-based chloroplast movement that is distinct from the mechanism used by other organelles.Key words: actin, Arabidopsis, blue light, kinesin, myosin, organelle movement, phototropinOrganelle movement and positioning are pivotal aspects of the intracellular dynamics in most eukaryotes. Although plants are sessile organisms, their organelles are quickly repositioned in response to fluctuating environmental conditions and certain endogenous signals. By and large, plant organelle movements and positioning are dependent on actin filaments, although microtubules play certain accessory roles in organelle dynamics.^1,2 Actin inhibitors effectively retard the movements of mitochondria,^3–6 peroxisomes,^5,7–11 Golgi stacks,^12,13 endoplasmic reticulum (ER),^14,15 and nuclei.^16–18 These organelles are co-aligned and associated with actin filaments.^{5,7,8,10–12,15,18} Recent progress in this field started to reveal the molecular motility system responsible for the organelle transport in plants.¹⁹Chloroplast movement is among the most fascinating models of organelle movement in plants because it is precisely controlled by ambient light conditions.^20,21 Weak light induces chloroplast accumulation response so that chloroplasts can capture photosynthetic light efficiently (Fig. 1A). Strong light induces chloroplast avoidance response to escape from photodamage (Fig. 1B).²² The blue light-induced chloroplast movement is mediated by the blue light receptor phototropin (phot). In some cryptogam plants, the red light-induced chloroplast movement is regulated by a chimeric phytochrome/phototropin photoreceptor neochrome.^23–25 In a model plant Arabidopsis, phot1 and phot2 function redundantly to regulate the accumulation response,²⁶ whereas phot2 alone is essential for the avoidance response.^27,28 Several additional factors regulating chloroplast movement were identified by analyses of Arabidopsis mutants deficient in chloroplast photorelocation.^29–32 In particular, identification of CHUP1 (chloroplast unusual positioning 1) revealed the connection between chloroplasts and actin filaments at the molecular level.²⁹ CHUP1 is a chloroplast outer membrane protein capable of interacting with F-actin, G-actin and profilin in vitro.^29,33,34 The chup1 mutant plants are defective in both the chloroplast movement and chloroplast anchorage to the plasma membrane,^22,29,33 suggesting that CHUP1 plays an important role in linking chloroplasts to the plasma membrane through the actin filaments. However, how chloroplasts move using the actin filaments and whether chloroplast movement utilizes the actin-based motility system similar to other organelle movements remained to be determined.Open in a separate windowFigure 1Schematic distribution patterns of chloroplasts in a palisade cell under different light conditions, weak (A) and strong (B) lights. Shown as a side view of mid-part of the cell and a top view with three different levels (i.e., top, middle and bottom of the cell). The cell was irradiated from the leaf surface shown as arrows. Weak light induces chloroplast accumulation response (A) and strong light induces the avoidance response (B).Here, we review the recent findings pointing to existence of a novel actin-based mechanisms for chloroplast movement and discuss the differences between the mechanism responsible for movement of chloroplasts and other organelles.     

The galactolipid monogalactosyldiacylglycerol (MGDG) contributes to photosynthesis-related processes in Arabidopsis thaliana

Processes putatively dependent on the galactolipid monogalactosyldiacylglycerol (MGDG) were recently studied using the knockdown monogalactosyldiacylglycerol synthase 1 (mgd1-1) mutant (∼40% reduction in MGDG). Surprisingly, targeting of chloroplast proteins was not affected in mgd1-1 mutants, suggesting they retain sufficient MGDG to maintain efficient targeting. However, in dark-grown mgd1-1 plants the photoactive to photoinactive protochlorophyllide (Pchlide) ratio was increased, suggesting that photoprotective responses are induced in them. Nevertheless, mgd1-1 could not withstand high light intensities, apparently due to impairment of another photoprotective mechanism, the xanthophyll cycle (and hence thermal dissipation). This was mediated by increased conductivity of the thylakoid membrane leading to a higher pH in the thylakoid interior, which impaired the pH-dependent activation of violaxanthin de-epoxidase (VDE) and PsbS. These findings suggest that MGDG contribute directly to the regulation of photosynthesis-related processes.Key words: conductivity, galactolipid, light stress, photosynthesis, plastid, xanthophyllThe galactolipid monogalactosyldiacylglycerol (MGDG), the major lipid in plastids,¹ is mainly synthesised in inner plastid envelopes,² where monogalactosyldiacylglycerol synthase 1 (MGD1) catalyses the last step of its production.³ Two MGDG-deficient mutants are known: the knockdown mgd1-1 mutant, which accumulates ∼40% less MGDG than wild type,⁴ and the null mutant mgd1-2, which displays extremely severe defects in chloroplast and plant development.⁵ Thus, the mgd1-1 mutant is more suitable for assessing putative roles of MGDG in processes such as protein targeting and photoprotection.There are conflicting indications regarding the involvement of galactolipids in chloroplast protein targeting: some suggest they play an important role,^6–10 but not all.^11,12 The data recently collected for mgd1-1 do not support MGDG''s involvement in protein targeting, since (inter alia) the level of MGDG in mgd1-1 mutants is clearly sufficient for efficient targeting.¹³ Further, the galactolipid associated with the TOC complex¹² is digalactosyldiacylglycerol (DGDG) and the digalactosyldiacylglycerol synthase 1 (dgd1) mutant,¹⁴ which has ∼10% of wild-type levels of DGDG, has impaired import efficiency.^15,16 Hence, this may indicate that DGDG is relatively more important for chloroplast import than MGDG.The prolamellar bodies (PLBs) of etioplasts have high lipid-to-protein ratios compared to thylakoids. Their major lipid and protein are MGDG and NADPH:Pchlide oxidoreductase (POR), respectively,¹⁷ and MGDG putatively plays an important role, interactively with POR, in the formation of PLBs.^18–20 The transformation of PLBs into thylakoids involves phototransformation of photoactive Pchlide (F656), a precursor of chlorophyll. Non-photoactive Pchlide (F631) is susceptible to photooxidative damage, but POR is believed to suppress this.^21,22 After excitation at 440 nm, mgd1-1 mutants display distinctly higher fluorescence emission peaks corresponding to photoactive Pchlide than wild type counterparts and (hence) higher photoactive:non-photoactive Pchlide ratios.¹³ These changes may be photoprotective responses that favour formation of photoactive Pchlide and optimize the plants'' opportunities to use light for chlorophyll production, enabling the transformation of etioplasts into chloroplasts.Interestingly,the xanthophyll cycle, another photoprotective mechanism, is impaired in mgd1-1.¹³ Normally, the xanthophyll cycle pigment violaxanthin is de-epoxidized into antheraxanthin, and then into zeaxanthin, by the enzyme VDE (Fig. 1), which is dependent on MGDG.²³ MGDG is also an integral component of photosynthetic complexes.^24–26 Thus, since mgd1-1 mutants have reduced total amounts of xanthophyll and chlorophyll pigments, but increased chlorophyll a/b ratios, their photosynthesis capacity is unsurprisingly reduced, even though the organization of their electron transport chains is not strongly affected by the MGDG deficiency.¹³Open in a separate windowFigure 1Reactions of the xanthophyll cycle (adapted from ref. ²⁹). VDE, violaxanthin de-epoxidase; ZE, zeaxanthin epoxidase.During short-term high light stress, antheraxanthin and zeaxanthin are thought to facilitate dissipation of excess light energy in the PSII antenna bed by non-photochemical quenching.^27,28 Upon high light stress the pH decreases, triggering photoprotective mechanisms via changes in the PSII antenna system. The PsbS protein, which is involved in thermal dissipation, is protonated and initiates a conformational change in the PSII antenna bed. This change is further stabilized by the de-epoxidation of violaxanthin to zeaxanthin by the luminal VDE.²⁸ However, the thermal dissipation is impaired in mgd1-1 mutants at high light intensities (>1000 µmol m⁻² s⁻¹) making them more susceptible to light stress. Surprisingly, this is not mediated by direct effects on VDE and PsbS activities, but by changes in the proton conductivity of the thylakoid membrane.¹³The steady-state capacity of the xanthophyll cycle is reduced in mgd1-1 mutants, due to a ∼40% reduction in the proton motive force (pmf) across their thylakoid membranes, indicating that they have impaired capacities to energize these membranes. Nevertheless, the pmf is more or less equal to wild type under light-limited conditions (200 µmol m⁻² s⁻¹ light); it is only the increase in pmf in high light intensities that is impaired in the mutants.¹³ This leads to the thylakoid lumen being less acidic in mgd1-1 than in wild type, hampering full activation of VDE and PsbS. Thus, the thylakoid lumen pH is above the threshold level required for full activation of PsbS and VDE under steady-state conditions and so de-epoxidation rates are retarded and the equilibrium between zeaxanthin and violaxanthin starts to shift slightly towards violaxanthin (Fig. 2).¹³ Thus, increased conductivity of the thylakoid membranes is probably responsible for the diminished non-photochemical quenching in mgd1-1, and the findings strongly indicate that MGDG is required for efficient photosynthesis and photoprotection, in addition to being a physical membrane constituent.Open in a separate windowFigure 2Schematic diagram illustrating the normal mode of action of the xanthophyll cycle. In standard light conditions, V is bound to the photosynthetic complexes and harvests light. In strong light, V is released from the complexes and converted to Z by VDE, which is unable to access V when it is associated with the photosynthetic complexes. The newly formed Z then binds to the photosynthetic complexes (at the PsbS protein), where it dissipates excess energy through NPQ. V, violaxanthin; A, antheraxanthin; Z, zeaxanthin; VDE, violaxanthin de-epoxidase; ZE, zeaxanthin epoxidase. Arrows indicate the directions of reactions.     

Prions: En route from structural models to structures

The prion hypothesis^1–3 states that the prion and non-prion form of a protein differ only in their 3D conformation and that different strains of a prion differ by their 3D structure.^4,5 Recent technical developments have enabled solid-state NMR to address the atomic-resolution structures of full-length prions, and a first comparative study of two of them, HET-s and Ure2p, in fibrillar form, has recently appeared as a pair of companion papers.^6,7 Interestingly, the two structures are rather different: HET-s features an exceedingly well-ordered prion domain and a partially disordered globular domain. Ure2p in contrast features a very well ordered globular domain with a conserved fold, and—most probably—a partially ordered prion domain.⁶ For HET-s, the structure of the prion domain is characterized at atomic-resolution. For Ure2p, structure determination is under way, but the highly resolved spectra clearly show that information at atomic resolution should be achievable.Key words: prion, NMR, solid-state NMR, MAS, structure, Ure2p, HET-sDespite the large interest in the basic mechanisms of fibril formation and prion propagation, little is known about the molecular structure of prions at atomic resolution and the mechanism of propagation. Prions with related properties to the ones responsible for mammalian diseases were also discovered in yeast and funghi^8,9 which provide convenient model system for their studies. Prion proteins described include the mammalian prion protein PrP, Ure2p,¹⁰ Rnq1p,¹¹ Sup35,¹² Swi1,¹³ and Cyc8,¹⁴ from bakers yeast (S. cervisiae) and HET-s from the filamentous fungus P. anserina. The soluble non-prion form of the proteins characterized in vitro is a globular protein with an unfolded, dynamically disordered N- or C-terminal tail.^15–18 In the prion form, the proteins form fibrillar aggregates, in which the tail adopts a different conformation and is thought to be the dominant structural element for fibril formation.Fibrills are difficult to structurally characterize at atomic resolution, as X-ray diffraction and liquid-state NMR cannot be applied because of the non-crystallinity and the mass of the fibrils. Solid-state NMR, in contrast, is nowadays well suited for this purpose. The size of the monomer, between 230 and 685 amino-acid residues for the prions of Figure 1, and therefore the number of resonances in the spectrum—that used to be large for structure determination—is now becoming tractable by this method.Open in a separate windowFigure 1Prions identified today and characterized as consisting of a prion domain (blue) and a globular domain (red).Prion proteins characterized so far were found to be usually constituted of two domains, namely the prion domain and the globular domain (see Fig. 1). This architecture suggests a divide-and-conquer approach to structure determination, in which the globular and prion domain are investigated separately. In isolation, the latter, or fragments thereof, were found to form β-sheet rich structures (e.g., Ure2p(1-89),^6,19 Rnq1p(153-405)²⁰ and HET-s(218-289)²¹). The same conclusion was reached by investigating Sup35(1-254).²² All these fragements have been characterized as amyloids, which we define in the sense that a significant part of the protein is involved in a cross-beta motif.²³ An atomic resolution structure however is available presently only for the HET-s prion domain, and was obtained from solid-state NMR²⁴ (vide infra). It contains mainly β-sheets, which form a triangular hydrophobic core. While this cross-beta structure can be classified as an amyloid, its triangular shape does deviate significantly from amyloid-like structures of smaller peptides.²³Regarding the globular domains, structures have been determined by x-ray crystallography (Ure2p^25,26 and HET-s²⁷), as well as NMR (mammal prions^15,28–30). All reveal a protein fold rich in α-helices, and dimeric structures for the Ure2 and HET-s proteins. The Ure2p fold resembles that of the β-class glutathione S-transferases (GST), but lacks GST activity.²⁵It is a central question for the structural biology of prions if the divide-and-conquer approach imposed by limitations in current structural approaches is valid. Or in other words: can the assembly of full-length prions simply be derived from the sum of the two folds observed for the isolated domains?     

MEK1/2 and p38-like MAP kinase successively mediate H2O2 signaling in Vicia guard cell

As a second messenger, H₂O₂ generation and signal transduction is subtly controlled and involves various signal elements, among which are the members of MAP kinase family. The increasing evidences indicate that both MEK1/2 and p38-like MAP protein kinase mediate ABA-induced H₂O₂ signaling in plant cells. Here we analyze the mechanisms of similarity and difference between MEK1/2 and p38-like MAP protein kinase in mediating ABA-induced H₂O₂ generation, inhibition of inward K⁺ currents, and stomatal closure. These data suggest that activation of MEK1/2 is prior to p38-like protein kinase in Vicia guard cells.Key words: H₂O₂ signaling, ABA, p38-like MAP kinase, MEK1/2, guard cellAn increasing number of literatures elucidate that reactive oxygen species (ROS), especially H₂O₂, is essential to plant growth and development in response to stresses,^1–4 and involves activation of various signaling events, among which are the MAP kinase cascades.^1–3,5 Typically, activation of MEK1/2 mediates NADPH oxidase-dependent ROS generation in response to stresses,^4,6–8 and the facts that MEK1/2 inhibits the expression and activation of antioxidant enzymes reveal how PD98059, the specific inhibitor of MEK1/2, abolishes abscisic acid (ABA)-induced H₂O₂ generation.^6,8,9 It has been indicated that PD98059 does not to intervene on salicylic acid (SA)-stimulated H₂O₂ signaling regardless of SA mimicking ABA in regulating stomatal closure.^2,6,8,10 Generally, activation of MEK1/2 promotes ABA-induced stomatal closure by elevating H₂O₂ generation in conjunction with inactivating anti-oxidases.Moreover, activation of plant p38-like protein kinase, the putative counterpart of yeast or mammalian p38 MAP kinase, has been reported to participate in various stress responses and ROS signaling. It has been well documented that p38 MAP kinase is involved in stress-triggered ROS signaling in yeast or mammalian cells.^11–13 Similar to those of yeast and mammals, many studies showed the activation of p38-like protein kinase in response to stresses in various plants, including Arabidopsis thaliana,^14–16 Pisum sativum,¹⁷ Medicago sativa¹⁸ and tobacco.¹⁹ The specific p38 kinase inhibitor SB203580 was found to modulate physiological processes in plant tissues or cells, such as wheat root cells,²⁰ tobacco tissue²¹ and suspension-cultured Oryza sativa cells.²² Recently, we investigate how activation of p38-like MAP kinase is involved in ABA-induced H₂O₂ signaling in guard cells. Our results show that SB203580 blocks ABA-induced stomatal closure by inhibiting ABA-induced H₂O₂ generation and decreasing K⁺ influx across the plasma membrane of Vicia guard cells, contrasting greatly with its analog SB202474, which has no effect on these events.^23,24 This suggests that ABA integrate activation of p38-like MAP kinase and H₂O₂ signaling to regulate stomatal behavior. In conjunction with SB203580 mimicking PD98059 not to mediate SA-induced H₂O₂ signaling,^23,24 these results generally reveal that the activation of p38-like MAP kinase and MEK1/2 is similar in guard cells.On the other hand, activation of p38-like MAP kinase^23,24 is not always identical to that of MEK1/2^8,25 in ABA-induced H₂O₂ signaling of Vicia guard cells. For example, H₂O₂^- and ABA-induced stomatal closure was partially reversed by SB203580. The maximum inhibition of both regent-induced stomatal closure were observed at 2 h after treatment with SB203580, under which conditions the stomatal apertures were 89% and 70% of the control values, respectively. By contrast, when PD98059 was applied together with ABA or H₂O₂, the effects of both ABA- and H₂O₂-induced stomatal closure were completely abolished (Fig. 1). These data imply that the two members of MAP kinase family are efficient in H₂O₂-stimulated stomatal closure, but p38-like MAP kinase is less susceptive than MEK1/2 to ABA stimuli.Open in a separate windowFigure 1Effects of SB203580 and PD98059 on ABA- and H₂O₂-induced stomatal closure. The experimental procedure and data analysis are according to the previous publication.^8,23,24It has been reported that ABA or NaCl activate p38 MAP kinase in the chloronema cells of the moss Funaria hygrometrica in 2∼10 min.²⁶ Similar to this, SB203580 improves H₂O₂-inhibited inward K⁺ currents after 4 min and leads it to the control level (100%) during the following 8 min (Fig. 2). However, the activation of p38-like MAP kinase in response to ABA need more time, and only recovered to 75% of the control at 8 min of treatment (Fig. 2). These results suggest that control of H₂O₂ signaling is required for the various protein kinases including p38-like MAP kinase and MEK1/2 in guard cells,^1,2,8,23,24 and the ABA and H₂O₂ pathways diverge further downstream in their actions on the K⁺ channels and, thus, on stomatal control. Other differences in action between ABA and H₂O₂ are known. For example, Köhler et al. (2001) reported that H₂O₂ inhibited the K⁺ outward rectifier in guard cells shows that H₂O₂ does not mimic ABA action on guard cell ion channels as it acts on the K⁺ outward rectifier in a manner entirely contrary to that of ABA.²⁷Open in a separate windowFigure 2Effect of SB203580 on ABA- and H₂O₂-inhibited inward K⁺ currents. The experimental procedure and data analysis are according to the previous publication.²⁴ SB203580 directs ABA- and H₂O₂-inactivated inward K⁺ currents across plasma membrane of Vicia guard cells. Here the inward K⁺ currents value is stimulated by −190 mV voltage.Based on the similarity and difference between PD98059 and SB203580 in interceding ABA and H₂O₂ signaling, we speculate the possible mechanism is that the member of MAP kinase family specially regulate signal event in ABA-triggered ROS signaling network,^1–4 and the signaling model as follows (Fig. 3).Open in a separate windowFigure 3Schematic illustration of MAP kinase-mediated H₂O₂ signaling of guard cells. The arrows indicate activation. The line indicates enhancement and the bar denotes inhibition.     

Mannan synthase activity in the CSLD family

Cellulose Synthase Like (CSL) proteins are a group of plant glycosyltransferases that are predicted to synthesize β-1,4-linked polysaccharide backbones. CSLC, CSLF and CSLH families have been confirmed to synthesize xyloglucan and mixed linkage β-glucan, while CSLA family proteins have been shown to synthesize mannans. The polysaccharide products of the five remaining CSL families have not been determined. Five CSLD genes have been identified in Arabidopsis thaliana and a role in cell wall biosynthesis has been demonstrated by reverse genetics. We have extended past research by producing a series of double and triple Arabidopsis mutants and gathered evidence that CSLD2, CSLD3 and CSLD5 are involved in mannan synthesis and that their products are necessary for the transition between early developmental stages in Arabidopsis. Moreover, our data revealed a complex interaction between the three glycosyltransferases and brought new evidence regarding the formation of non-cellulosic polysaccharides through multimeric complexes.Key words: mannan, mannose, plant cell wall, glycosyltransferase, cellulose synthase like, CSL, biosynthesis, hemicelluloseThe plant cell wall is mainly composed of polysaccharides, which are often grouped into cellulose, hemicelluloses and pectin. Since the discovery of the first cellulose synthase (CESA) genes in cotton fibers,¹ the synthesis of cellulose has been extensively studied.² In contrast, the glycosyltransferases responsible for synthesizing hemicelluloses and pectin are still largely unidentified.^3,4,5 The CESA genes are members of a superfamily that includes genes with a high sequence similarity with CESA genes and are named Cellulose Synthase Like (CSL).⁶ The CSL genes have themselves been grouped into nine families designated CSLA, -B, -C, -D, -E, -F, -G, -H and -J (Figure 1A).^5,6 Mannan and glucomannan synthase activity has been demonstrated in the CSLA family,^7,8,9 while members of the CSLC family have been implicated in synthesis of the xyloglucan backbone.¹⁰ CSLF and CSLH, which are found only in grasses, are involved in synthesis of mixed linkage glucan.^11,12 The function of the remaining CSL families has not been determined. We have reported our research on the CSLD family in a recent publication.¹³ Of all the CSL families, CSLD possesses the most ancient intron/exon structure and is the most similar to the CESA family.⁶ CSLD genes are found in all sequenced genomes of terrestrial plants including Physcomitrella and Selaginella suggesting a highly conserved function throughout the plant kingdom (Figure 1A). Five genes (CSLD1 to CSLD5) and one apparent pseudogene (CSLD6) have been identified in Arabidopsis thaliana.¹⁴ Bernal et al.^14,15 studied knock-out mutants of the individual genes and presented evidence for a role in cell wall biosynthesis for each Arabidopsis CSLD. To elucidate the activity of the CSLD proteins and obtain further understanding of their biological role, we generated double mutants csld2/csld3, csld2/csld5, csld3/csld5 and the triple mutant csld2/csld3/csld5. Immunochemical, biochemical and complementation assays brought evidence that CSLD5 or CSLD2 in concomitance with CSLD3 act as mannan synthases.Open in a separate windowFigure 1(A) Schematic representation of the CESA superfamily phylogeny. The inset on the right is a detailed phylogenetic tree of CSLDs from Selaginella moellendorffii, Arabidopsis thaliana and Oryza sativa. The figure is modified from Ulvskov and Scheller.⁵ (B) Comparison of csld2, csld3, csld5 with Col-0 20 days after germination. The inflorescences of csld2 and csld3 were similar to Col-0 whereas csld5 had a delayed growth. Scale bar: 1 cm. (C) Col-0 and csld2/csld3/csld5 (triple mutant, TM) 40 days after germination. After 40 days, the triple mutant was barely developed and, as shown in the magnified inset, displayed purple coloration indicating accumulation of anthocyanins, a typical stress response. Scale bar: 2 mm.     

Is membrane occupation and recognition nexus domain functional in plant phosphatidylinositol phosphate kinases?

Phosphatidylinositol phosphate kinase (PIPK) catalyzes a key step controlling cellular contents of phosphatidylinositol-4,5-bisphosphate [PtdIns(4,5)P₂], a critical intracellular messenger involved in vesicle trafficking and modulation of actin cytoskeleton and also a substrate of phospholipase C to produce the two intracellular messengers, diacylglycerol and inositol-1,4,5-trisphosphate. In addition to the conserved C-terminal PIPK catalytic domain, plant PIPKs contain a unique structural feature consisting of a repeat of membrane occupation and recognition nexus (MORN) motifs, called the MORN domain, in the N-terminal half. The MORN domain has previously been proposed to regulate plasma membrane localization and phosphatidic acid (PA)-inducible activation. Recently, the importance of the catalytic domain, but not the MORN domain, in these aspects was demonstrated. These conflicting data raise the question about the function of the MORN domain in plant PIPKs. We therefore performed analyses of PpPIPK1 from the moss Physcomitrella patens to elucidate the importance of the MORN domain in the control of enzymatic activity; however, we found no effect on either enzymatic activity or activation by PA. Taken together with our previous findings of lack of function in plasma membrane localization, there is no positive evidence indicating roles of the MORN domain in enzymatic and functional regulations of PpPIPK1. Therefore, further biochemical and reverse genetic analyses are necessary to understand the biological significance of the MORN domain in plant PIPKs.Key words: membrane occupation and recognition nexus (MORN) domain, phosphatidylinositol phosphate kinase, phosphatidic acid, Physcomitrella patensPhosphoinositides (PIs) are minor membrane phospholipds that play pivotal roles in various signal transduction cascades involved in development and stress response via the regulation of cytoskeletal organization, ion channel activation and vesicle trafficking.^1,2 These are derivatives of phosphatidylinositol (PtdIns) produced by phosphorylation of the 3-, 4- and 5- positions of the inositol ring.² To address the roles of PIs, enzymes involved in their production have been extensively studied using biochemical and molecular biological approaches. Of these enzymes, phosphatidylinositol monophosphate kinases (PIPKs) catalyze the reaction producing phosphatidylinositol-4,5-bisphosphate [PtdIns(4,5)P₂] that is a substrate of phospholipase C and phosphatidylinositol 3-kinase, and also acts as an intracellular messenger involved in the regulation of F-actin organization and activity of ion channels.^1–3 Although PtdIns(4,5)P₂ is produced by sequential phosphorylation by phosphatidylinositol 4-kinase, producing phosphatidylinositol-4-phosphate [PtdIns(4)P], and then by PIPK,^1,2 the cellular levels of PtdIns(4)P are much higher compared to PtdIns(4,5)P₂.^4–6 Thus, a restriction step controlling cellular PtdIns(4,5)P₂ contents is mediated by PIPKs, indicating the importance of PIPK regulation in various kinds of physiological processes.The roles of plant PIPKs have been established in growth regulation, such as polarized tip growth of root hairs and pollen tubes, via their localization at plasma membranes.^7–12 It is worth to note that plant PIPKs contain a unique structure consisting of a repeat of a membrane occupation recognition nexus (MORN) motifs, called MORN domain, at the N-terminal region and a C-terminal PIPK catalytic domain, except for AtPIP5K10 and AtPIP5K11 from Arabidopsis thaliana, which lack the N-terminal MORN domain.¹³ The MORN domain was first identified as plasma membrane-binding module in junctophilin¹⁴ and the involvement of the MORN domain in plasma membrane localization was proposed for A. thaliana AtPIP5K1 and AtPIP5K3.^9,15,16Another remarkable feature of eukaryotic PIPKs is dependency of the enzymatic activity on phosphatidic acid (PA).^17,18 Indeed, PA-dependent activation of PIPKs has been observed in A. thaliana and in the moss Physcomitrella patens,^6,19,20 as with animal type I PIPKs.²¹ Although much less is known about how PA activates PIPKs in plants, biochemical analyses suggested the involvement of the MORN domain in PA-dependent activation of AtPIP5K1.¹⁵Based on above findings, it was proposed that plasma membrane-localization and PA-dependent activation of plant PIPKs might be regulated by the MORN domain.^9,15,16 In contrast, we recently demonstrated the critical involvement of the C-terminal half containing the catalytic domain of plant PIPKs in both plasma membrane-localization and PA-dependent activation.²² Thus, the function of the MORN domain remains elusive in plant PIPKs.As shown earlier, the N-terminal half of P. patens PpPIPK1 containing the MORN domain enhances its catalytic activity.²² Thus, to identify the region required for the activation of PpPIPK1, we further dissected the N-terminal half into 3 regions; the N-terminal region (amino acid nos. 1–154), the MORN repeat (amino acid nos. 155–316) and the linker region (amino acid nos. 338–452), and made deletion mutants of PpPIPK1 as shown in Figure 1A. Using Pfu Turbo DNA polymerase (Stratagene, La Jolla, USA), DNA fragments corresponding to deletion mutants lacking the N-terminal and N-terminal plus the MORN repeat, designated PpPIPK1ΔN and PpPIPK1ΔN-MORN, respectively, were amplified with primer sets; one is M_PIPK1_fb (5′-GGC AAG CAC GTG TAT AAT GTC TGA AGG GCT T-3′) and XhoIPIPK1 (5′-TAA ACT CGA GTT AGC TGG GTA GGA GGA AA-3′) and the other is M_PIPK1_f7 (5′-AGA GAA CAC GTG TAT AAT GTC TGA CTT CTA CGT CGG T-3′) and XhoIPIPK1. For building an expression plasmid for a deletion mutant lacking the MORN repeat, designated PpPIPK1ΔMORN, the N-terminal region and PpPIPK1ΔN-MORN were amplified with primer sets, M_PIPK1_fb and M_PIPK1_r3 (5′-TTG TAA GTC TCG GGT GCC ATT TGA GAG CTC-3′) M_PIPK1_f6 (5′-GAG CTC TCA AAT GGC ACC CGA GAC TTA CAA-3′) and XhoIPIPK1, respectively, using Pfu Turbo DNA polymerase and resultant DNA fragments were fused by PCR with a primer set, M_PIPK1_fb and XhoIPIPK1 using the same enzyme. These PCR products were digested with Pml1 and XhoI and inserted into Pml1-XhoI digested pPICZB (Invitrogen) to construct expression plasmids, pPICZB-PpPIPK1ΔN, pPICZB-PpPIPK1ΔN-MORN and pPICZB-PpPIPK1ΔMORN. Transformation of P. pastoris X-33 cells with the above expression plasmids, colony PCR of transformants and following expression, purification and western blot analysis of His-tagged recombinant proteins were performed as described previously.⁶ The PIPK activity assay using purified His-tagged proteins was carried out as described previously²³ with the modifications.⁶Open in a separate windowFigure 1Functional dissection of the N-terminal region of PpPIPK1 identifies positive regulatory regions. (A) His-tagged recombinant PpPIPK1 proteins. A repetition of eight MORN motifs (grey boxes) and the conserved catalytic domain (black box) are indicated in wild type and mutant PpPIPK1s. The MORN repeat and junction of internal deletion are indicated by amino acid position numbers. (B) In vitro lipid kinase activity of His-tagged recombinant proteins. The activities of recombinant proteins bound to Ni-NTA agarose beads were assayed with PtdIns4P. (C) In vitro PA-dependent lipid kinase activity of His-tagged proteins. The activities of recombinant proteins bound to Ni-NTA agarose beads were assayed with PtdIns4P with 143 µM PA. Top and bottom arrowheads represent reaction products PtdIns(4,5)P₂ and lysoPtdIns(4,5)P₂, respectively.Biochemical analyses of these enzymes after expression in yeast P. pastoris X-33 cells followed by purification showed that deletion of the N-terminal region (PpPIPK1ΔN) reduced PpPIPK1 activity ca 40% compared to the full length enzyme, whereas loss of the MORN repeat (PpPIPK1ΔMORN) had no significant effect (Fig. 1B). In agreement, a mutant lacking four MORN repeats of the total eight repeats showed no difference in the activity compared the full length enzyme (data not shown). These results indicate a positive role of the N-terminal region, but not the MORN repeats, on PpPIPK1 activity. However, these findings differ from those obtained with AtPIP5K1, where the MORN domain represses enzymatic activity.¹⁵ Interestingly, PpPIPK1ΔN-MORN containing the linker and catalytic regions showed higher enzymatic activity of ca 23 % compared to the full length PpPIPK1 (Fig. 1B). The C-terminal half only containing the catalytic domain of PpPIPK1 and thus lacking the linker region showed a reduced activity.²² It is therefore proposed that the linker region carries a positive regulatory element. Although details are unknown, negligible effects of the N-terminal and MORN domains for the enzymatic activity has been indicated in AtPIP5K3 from A. thaliana.¹¹ Moreover, it is noteworthy that PA-dependent activation was not affected by any deletion as shown in Figure 1C, confirming that the N-terminal half is not involved in PA dependency of the PpPIPK1 activity.²²Our results indicated that the MORN domain is not involved in the regulation of the catalytic activity in PpPIPK1. Similarly, the function of the MORN domain found in the accumulation and replication of chloroplasts 3 (ARC3) was not resolved. ARC3 is an FtsZ homologue involved in chloroplast division²⁴ and the only protein containing the MORN repeats other than PIPKs in A. thaliana. It was shown that the ARC3 MORN domain did not interact with any stromal plastid division components.²⁵ Moreover, there are reports representing functions of the MORN domain other than plasma membrane binding. Human amyotrophic lateral sclerosis 2 (ALS2), a guanine nucleotide exchange factor (GEF) specific to the small GTPase Rab5, contains the MORN domain at the central region that is essential for the GEF activity but not for interaction with Rab5.²⁶ In contrast, specific interaction of the MORN domain with Rab-E GTPases and resultant enzymatic activation was recently demonstrated for AtPIP5K2.¹² It is interesting that these results are inconsistent with each other in terms of interaction of the MORN domain with small GTPases.Taken together, with no function of the MORN domain in plasma membrane localization of PpPIPK1 and AtPIP5K1,²² the function of the MORN domain is still unknown, despite its high conservation plants PIPKs. Alternatively, based on the findings of ARC3, ALS2 and AtPIP5K2,^12,25,26 the function of the MORN domain possibly varies among PIPK isoforms and may thus have multifunctional roles. Therefore, it is necessary to identify interaction partners for the MORN domain of each plant PIPKs and to analyze phenotypes of transgenic plants carrying MORN domain-lacking PIPKs during developmental process and environmental stress responses.     

Reactive oxygen species as transducers of sphinganine-mediated cell death pathway

Long chain bases or sphingoid bases are building blocks of complex sphingolipids that display a signaling role in programmed cell death in plants. So far, the type of programmed cell death in which these signaling lipids have been demonstrated to participate is the cell death that occurs in plant immunity, known as the hypersensitive response. The few links that have been described in this pathway are: MPK6 activation, increased calcium concentrations and reactive oxygen species (ROS) generation. The latter constitute one of the more elusive loops because of the chemical nature of ROS, the multiple possible cell sites where they can be formed and the ways in which they influence cell structure and function.Key words: hydrogen peroxide, long chain bases, programmed cell death, reactive oxygen species, sphinganine, sphingoid bases, superoxideA new transduction pathway that leads to programmed cell death (PCD) in plants has started to be unveiled.^1,2 Sphingoid bases or long chain bases (LCBs) are the distinctive elements in this PCD route that naturally operates in the entrance site of a pathogen as a way to contend its spread in the plant tissues.^2,3 This defense strategy has been known as the hypersensitive response (HR).^4,5As a lately discovered PCD signaling circuit, three connected transducers have been clearly identified in Arabidopsis: the LCB sphinganine (also named dihydrosphingosine or d18:0); MPK6, a mitogen activated kinase and superoxide and hydrogen peroxide as reactive oxygen species (ROS).^1,2 In addition, calcium transients have been recently allocated downstream of exogenously added sphinganine in tobacco cells.⁶Contrary to the signaling lipids derived from complex glycerolipid degradation, sphinganine, a metabolic precursor of complex sphingolipids, is raised by de novo synthesis in the endoplasmic reticulum to mediate PCD.^1,2 Our recent work demonstrated that only MPK6 and not MPK3 (commonly functionally redundant kinases) acts in this pathway and is positioned downstream of sphinganine elevation.² Although ROS have been identified downstream of LCBs in the route towards PCD,¹ the molecular system responsible for this ROS generation, their cellular site of formation and their precise role in the pathway have not been unequivocally identified. ROS are produced in practically all cell compartments as a result of energy transfer reactions, leaks from the electron transport chains, and oxidase and peroxidase catalysis.⁷Similar to what is observed in pathogen defense,³ increases in endogenous LCBs may be elicited by addition of fumonisin B1 (FB1) as well; FB1 is a mycotoxin that inhibits ceramide synthase. This inhibition results in an accumulation of its substrate, sphinganine and its modified forms, leading to the activation of PCD.^1,2,8 The application of FB1 is a commonly used approach for the study of PCD elicitation in Arabidopsis.^1,2,9–11An early production of ROS has been linked to an increase of LCBs. For example, an H₂O₂ burst is found in tobacco cells after 2–20 min of sphinganine supplementation,¹² and superoxide radical augmented in the medium 60 min after FB1 or sphinganine addition to Arabidopsis protoplasts (Fig. 1A). In consonance with this timing, both superoxide and H₂O₂ were detected in Arabidopsis leaves after 3–6 h exposure to FB1 or LCBs.¹ However, the source of ROS generation associated with sphinganine elevation seems to not be the same in both species: in tobacco cells, ROS formation is apparently dependent on a NADPH oxidase activity, a ROS source consistently implicated in the HR,^13,14 while in Arabidopsis, superoxide formation was unaffected by diphenyliodonium (DPI), a NADPH oxidase inhibitor (Fig. 1A). It is possible that the latter oxidative burst is due to an apoplastic peroxidase,¹⁵ or to intracellular ROS that diffuse outwards.^16,17 These results also suggest that both tobacco and Arabidopsis cells could produce ROS from different sources.Open in a separate windowFigure 1ROS are produced at early and long times in the FB1-induced PCD in Arabidopsis thaliana (Col-0). (A) Superoxide formation by Arabidopsis protoplasts is NADPH oxidase-independent and occurs 60 min after FB1 or sphinganine (d18:0) exposure. Protoplasts were obtained from a cell culture treated with cell wall lytic enzymes. Protoplasts were incubated with 10 µM FB1 or 10 µM sphinganine for 1 h. Then, cells were vacuum-filtered and the filtrate was used to determine XTT [2,3-bis-(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide, disodium salt] reduction as described in references ²⁸ and ²⁹. DPI was used at 50 µM. (B) H₂O₂ formation in Arabidopsis wt and lcb2a-1 mutant in the presence and absence of FB1. Arabidopsis seedlings were exposed to 10 µM FB1 and after 48 h seedlings were treated with DA B (3,3-diaminobencidine) to detect H₂O₂ according to Thordal-Christensen et al.³⁰It has been suggested that the H₂O₂ burst associated with the sphinganine signaling pathway leads to the expression of defense-related genes but not to the PCD itself in tobacco cells.¹² It is possible that ROS are involved in the same way in Arabidopsis, since defense gene expression is also induced by FB1 in Arabidopsis.⁹ In this case, it will be important to define how the early ROS that are DPI-insensitive could contribute to the PCD manifestation mediated by sphinganine.The generation of ROS (4–60 min) found in Arabidopsis was associated to three conditions: the addition of sphinganine (Fig. 1A), FB1 (Fig. 1A) or pathogen elicitors.¹⁵ This is consistent with the MPK6 activation time, which is downstream of sphinganine elevation and occurs as early as 15 min of FB1 or sphinganine exposure.² All of them are events that appear as initial steps in the relay pathway that produces PCD.In order to explore a possible participation of ROS at more advanced times of PCD progression, we detected in situ H₂O₂ formation in Arabidopsis seedlings previously exposed to FB1 for 48 h. As shown in Figure 1B, formation of the brown-reddish precipitate corresponding to the reaction of H₂O₂ with 3,3′-diaminobenzidine (DAB) was only visible in the FB1-exposed wild type plants, as compared to the non-treated plants. However, when lcb2a-1 mutant seedlings were used, FB1 exposure had a subtle effect in ROS formation. This mutant has a T-DNA insertion in the gene encoding subunit LCB2a from serine palmitoyltransferase (SPT), which catalyzes the first step in sphingolipid synthesis¹⁸ and the mutant has a FB1-resistant phenotype.² These results indicate that mutations in the LCB1¹ and LCB2a² genes (coding for the subunits of the heterodimeric SPT) that lead to a non-PCD phenotype upon the FB1 treatment, are unable to produce H₂O₂. In addition, they suggest that high levels of hydrogen peroxide are produced at advanced times in the PCD mediated by LCBs in Arabidopsis.Exposure of Arabidopsis to an avirulent strain of Pseudomonas syringae produces an endogenous elevation of LCBs as a way to implement defense responses that include HR-PCD.³ In this condition, we clearly detected H₂O₂ formation inside chloroplasts (Fig. 2A). When ultrastructure of the seedlings tissues exposed to FB1 for 72 h was analyzed, integrity of the chloroplast membrane system was severely affected in Arabidopsis wild-type seedlings exposed to FB1.² Therefore, we suggest that ROS generation-LCB induced in the chloroplast could be responsible of the observed membrane alteration, as noted by Liu et al. who found impairment in chloroplast function as a result of H₂O₂ formation in this organelle from tobacco plants. Interestingly, these plants overexpressed a MAP kinase kinase that activated the kinase SIPK, which is the ortholog of the MPK6 from Arabidopsis, a transducer in the PCD instrumented by LCBs.²Open in a separate windowFigure 2Conditions of LCBs elevation produce H₂O₂ formation in the chloroplast and perturbation in the membrane morphology of mitochondria. (A) Exposure of Arabidopsis leaves to the avirulent strain Pseudomonas syringae pv. tomato DC3000 (avrRPM1) (or Pst avrRPM1) induces H₂O₂ formation in the chloroplast. Arabidopsis leaves were infiltrated with 1 × 10⁸ UFC/ml Pst avrRPM1 and after 18 h, samples were treated to visualize H₂O₂ formation with the DAB reaction. Controls were infiltrated with 10 mM MgCl₂ and then processed for DAB staining. Then, samples were analyzed in an optical photomicroscope Olympus Provis Model AX70. (B) Effect of FB1 on mitochondria ultrastructure. Wild type Arabidopsis seedlings were treated with FB1 for 72 h and tissues were processed and analyzed according to Saucedo et al.² Ch, chloroplast; M, mitochondria; PM, plasma membrane. Arrows show mitochondrial cisternae. Bars show the correspondent magnification.In addition, we have detected alterations in mitochondria ultrastructure as a result of 72 h of FB1 exposure (Fig. 2B). These alterations mainly consist in the reduced number of cristae, the membrane site of residence of the electron transport complexes. In this sense, it has been shown that factors that induce PCD such as the victorin toxin, methyl jasmonate and H₂O₂ produce alterations in mitochondrial morphology.^20–22 In fact, some of these studies propose that ROS are formed in the mitochondria and then diffuse to the chloroplasts.^22–24It is reasonable to envisage that damage of the membrane integrity of these two organelles reflects the effects of vast amounts of ROS produced by the electron transport chains.^25,26 Recent evidence supports the destruction of the photosynthetic apparatus associated to the generation of ROS in the HR.²⁶ At this time of PCD progression, ROS could be contributing to shut down the energy machinery in the cell, which ultimately would become the point of no-return of PCD²⁷ as part of the execution program of the cell death mediated by LCBs.In conclusion, we propose that ROS can display two different functional roles in the PCD process driven by LCBs. These roles depend on the time of ROS expression, the cellular site where they are generated, the enzymes that produce them, and the magnitude in which they are formed.