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.     

Do Plant Cells Secrete Exosomes Derived from Multivesicular Bodies?

Multivesicular bodies (MVBs) are spherical endosomal organelles containing small vesicles formed by inward budding of the limiting membrane into the endosomal lumen. In mammalian red cells and cells of immune system, MVBs fuse with the plasma membrane in an exocytic manner, leading to release their contents including internal vesicles into the extracellular space. These released vesicles are termed exosomes. Transmission electron microscopy studies have shown that paramural vesicles situated between the plasma membrane and the cell wall occur in various cell wall-associated processes and are similar to exosomes both in location and in morphology. Our recent studies have revealed that MVBs and paramural vesicles proliferate when cell wall appositions are rapidly deposited beneath fungal penetration attempts or during plugging of plasmodesmata between hypersensitive cells and their intact neighboring cells. This indicates a potential secretion of exosome-like vesicles into the extracellular space by fusion of MVBs with the plasma membrane. This MVB-mediated secretion pathway was proposed on the basis of pioneer studies of MVBs and paramural vesicles in plants some forty years ago. Here, we recall the attention to the occurrence of MVB-mediated secretion of exosomes in plants.Key Words: cell wall, endocytosis, endosome, exocytosis, exosome, multivesicular body, paramural bodyMultivesicular bodies (MVBs) are spherical endosomal organelles containing a number of small vesicles formed by inward budding of the limiting membrane into the endosomal lumen.¹ MVBs contain endocytosed cargoes and deliver them into lysosomal/vacuolar compartments for degradation. They also incorporate newly synthesized proteins destined for lysosomal/vacuolar compartments.² In mammalian cells of hematopoietic origin, endosomal MVBs function in removal of endocytosed surface proteins in an exocytic manner. They are redirected to the plasma membrane, where they release their contents including internal vesicles into the extracellular space by membrane fusion. The released vesicles are termed exosomes.³ During reticulocyte maturation to erythrocyte, a group of surface proteins, such as the transferrin receptor, become obsolete and are discarded via MVB-mediated secretion.³ Time-course transmission electron microscopy (TEM) first revealed that colloidal gold-transferrin was internalized into MVBs via receptor-mediated endocytosis and then transferrin together with its receptor were delivered into the extracellular space via the fusion of MVBs with the plasma membrane of reticulocytes.⁴ Some other cell types of hematopoietic origin, such as activated platelets, cytotoxic T cells and antigen-presenting cells, also secrete exosomes. Exosomes thus may play a role in various physiological processes other than discarding obsolete proteins.³Our recent TEM studies provided ultrastructural evidence on the enhanced vesicle trafficking in barley leaf cells attacked by the biotrophic powdery mildew fungus. Multivesicular compartments including MVBs, intravacuolar MVBs, and paramural bodies turned out to proliferate in intact host cells during formation of cell wall appositions (papilla response), in the hypersensitive response, and during accommodation of haustoria.^5,6 MVBs proliferated in the cytoplasm of haustorium-containing epidermal cells during compatible interactions and near sites of cell wall-associated oxidative microburst either during the papilla response or during the hypersensitive response. Because MVBs in plant cells have been demonstrated to be endosomal compartments,^7–9 they may participate in internalization of nutrients from the apoplast of intact haustorium-containing epidermal cells and sequestration of damaged membranes and deleterious materials originating from the oxidative microburst.^5,6 The presence of intravacuolar MVBs with double limiting membranes (Fig. 1A) indicates an engulfment of MVBs by the tonoplast and a vacuole-mediated autophagy of MVBs.^5,6 MVBs, as prevacuolar compartments in plant cells,⁹ thus probably deliver their contents into the central vacuole via both the fusion with the tonoplast and the engulfment by the tonoplast (Fig. 2A and B). On the other hand, paramural bodies, in which small vesicles are situated between the cell wall and the plasma membrane, were associated with cell wall appositions deposited beneath fungal penetration attempts (Fig. 1B) or around hypersensitive cells including sites of plugged plasmodesmata (Fig. 1C and D).^5,6 Because paramural vesicles are similar to exosomes both in location and in morphology, we speculated that MVBs fuse with the plasma membrane in an exocytic manner to form paramural bodies.^5,6 Endocytosed cell surface materials in endosomal MVBs may be reused and delivered together with newly synthesized materials in Golgi apparatus-derived vesicles to cell wall appositions, which are deposited rapidly to prevent fungal penetration (Fig. 2A) or to contain hypersensitive cell death (Fig. 2B). MVBs thus may be driven along two distinct pathways to deliver their contents into either central vacuole or extracellular space.Open in a separate windowFigure 1Multivesicular compartments in intact cells in barley leaves attacked by the barley powdery mildew fungus. (A) An intravacuolar multivesicular body (MVB) with double limiting membranes in an intact epidermal cell (EC) adjacent to a hypersensitive epidermal cell (EC*). The arrows point to the outer limiting membrane, which is seemingly derived from the tonoplast. Note that neighboring intravacuolar vesicles (in between two arrowheads) may result from degradation of double limiting membranes of intravacuolar MVBs or may be delivered into the vacuole by MVB-fusion with the tonoplast. (B) Paramural vesicles (arrowheads) in a paramural body associated with cell wall appositions (asterisk) deposited by an intact epidermal cell. (C) A multivesicular body (MVB) in contact with a paramural body (PMB) (a nonmedian section) associated with cell wall appositions (asterisk) deposited by an intact mesophyll cell adjacent to a hypersensitive mesophyll cell. Note that cell wall appositions deposit beside an intercellular space (IS). The arrows point to the tonoplast. (D) A paramural body (PMB) associated with cell wall appositions (asterisks) blocking plasmodesmata (in between two arrowheads) at the side of an intact mesophyll cell (MC) underlying a hypersensitive epidermal cell (EC*). The arrows point to the tonoplast. CV, central vacuole; CW, cell wall; MB, microbody. Bars, 1µm.Open in a separate windowFigure 2Hypothetical diagram of delivery of endocytosed cell surface materials via MVBs into the central vacuole or the extracellular space where intact barley cells deposit cell wall appositions. (A) Deposition of cell wall appositions (asterisk) beneath powdery mildew penetration attempts. AGT, appressorial germ tube; PP, penetration peg. (B) Deposition of cell wall appositions (asterisks) against constricted plasmodesmata (PD) between a hypersensitive epidermal cell (EC) penetrated by the powdery mildew fungus and an underlying mesophyll cell (MC). H, haustorium. Arrows and numbers show pathways of vesicle trafficking. 1, Secretion of Golgi-derived vesicles containing newly synthesized materials; G, Golgi body; TGN, trans-Golgi network; 2, Endocytosis of cell surface materials from coated pits (coated open circles) via coated vesicles (coated circles) to multivesicular bodies (MVB); 3, Delivery of endocytosed materials for degradation inside the central vacuole (CV) via membrane fusion between MVBs and the tonoplast (T); small broken circles, vesicles in degradation; 4, Delivery of endocytosed materials for degradation inside the central vacuole via engulfment of MVBs by the tonoplast; large broken circles; MVB limiting membranes in degradation; 5, delivery of endocytosed materials into the extracellular space for deposition of cell wall appositions (asterisks) via membrane fusion between MVBs and the plasma membrane (PM). CW, cell wall; PMB, paramural body. PD0, 1, 2, 3 and 4 represent stages of plugging plasmodesmata. PD0, open plasmodesmata between two intact mesophyll cells (MC) subjacent to the hypersensitive epidermal cell (EC); PD1, constriction of plasmodesmata by callose (grey dots) deposition at plasmodesmal neck region; PD2, constricted plasmodesmata associated with plasmodesma-targeted secretion; PD3, further blocking of plasmodesmata by deposition of cell wall appositions; PD4, completely blocked plasmodesmata.Earlier than the discovery in animal cell systems,⁴ it was proposed in two independent papers in 1967 that the fusion of MVBs with the plasma membrane might result in the release of small vesicles into the extracellular space in fungi and in higher plants.^10,11 Several lines of evidence support the occurrence of MVB-mediated secretion of exosome-like vesicles in plants. First, vesicles of the same morphology as MVB internal vesicles have been observed in extracellular spaces or paramural spaces in various types of plant cells in various plant species by TEM.¹² An early study on endocytosis by soybean protoplasts also showed small extracellular vesicles attaching on the plasma membrane.⁸ Second, cooccurrence of MVBs and paramural vesicles has been observed in processes of cell proliferation, cell differentiation, and cell response to abiotic and biotic stress. Examples are cell plate formation,^13,14 secondary wall thickening,^15,16 cold hardness,^17,18 and deposition of cell wall appositions upon pathogen attack.^5,6,19–21 Third, identical molecular components, such as arabinogalactan proteins^22,23 and peroxidases,⁶ have been immunolocalized in both MVBs and paramural bodies. Despite these pieces of evidence, a conclusive demonstration of MVB-mediated secretion of exosomes in plants requires further exploration.The presently available experimental systems, approaches, and membrane markers may allow future demonstration of MVB-mediated secretion of exosomes in plants. Recent in vivo real-time observation and colocalization of cell surface and endosomal markers have already revealed that endosomes filled with endocytosed preexisting cell wall and plasma membrane materials are rapidly delivered to cytokinetic spaces to form cell plates in dividing tobacco, Arabidopsis, and maize cells.²⁴ Because TEM observed paramural bodies attaching to cell plates¹³ and MVBs in the vicinity of cell plates during all stages of cell plate formation,^14,25,26 MVBs and paramural bodies may participate in delivery of endocytosed building blocks to cell plates. Jiang''s and Robinson''s labs together developed a transgenic tobacco BY-2 cell line stably expressing a YFP-labeled vacuolar sorting receptor protein and antibodies against the vacuolar sorting receptor protein localized to the limiting membrane of MVBs.⁹ These tools together with live cell imaging and immunoelectron microscopy may allow visualization of MVB-fusion to the new plasma membrane, of vacuolar sorting receptors in both the limiting membrane of MVBs and the new plasma membrane, and of identical cell plate components in both internal vesicles of MVBs and paramural vesicles.In spite of obvious differences in plant and animal cytokinesis, the generation of cell plates by cell-plate-directed fusion of endosomes resembles the plugging of midbody canals by midbody-directed endosomes to separate daughter cells at the terminal phase of animal cytokinesis.²⁷ Likely, functional similarities of the fusion between endosomal MVBs and the plasma membrane to eliminate unwanted cell contents may also exist in maturation of mammalian red blood cells and plant sieve elements in the sense that the fusion of MVBs with the plasma membrane may occur during maturation of the latter.²⁸ On the other hand, although plant cells may secrete MVB-derived exosomes in defense response upon pathogen attack,^5,6 plant cell walls rule out the direct intercellular communication during the immune response mediated by exosomes in the circulation of mammals.³ In contrast, plasmodesma-directed secretion of exosomes would block the cell-to-cell communication between hypersensitive cells and their neighboring cells during hypersensitive response.⁵ Further exploration will lead us to a better understanding of similarities and differences of exosome secretion between plants and animals.     

The Arabidopsis hit1-1 mutant has a plasma membrane profile distinct from that of wild-type plants at optimal growing temperature

High temperatures alter the physical properties of the plasma membrane and cause loss-of-function in the embedded proteins. Effective membrane and protein recycling through intracellular vesicular traffic is vital to maintain the structural and functional integrity of the plasma membrane under heat stress. However, in this regard, little experimental data is available. Our characterization of the Arabidopsis hit1-1 mutant, linking a subunit of a vesicle tethering complex to plasma membrane thermostability, provided valuable information to this end. We further dissected the effect of the hit1-1 mutation on plasma membrane properties and found that even at optimal growth temperature (23°C), the hit1-1 mutant exhibited a plasma membrane protein profile distinct from that of wild-type plants. This result implies that the hit1-1 mutation essentially alters vesicle trafficking and results in changes in the plasma membrane components under non-stress conditions. Such changes do not affect normal plant growth and development, but is significant for plant survival under heat stress.Key words: GARP complex, heat stress, heat intolerant, HIT1, membrane trafficking, vesicle tethering factor, Vps53The plasma membrane is indispensable to all living cells. It serves as a barrier separating the interior and exterior of the cell, and consists of a variety of proteins that accomplish vital biological functions. High temperature can fluidize the plasma membrane and damage the functions of its embedded proteins. Severe heat stress may even disrupt the integrity of the plasma membrane, causing escape of essential cytoplasmic constituents and leading to cell death.^1,2 Although it is predictable that effective vesicle trafficking machinery, which is involved in the removal of proteins and lipid molecules from and the delivery of freshly synthesized components to the plasma membrane, is important for plasma membrane rejuvenation and should participate in plant thermotolerance,³ little empirical data has come to light to elucidate the protective mechanism by which vesicle trafficking improves plant tolerance to heat stress.The Arabidopsis hit1-1 mutant was originally isolated by its heat-intolerant phenotype.⁴ Map-based cloning led to the identification of HIT1 as a homolog of yeast Vps53p,⁵ which is a subunit of the Golgi-associated retrograde protein (GARP) tethering complex that is involved in vesicle trafficking from the endosome to the trans-Golgi network (TGN).⁶ Taking advantage of the available hit mutants, Wang et al. investigated the causality between HIT1 and plasma membrane thermostability,^7,8 and demonstrated that, while the anti-oxidative capability of hit1-1 plants was equal to that of wild-type plants, significantly more electrolyte leakage from hit1-1 leaves was detected after long term heat exposure (37°C for 24 h). Furthermore, hit1-1 was not sensitive to sudden heat shock (44°C for 30 min).⁷ These findings demonstrated that there is indeed a vesicle trafficking mediator of plasma membrane thermal adaptation, and this adaptation is probably more involved in remodeling than in repair. Such remodeling enables the membrane to withstand elevated temperatures, circumvent heat-induced damage, and thus is specifically significant for tolerance of long-term heat stress.⁷Since the hit1-1 allele is a point mutation leading to a Ser-to-Tyr amino acid substitution,⁵ one may suspect that the temperature-sensitive phenotype of hit1-1 plants is produced by the thermolabile properties of the hit1-1 protein. Nevertheless, because hit1-1 plants are more sensitive than wild-type to osmotic and saline stress inhibition during seedling germination and development,^4,5,7 it is unlikely that the heat-intolerant phenotype of hit1-1 is derived from a simple alternation in the thermal stability of a gene product. To answer this question and to provide further insight into the roles of HIT1 in plasma membrane acclimation to heat stress, electrophoretic patterns of plasma membrane proteins were analyzed. Even at the optimal growth temperature (23°C), wild-type, and hit1-1 plants have different plasma membrane protein profiles (Fig. 1). This result suggests that, regardless of growing temperature, the hit1-1 mutation essentially alters regular recycling of plasma membrane components, and this alteration does not affect plant growth and development under non-stress conditions, but is unfavorable or even lethal for plants growing under certain stress conditions.Open in a separate windowFigure 1One-dimensional SDS gel electrophoretic banding patterns of plasma membrane proteins from 4-wk-old, 23°C-grown wild-type (WT) and hit1-1 plants. Plasma membrane proteins were prepared as previously described in reference ¹³, with minor modification, and then separated on a 12.5% SDS-PAGE gel, and the protein bands were detected by silver staining. Equal amounts of proteins were loaded in each lane. Arrows indicate some protein bands showing noticeable differences in staining intensity, reflecting that these proteins were present in different abundances in the wild-type and hit1-1 plasma membranes. Molecular mass markers are shown on the left.Modification of lipid saturation levels is a well known mechanism for membrane acclimation to heat,¹ and is more significant for plant tolerance of longer-term heat stress than heat shock.⁹ The hit1-1 heat-sensitive phenotype correlates with this notion. Meanwhile, mutants that have a defect in a gene that encodes digalactosyldiacylglycerol (DGDG) synthase become thermosensitive. This thermosensitivity is associated with an inability to increase the ratio of DGDG to monogalactosyldiacylglycerol (MGDG) upon exposure to high temperature.¹⁰ DGDG is normally a plastid-specific lipid but has been found in the plasma membrane upon phosphate deprivation.^11,12 These data suggest that modification of membrane lipids for high temperature adaptation may not be restricted to changes in fatty acid saturation level. How vesicle trafficking machinery participates in this global reorganization of membrane lipids and to what extent it affects the heat tolerance are largely unclear, and the hit1-1 mutant holds great potential to provide novel insight to these questions.     

Calcium/calmodulin-regulated receptor-like kinase CRLK1 interacts with MEKK1 in plants

Recently we reported that CRLK1, a novel calcium/calmodulin-regulated receptor-like kinase plays an important role in regulating plant cold tolerance. Calcium/calmodulin binds to CRLK1 and upregulates its activity. Gene knockout and complementation studies revealed that CRLK1 is a positive regulator of plant response to chilling and freezing temperatures. Here we show that MEKK1, a member of MAP kinase kinase kinase family, interacts with CRLK1 both in vitro and in planta. The cold triggered MAP kinase activation in wild-type plants was abolished in crlk1 knockout mutants. Similarly, the cold induced expression levels of genes involved in MAP kinase signaling are also altered in crlk1 mutants. These results suggest that calcium/calmodulinregulated CRLK1 modulates cold acclimation through MAP kinase cascade in plants.Key words: calcium, calmodulin, cold stress, MAPK, Arabidopsis, protein phosphorylationCalcium, a universal second messenger in eukaryotic cells, mediates changes in external and internal signals leading to the physiological responses.^1–4 Calcium/calmodulin (Ca²⁺/CaM)-dependent protein kinases (CaMKs) are very important players in calcium/calmodulin mediated signaling in mammalian cells.⁵ In plants, Ca²⁺/CaM-dependent protein phosphorylation was observed more than 25 years ago.⁶ Several calmodulin-regulated protein kinases have been identified and characterized.^7,8 For example, plants have a unique chimeric Ca²⁺/CaM-dependent protein kinase (CCaMK), which exhibits Ca²⁺-dependent autophosphorylation and Ca²⁺/CaM-dependent substrate phosphorylation.⁹ CCaMK is required for bacterial and fungal symbioses in plants.^10–12 Recently, we characterized a novel plant-specific calcium/CaM-regulated receptor-like kinase, CRLK1.¹³ Ca²⁺/CaM binds to CRLK1 and stimulates its kinase activity. Functional studies with CRLK1 indicate that CRLK1 acts as a positive regulator in plant response to chilling and freezing temperatures. To further define the CRLK1-mediated signal pathway, we isolated CRLK1 interacting proteins by co-immunoprecipitation using an anti-CRLK1 antibody. Since cold increases the amount of CRLK1 protein, wildtype plants (WT) were treated at 4°C for 1 hr before co-immunoprecipitation. The resulting CRLK1 immunocomplex was separated by SDS-PAGE. We observed several bands of different sizes only in the wild-type but not in the crlk1 knockout mutant plants (Fig. 1A). Furthermore, the intensity of these bands increased upon cold treatment, suggesting that they are the putative partners or associated proteins of the CRLK1 immunocomplex.Open in a separate windowFigure 1CRLK1 Interacts with MEKK1. (A) One-dimension SDS-PAGE of anti-CRLK1 immunocomplexes from 3-week-old WT or crlk1 plants with or without cold treatment. One mg of total protein was used for immunoprecipitation. (B) A list of putative CRLK1-interacting proteins determined by MALDI-TOF-MS analysis. (C) CRLK1 interacts with MEKK1 as shown by GST pull-down assay. (D) BiFC analysis show that CR LK associates with MEKK1 in vivo. Upper row shows that CRLK and MEKK1 associate both on cell membrane and in endosomes. The middle and last rows are controls. Bar = 10 µm.To determine the identities of these proteins, mass spectrometric analysis was performed with the total immunocomplex.¹⁴ In addition to CRLK1, there were 12 other proteins which matched the Arabidopsis database. Several of them appeared in the pull-down complex from WT, but not from crlk1 mutants. These putative interacting proteins included MEKK1, another unknown protein kinase, a type 2C phosphatase and CaM (Fig. 1B). MEKK1 is one of the 60 putative MAPKKKs in the Arabidopsis genome, and sits on the top of mitogen-activated protein kinase (MAPK) cascade. The MAPK signaling consists of a cascade of three consecutively acting protein kinases, a MAP kinase kinase kinase (MAPKKK), a MAP kinase kinase (MAPKK) and a MAP kinase (MAPK). Plants possess multiple MAPKKKs, MAPKKs and MAPKs, which respond to different upstream signals and activate distinct downstream pathways.^15–17 The specific MAPK module responding to lower temperature has been determined in Arabidopsis.^18,19 MEKK1, a member of MAPKKKs, specifically interacts and phosphorylates MKK2 and regulates COR genes expression in response to cold stress.¹⁹ MEKK1 has been shown to play a role in mediating reactive oxygen species homeostasis.^20,21 Therefore we selected MEKK1 from the putative CRLK1 partners for further studies.     

AtAGP18, a lysine-rich arabinogalactan protein in Arabidopsis thaliana,functions in plant growth and development as a putative co-receptor for signal transduction

Arabinogalactan-proteins (AGPs) are a class of hyperglycosylated, hydroxyproline-rich glycoproteins that are widely distributed in the plant kingdom. AtAGP17, 18 and 19 are homologous genes encoding three classical lysine-rich AGPs in Arabidopsis. We observed subcellular localization of AtAGP18 at the plasma membrane by expressing a translational fusion gene construction of AtAGP18 attached to a green fluorescent protein (GFP) tag in Arabidopsis plants. We also overexpressed AtAGP18 without the GFP tag in Arabidopsis plants, and the resulting transgenic plants had a short, bushy phenotype. Here we discuss putative roles of AtAGP18 as a glycosylphosphatidylinositol (GPI)-anchored protein involved in a signal transduction pathway regulating plant growth and development.Key words: Arabidopsis thaliana, arabinogalactan-proteins, co-receptor, glycosylphosphatidylinositol, lipid rafts, overexpressionArabinogalactan-proteins (AGPs) are plant cell surface glycoproteins or proteoglycans which are thought to play important roles in various aspects of plant growth and development, such as somatic embryogenesis, cell proliferation and elongation, pattern formation and hormone signaling.¹ The lysine-rich classical AGP subfamily in Arabidopsis contains three members: AtAGP17, 18 and 19. The subcellular localization of AtAGP17 and AtAGP18 was previously studied in our laboratory by expressing GFP-AtAGP17/18 fusion proteins in tobacco cell cultures.^2,3 In a recent report, we used Arabidopsis plants to overexpress GFP-AtAGP17/18/19 fusion proteins to observe subcellular localization of the lysine-rich AGPs in planta, in contrast to our previous plant cell culture work.⁴ Moreover, the lysine-rich AGPs alone (i.e., AtAGP17/18/19 without the GFP tag) were overexpressed in Arabidopsis plants, and only AtAGP18 overexpressors had a distinctive phenotype. This phenotype included shorter stems, more branches and less seeds, indicating a role for AtAGP18 in plant growth and development.⁴ In this addendum, we further discuss the putative biological role of AtAGP18 on a molecular level and its possible mode of action in cellular signaling.Classical AGPs are frequently predicted to have a glycosylphosphatidylinositol (GPI) anchor, which would allow for the localization of such AGPs to the outer surface of the plasma membrane. Biochemical analyses were carried out to support this hypothesis in tobacco, pear,⁵ rose⁶ and Arabidopsis.⁷ The lysine-rich classical AGPs, AtAGP17 and 18, were predicted to have a GPI anchor.⁸ To test this idea, tobacco cell cultures expressing GFP-AtAGP17/18 fusion proteins were plasmolyzed and GFP fluorescence was observed on the plasma membrane.^2,3 To corroborate this finding in planta, GFP-AtAGP17/18 were expressed in Arabidopsis plants and leaf trichome cells were plasmolyzed. Enhanced GFP fluorescence was observed at the plasma membrane of these transgenic trichome cells, indicating the presence of GFP-AtAGP17/18 at the plasma membrane.⁴ The localization of these lysine-rich classical AGPs at the plasma membrane suggests possible biological roles in sensing extracellular signals. They are likely associated with lipid rafts involved in cell signaling for the following reasons. In plants as well as animals, there are sterol-enriched, detergent-resistant plasma membrane microdomains called lipid rafts. Lipid rafts are known to be involved in signal transduction and are enriched in transmembrane receptors and GPI-anchored proteins, including AGPs.^9–11 The accumulation of these proteins in such microdomains may allow for interactions between these proteins in sensing extracellular signals which lead to various intracellular events. Interestingly, a recent study shows that lipid rafts from hybrid aspen cells contain callose synthase and cellulose synthase, and these enzymes are active since in vitro polysaccharide synthesis by the isolated detergent-resistant membranes was observed. These results demonstrate that lipid rafts are involved in cell wall polysaccharide biosynthesis.¹² In addition, an Arabidopsis pnt mutant study shows GPI-anchored proteins are required in cell wall synthesis and morphogenesis.¹³ These observations, coupled with previous observations that cellulose synthases as well as AGPs interact with microtubules, suggest that AGPs in lipid rafts may have a role in signal events, including those regulating cellulose and/or callose biosynthesis or deposition.^14,15To examine the role of LeAGP-1, a lysine-rich AGP in tomato, transgenic tomato plants were produced which expressed GFP-LeAGP-1 under the control of the cauliflower mosaic virus 35S promoter.¹⁶ The tomato LeAGP-1 overexpressors and Arabidopsis AtAGP18 overexpressors both have a bushy phenotype similar to transgenic tobacco plants overproducing cytokinins.^4,16,17 Cytokinins are an important class of plant hormones involved in many plant growth and development processes, such as cell growth and division, differentiation and other physiological processes.¹⁸ Therefore, Sun et al. proposed that LeAGP-1 might function in concert with the cytokinin signal transduction pathway.¹⁶ Since the overexpression phenotypes of AtAGP18 are similar to those of LeAGP-1, AtAGP18 is also likely associated with the cytokinin signal transduction pathway. The prevailing model for cytokinin signaling in Arabidopsis is similar to the two-component system in bacteria and yeast. In this model, the cytokinin receptor contains an extracellular domain, a kinase domain and a receiver domain. When the cytokinin receptor senses cytokinin signals, it auto-phosphorylates at a His residue in the kinase domain. The phosphoryl group is then transferred to an Asp residue in the receiver domain. Subsequently, the phosphoryl group is transferred to a His residue in the histidine phosphotransfer protein (Hpt) and the Hpt translocates to the nucleus and transfers the phosphoryl group to an Asp residue in a downstream response regulator to activate it.¹⁹ This model is consistent with our hypothesis since the cytokinin receptor in this model is a receptor kinase located in the plasma membrane with an extra-cellular domain that can potentially interact with AtAGP18. AtAGP18 may function as a co-receptor that first binds to cytokinins, then either directly interacts with cytokinin receptors or brings the cytokinins to cytokinin receptors in the plasma membrane. The first scenario is analogous to the interaction of contactin and contactin-associated protein (Caspr) in neurons. In this model, contactin is a GPI-anchored protein on the cell surface that binds to signal molecules and interacts with the transmembrane receptor Caspr to transmit signals to the cell interior.²⁰ The second scenario is analogous to fibroblast growth factor (FGF) signal activation in which heparan sulfate proteoglycans bind to FGF molecules and bring them to the FGF receptor.²¹Based on all the above observations and findings, a hypothetical model for AtAGP18 function is proposed in Figure 1. The model shows AtAGP18 located on the outer surface of the plasma membrane in lipid rafts where it could act as a co-receptor to sense extracellular signals (such as cytokinin) and interact with transmembrane proteins, possibly receptor kinases or ion channels, in the lipid rafts to initiate signaling by triggering various intracellular events. Interestingly, receptor tyrosine kinases and ion channels are known to be present in lipid rafts.^9,22 Moreover, AGPs are likely associated with ion channels since addition of the AGP-binding reagent Yariv phenylglycoside resulted in elevated cytoplasmic calcium concentrations in tobacco cells and lily pollen tubes.^15,23,24 Clearly, additional work will be required to verify such a model, and to better understand how AtAGP18 might sense extracellular signals and interact with the transmembrane proteins in the lipid rafts.Open in a separate windowFigure 1Model for atAGP18 functioning in cellular signaling to control plant growth and development. In this model, lipid rafts are enriched in glycosphingolipids, sterols, transmembrane proteins (such as receptors, receptor kinases and ion channel proteins) and GPI-anchored proteins including AtAGP18. (a) AtAGP18 acts as a co-receptor by binding to signaling molecules and directly interacting with transmembrane proteins in the lipid rafts. (B) AtAGP18 acts as a co-receptor by binding to signaling molecules and bringing the signaling molecules to transmembrane proteins in the lipid rafts. Upon activation by the extracellular signals, the transmembrane proteins initiate signaling and lead to various intracellular events (e.g., phosphorylation similar to the two-component signaling system, influx of calcium ions). The different components of the AtAGP18 molecule and the various lipid components of lipid rafts and plasma membrane are shown in the boxed inset. Hpt, histidine phosphotransfer protein.     

Structural basis of transmembrane domain interactions in integrin signaling

Cell surface receptors of the integrin family are pivotal to cell adhesion and migration. The activation state of heterodimeric αβ integrins is correlated to the association state of the single-pass α and β transmembrane domains. The association of integrin αIIbβ3 transmembrane domains, resulting in an inactive receptor, is characterized by the asymmetric arrangement of a straight (αIIb) and tilted (β3) helix relative to the membrane in congruence to the dissociated structures. This allows for a continuous association interface centered on helix-helix glycine-packing and an unusual αIIb(GFF) structural motif that packs the conserved Phe-Phe residues against the β3 transmembrane helix, enabling αIIb(D723)β3(R995) electrostatic interactions. The transmembrane complex is further stabilized by the inactive ectodomain, thereby coupling its association state to the ectodomain conformation. In combination with recently determined structures of an inactive integrin ectodomain and an activating talin/β complex that overlap with the αβ transmembrane complex, a comprehensive picture of integrin bi-directional transmembrane signaling has emerged.Key words: cell adhesion, membrane protein, integrin, platelet, transmembrane complex, transmembrane signalingThe communication of biological signals across the plasma membrane is fundamental to cellular function. The ubiquitous family of integrin adhesion receptors exhibits the unusual ability to convey signals bi-directionally (outside-in and inside-out signaling), thereby controlling cell adhesion, migration and differentiation.^1–5 Integrins are Type I heterodimeric receptors that consist of large extracellular domains (>700 residues), single-pass transmembrane (TM) domains, and mostly short cytosolic tails (<70 residues). The activation state of heterodimeric integrins is correlated to the association state of the TM domains of their α and β subunits.^6–10 TM dissociation initiated from the outside results in the transmittal of a signal into the cell, whereas dissociation originating on the inside results in activation of the integrin to bind ligands such as extracellular matrix proteins. The elucidation of the role of the TM domains in integrin-mediated adhesion and signaling has been the subject of extensive research efforts, perhaps commencing with the demonstration that the highly conserved GFFKR sequence motif of α subunits (Fig. 1), which closely follows the first charged residue on the intracellular face, αIIb(K989), constrains the receptor to a default low affinity state.¹¹ Despite these efforts, an understanding of this sequence motif had not been reached until such time as the structure of the αIIb TM segment was determined.¹² In combination with the structure of the β3 TM segment¹³ and available mutagenesis data,^6,9,10,14,15 this has allowed the first correct prediction of the overall association of an integrin αβ TM complex.¹² The predicted association was subsequently confirmed by the αIIbβ3 complex structure determined in phospholipid bicelles,¹⁶ as well as by the report of a similar structure based on molecular modeling using disulfide-based structural constraints.¹⁷ In addition to the structures of the dissociated and associated αβ TM domains, their membrane embedding was defined^{12,13,16,18,19} and it was experimentally recognized that, in the context of the native receptor, the TM complex is stabilized by the inactive, resting ectodomain.¹⁶ These advances in integrin membrane structural biology are complemented by the recent structures of a resting integrin ectodomain and an activating talin/β cytosolic tail complex that overlap with the αβ TM complex,^20,21 allowing detailed insight into integrin bi-directional TM signaling.Open in a separate windowFigure 1Amino acid sequence of integrin αIIb and β3 transmembrane segments and flanking regions. Membrane-embedded residues^{12,13,16,18,19} are enclosed by a gray box. Residues 991–995 constitute the highly conserved GFFKR sequence motif of integrin α subunits.     

Challenges to understand plant responses to wind

Understanding plant response to wind is complicated as this factor entails not only mechanical stress, but also affects leaf microclimate. In a recent study, we found that plant responses to mechanical stress (MS) may be different and even in the opposite direction to those of wind. MS-treated Plantago major plants produced thinner more elongated leaves while those in wind did the opposite. The latter can be associated with the drying effect of wind as is further supported by data on petiole anatomy presented here. These results indicate that plant responses to wind will depend on the extent of water stress. It should also be recognized that the responses to wind may differ between different parts of a plant and between plant species. Physiological research on wind responses should thus focus on the signal sensing and transduction of both the mechanical and drought signals associated with wind, and consider both plant size and architecture.Key words: biomechanics, leaf anatomy, phenotypic plasticity, plant architecture, signal transduction thigmomorphogenesis, windWind is one of the most ubiquitous environmental stresses, and can strongly affect development, growth and reproductive yield in terrestrial plants.^1–3 In spite of more than two centuries of research,⁴ plant responses to wind and their underlying mechanisms remain poorly understood. This is because plant responses to mechanical movement themselves are complicated and also because wind entails not only mechanical effects, but also changes in leaf gas and heat exchange.^5–7 Much research on wind has focused primarily on its mechanical effect. Notably, several studies that determine plant responses to mechanical treatments such as flexing, implicitly extrapolate their results to wind effects.^8–10 Our recent study¹¹ showed that this may lead to errors as responses to wind and mechanical stimuli (in our case brushing) can be different and even in the opposite direction. In this paper, we first separately discuss plant responses to mechanical stimuli, and other wind-associated effects, and then discuss future challenges for the understanding of plant responses to wind.It is often believed that responses to mechanical stress (thigmomorphogenesis) entail the production of thicker and stronger plant structures that resist larger forces. This may be true for continuous unidirectional forces such as gravity, however for variable external forces (such as wind loading or periodic flooding) avoiding such mechanical stress by flexible and easily reconfigurable structures can be an alternative strategy.^12–14 How plants adapt or acclimate to such variable external forces depends on the intensity and frequency of stress and also on plant structures. Reduced height growth is the most common response to mechanical stimuli.^15,16 This is partly because such short stature increases the ability of plants to both resist forces (e.g., real-locating biomass for radial growth rather than elongation growth), and because small plants experience smaller drag forces (Fig. 1). Some plant species show a resistance strategy in response to mechanical stress by increasing stem thickness^1,10 and tissue strength.⁷ But other species show an avoidance strategy by a reduction in stem or petiole thickness and flexural rigidity in response to MS.^11,15–18 These different strategies might be associated with plant size and structure. Stems of larger plants such as trees and tall herbs are restricted in the ability to bend as they carry heavy loads^7,10,19 (Fig. 1). Conversely short plants are less restricted in this respect and may also be prone to trampling for which stress-avoidance would be the only viable strategy.^18,20 Systematic understanding of these various responses to mechanical stress remains to be achieved.Open in a separate windowFigure 1A graphical representation of how wind effects can be considered to entail both a drying and a mechanical effect. Adaptation or acclimation to the latter can be through a force resistance strategy or a force avoidance strategy, the benefit of which may depend on the size and architecture of plants as well as the location of a given structure within a plant.Wind often enhances water stress by reducing leaf boundary layers and reduces plant temperature by transpiration cooling. The latter effect may be minor,¹¹ but the former could significantly affect plant development. Anten et al. (2010) compared phenotypic traits and growth of Plantago major that was grown under mechanical stimuli by brushing (MS) and wind in the factorial design. Both MS and wind treatments reduced growth and influenced allocation in a similar manner. MS plants, however, had more slender petioles and narrower leaf blades while wind exposed plants exhibited the opposite response having shorter and relatively thicker petioles and more round-shaped leaf blades. MS plants appeared to exhibit stress avoidance strategy while such responses could be compensated or overridden by water stress in wind exposure.¹¹ A further analysis of leaf petiole anatomy (Fig. 2) supports this view. The vascular fraction in the petiole cross-section was increased by wind but not by MS, suggesting that higher water transport was required under wind. Our results suggest that drying effect of wind can at least to some extent override its mechanical effect.Open in a separate windowFigure 2Representative images of petiole cross-sections of Plantago major grown in 45 days in continuous wind and/or mechanical stimuli (A–D). Petiole cross-section area (E) and vascular bundle fraction in the cross-section of petiole (F). mean + SD (n = 12) are shown. Significance levels of ANOVA; ***p < 0.001, **p < 0.01, *p < 0.05, ns p > 0.05.Physiological knowledge on plant mechanoreception and signal transduction has been greatly increased during the last decades. Plants sense mechanical stimuli through membrane strain with stretch activated channels²¹ and/or through some linker molecules connecting the cell wall, plasma membrane and cytoskeleton.^4,22,23 This leads to a ubiquitous increase in intracellular Ca²⁺ concentration. The increased Ca²⁺ concentration is sensed by touch induced genes (TCHs),^24,25 which activates downstream transduction machineries including a range of signaling molecules and phytohormones, consequently altering physiological and developmental processes.²⁶ Extending this knowledge to understand plant phenotypic responses to wind however remains a challenge. As responses to wind have been found to differ among parts of a plant (e.g., terminal vs. basal stem) and also across species, physiological studies should be extended to the whole-plant as integrated system rather than focusing on specific tissue level. Furthermore to understand the general mechanism across species, it is required to study different species from different environmental conditions. Advances in bioinformatics, molecular and physiological research will facilitate cross-disciplinary studies to disentangle the complicated responses of plants to wind.     

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.     

A new callose function: Involvement in differentiation and function of fern stomatal complexes

Callose in polypodiaceous ferns performs multiple roles during stomatal development and function. This highly dynamic (1→3)-β-D-glucan, in cooperation with the cytoskeleton, is involved in: (a) stomatal pore formation, (b) deposition of local GC wall thickenings and (c) the mechanism of stomatal pore opening and closure. This behavior of callose, among others, probably relies on the particular mechanical properties as well as on the ability to form and degrade rapidly, to create a scaffold or to serve as a matrix for deposition of other cell wall materials and to produce fibrillar deposits in the periclinal GC walls, radially arranged around the stomatal pore. The local callose deposition in closing stomata is an immediate response of the external periclinal GC walls experiencing strong mechanical forces induced by the neighboring cells. The radial callose fibrils transiently co-exist with radial cellulose microfibrils and, like the latter, seem to be oriented via cortical MTs.Key words: callose, cytoskeleton, fern stomata, guard cell wall thickening, stomatal function, stomatal pore formationCallose represents a hemicellulosic matrix cell wall component, usually of temporal appearance, which is synthesized by callose synthases, enzymes localized in the plasmalemma and degraded by (1→3)-β-glucanases.^1–4 It consists of triple helices of a linear homopolymer of (1→3)-β-glucose residues.^5–7 The plant cell is able to form and degrade callose in a short time. On the surface of the plasmolyzed protoplast a thin callose surface film may arise within seconds.⁸ Callose is the only cell wall component that is implicated in a great variety of developmental plant processes, like cell plate formation,^9–11 microspore development,^12–14 trafficking through plasmodesmata,^15,16 formation and closure of sieve pores,¹⁶ response of the plant cells to multiple biotic and abiotic stresses,^4,5 establishment of distinct “cell cortex domains”,¹⁷ etc.Despite the widespread occurrence of callose, its general function(s) is (are) not well understood (reviewed in refs. ⁴ and ⁵). It may serve as: a matrix for deposition of other cell wall materials, as in developing cell plates;⁹ a cell wall-strengthening material, as in cotton seed hairs and growing pollen tubes;¹⁸ a sealing or plugging material at the plasma membrane of pit fields, plasmodesmata and sieve plate pores;¹⁶ a mechanical obstruction to growth of fungal hyphae or a special permeability barrier, as in pollen mother cell walls and muskmelon endosperm envelopes.^4,19,20 The degree of polymerization, age and thickness of callose deposits may cause variation in its physical properties.⁵Evidence accumulated so far showed that a significant number of ferns belonging to Polypodiales and some other fern classes forms intense callose deposits in the developing GC wall thickenings.^21–28 This phenomenon has not been observed in angiosperm stomata, although callose is deposited along the whole surface of the young VW and in the VW ends of differentiating and mature stomata (our unpublished data; reviewed in refs ²⁹ and ³⁰).Stomata are specialized epidermal bicellular structures (Fig. 1A) regulating gas exchange between the aerial plant organs and the external environment. Their appearance in the first land plants was crucial for their adaptation and survival in the terrestrial environment. The constituent GCs have the ability to undergo reversible changes in shape, leading to opening and closure of the stomatal pore (stomatal movement). The mechanism by which GCs change shape is based on: (a) the particular mechanical properties of GC walls owed to their particular shape, thickening, fine structure and chemical composition and (b) the reversible changes in vacuole volume, in response to environmental factors, through fairly complicated biochemical pathways.^30–33Open in a separate windowFigure 1(A) Diagrammatic representation of an elliptical stoma. (B–E) Diagram to show the process of stomatal pore formation in angiosperms (B and C) and Polypodiales ferns (D and E). The arrows in (B) indicate the forming stomatal pore. DW, dorsal wall; EPW, external periclinal wall; GC, guard cell; IPW, internal periclinal wall; ISP, internal stomatal pore; PE polar ventral wall end; VW, ventral wall.The present review is focused on the multiple-role of callose in differentiating and functioning fern stomata, as they are substantiated by the available information, including some unpublished data, and in particular in: stomatal pore formation, deposition of GC wall thickenings and opening and closure of the stomatal pore. The mode of deposition of fibrillar callose deposits in GC walls and the mechanism of their alignment are also considered.     

Step by Step: Deciphering Ion Transport in the Root Xylem Parenchyma

Proton pumps produce electrical potential differences and differences in pH across the plasma membrane of cells which drive secondary ion transport through sym- and antiporters. We used the patch-clamp technique to characterize an H⁺-pump in the xylem parenchyma of barley roots. This cell type is of special interest with respect to xylem loading. Since it has been an ongoing debate whether xylem loading is a passive or an active process, the functional characterization of the H⁺-pump is of major interest in the context of previous work on ion channels through which passive salt efflux into the xylem vessels could occur. Cell-type specific features like its Ca²⁺ dependence were determined, that are important to interpret its physiological role and eventually to model xylem loading. We conclude that the electrogenic pump in the xylem parenchyma does not participate directly in the transfer of KCl and KNO₃ to the xylem but, in combination with short-circuiting conductances, plays a crucial role in controlling xylem unloading and loading through modulation of the voltage difference across the plasma membrane. Here, our recent results on the H⁺ pump are put in a larger context and open questions are highlighted.Key Words: plant nutrition, H⁺-ATPase, anion conductance, K⁺ channel, electrophysiology, signaling networkThe root xylem parenchyma is of major interest with respect to nutrient (and signal) traffic between root and shoot. One of its main functions appears to be xylem loading. However, the cell walls of the vascular tissue provide apoplastic paths between xylem and phloem that represent the upward and downward traffic lanes, allowing nutrient circulation¹ (Fig. 1). Therefore mechanisms for ion uptake and for ion release must exist side by side. In the last 15 years major progress has been made in the investigation of transport properties of xylem-parenchyma cells, and both uptake and release channels and transporters were identified. Today, we have good knowledge on the role of K⁺ and anion conductances in xylem loading with salts.² Note, that from the functionally well characterized conductances only the molecular structure of K⁺ channels is known. In contrast, many transporters are identified on the molecular level, but functional data are scarce.Open in a separate windowFigure 1Distribution of tissues in the periphery of the stele. The stippled area marks the region from which early metaxylem protoplasts originated. E, Endodermis with Casparian strip; eMX, ‘early’ metaxylem vessel; IMX, ‘late’ metaxylem vessel; Mph, metaphloem (sieve tube); Pph, protophloem (sieve tube); P, pericycle; Cx, cortex. Symplasmic and apoplasmic transport routes are indicated in red and black, respectively. The Casparian strip prevents apoplastic transport into the stele. Plasmodesmata are shown exemplarily for the indicated symplastic pathway. All cells of the symplast are connected via plasmodesmata. Sites of active uptake into the root symplast and of release into the stelar apoplast are indicated by a black and an orange arrow. Modified from Wegner and Raschke, 1994.³A challenging question to deal with was the dispute about xylem loading with ions being a passive or active process. While it is clear that energy through electrogenic H⁺ efflux is needed to take up nutrient ions from the soil against their electrochemical gradient into the cortical symplast, it has been a matter of debate if ion release into xylem vessels also is energy-linked or if the electrochemical potentials of ions are raised high enough to allow a thermodynamically passive flux.^2,3 The Casparian strip prohibits apoplastic transport of nutrients into the stele and electrically insulates the stelar from the cortical apoplast. Therefore the electrical potential difference of the cells in the xylem parenchyma could be independent from the cortical potential difference but be subject to control, for instance, from the shoot.⁴ Indeed, evidence points to xylem loading as a second control point in nutrient transfer to the shoot.^5,6 The identification and characterization of K⁺ and anion conductances clearly showed that release of KCl and KNO₃ into the xylem can be passive through voltage-dependent ion channels.^2,3,7–9 No need appeared for a pump energizing the transfer of salts to the xylem.However, H⁺ pumps are ubiquitous. H⁺-ATPases are encoded by a multigene family and heterologous expression in yeast showed that isoforms have distinct enzymatic properties.^10,11 As the example of the amino acid transporter AAP6 from the xylem parenchyma shows, a cell-type specific functional characterization of transporters is essential to draw conclusions on their physiological role. AAP6 is the only member of a multigene family with an affinity for aspartate in the physiologically relevant range. The actual apoplastic concentration of amino acids and the pH will determine what is transported in vivo.^12,13 Xylem-parenchyma cells of barley roots were strongly labelled by antibodies against the plasma membrane H⁺-ATPase.¹⁴ In a recent publication in Physiologia Plantarum we report the functional analysis of the electrogenic pump from the plasma membrane of xylem parenchyma from barley roots that was done with the patch-clamp technique after specific isolation of protoplasts from this cell type. It displayed characteristics of an H⁺-ATPase: current-voltage relationships were characteristic for a ‘rheogenic’ pump¹⁵ and currents were stimulated by fusicoccin or by an enlarged transmembrane pH gradient and inhibited by dicyclohexylcarbodiimide (DCCD). Importantly, it also showed distinct characteristics. Neither intracellular pH nor the intracellular Ca²⁺ concentration affected its activity. Noteworthy, K⁺ and anion conductances from the same cell type are controlled by intracellular [Ca²⁺]^7,9 (Fig. 2). It was proposed that the effect of abscisic acid (ABA) on anion conductances is mediated via an increase in the cytosolic Ca²⁺ concentration.¹⁶ Very likely stelar H⁺ pumps are stimulated by ABA.¹⁷ Thus, a Ca²⁺ independent control has to be hypothesized in this case.Open in a separate windowFigure 2Control of ion conductances in the plasma membrane of xylem-parenchyma cells. Arrowheads indicate stimulation and bars indicate inhibition by an increase in cytosolic [Ca²⁺],^7,9,16 by ABA,^16,17,21 by cytosolic and apoplastic acidification,^4,22 by G-proteins²³ and by an increase in apoplastic [K⁺]⁷ and [NO₃⁻].²⁴ Apoplastic [K⁺] and [NO₃⁻] modify the voltage dependence exerting negative feedback on K⁺ efflux and a positive feedback on NO₃⁻ efflux. Abscisic acid has an immediate effect on ion channel activity, most likely via [Ca²⁺], and causes a change in gene expression as indicated by circles (up) and bars (down). ABA perception is not clear. A Ca²⁺ influx could occur through a hyperpolarization activated cation conductance (HACC).^16,25 Cation transporters are NORC, nonselective cation conductance, KORC, K⁺-selective outwardly rectifying conductance (=SKOR⁸), and KIRC, K⁺-selective inwardly rectifying conductance, and anion conductances with different voltage-dependencies and gating characteristics are X-QUAC, quickly activating anion conductance, X-SLAC, slowly activating anion conductance, and X-IRAC, inwardly rectifying anion channel.^2,3,9,16,26 Transported ions and direction of flux are plotted.To date, we know that besides Ca²⁺ and abscisic acid also the pH, nonhydrolyzable GTP analogs and extracellular NO₃⁻ and K⁺ affect membrane transport capacities of root xylem-parenchyma cells (Fig. 2). Other control mechanisms by metabolites, the redox potential and phytohormones have to be included, especially if they represent signals in xylem loading or root-shoot communication. The composition of the xylem sap changes during the course of a day, depending on nutrient supply and various stresses, and the apoplastic ion concentration is considered to be an important factor in ion circulation.^6,18,19 ABA is such a signal. It is known to increase solute accumulation within the root by inhibiting release of ions into the xylem.¹⁷ Any change in transport activity has an impact on the membrane potential. This again determines whether salt release or uptake takes place. Passive salt release is restricted to a limited range of membrane potentials in which conductances for anions and cations are active simultaneously, that is with depolarization. Negative membrane voltages will be required for reabsorption of NO₃⁻ by a putative NO₃⁻/H⁺-symporter and for the uptake of K⁺ and amino acids.^3,13 As shown in our recent paper, the balance between the activities of the H⁺-pump and the anion conductances could affect the position between a depolarized and a hyperpolarized state of the parenchymal membrane. Thus, H⁺ pump activity is crucial in membrane voltage control. Furthermore, the simultaneous activities of H⁺ pumps and anion conductances make the generation of a high pH gradient possible, whilst maintaining electroneutrality. The proton gradient could be used for ion transport through cotransporters and antiporters as suggested for the loading of borate into the xylem through the boron transporter BOR1.²⁰ So we are on the way to decipher xylem loading in roots and this exciting field will also provide information about small-scale nutrient cycling and root-shoot communication. To determine how the activities of pumps, channels and transporters are adjusted among each other is the next challenge. Further insight has to be obtained by experimentation as well as by biophysical modeling.     

Regulation of stomatal opening by the guard cell expansin AtEXPA1

Stomatal movement is strictly regulated by various intracellular and extracellular factors in response environmental signals. In our recent study, we found that an Arabidopsis guard cell expressed expansin, AtEXPA1, regulates stomatal opening by altering the structure of the guard cell wall. This addendum proposes a mechanism by which guard cell expansins regulate stomatal movement.Key words: expansin, stomatal movement, AtEXPA1, guard cell, wall looseningStomatal movement is the most popular model system for cellular signaling transduction research. A complicated complex containing many proteins has been proposed to control stomatal responses to outside stimuli. The known regulation factors are primarily located in the nucleus, cytoplasm, plasma membrane and other intracellular organelles.^1,2 Although the cell wall structure of the stomata is different from that of other cells,^3,4 the presence of stomatal movement regulation factors in the cell wall has seldom been reported in reference ⁵. In our previous work, we found that extracellular calmodulin stimulates a cascade of intracellular signaling events to regulate stomatal movement.⁶ The involvement of this signaling pathway is the first evidence that cell wall proteins play an important role in regulation of stomatal opening. Cell wall-modifying factors constitute a major portion of cell wall proteins. However, the role of these factors in the regulation of stomatal movement is not yet known.Expansins are nonenzymatic proteins that participate in cell wall loosening.^7–9 Expansins were first identified as “acid-growth” factors because they have much higher activities at acidic pHs.^10,11 It has been reported that expansins play important roles in plant cell growth, fruit softening, root hair emergence and other developmental processes in which cell wall loosening is involved.^7–9,12,13 Wall loosening is an essential step in guard cell swelling and the role of stomatal expansins was investigated. AtEXPA1 is an Arabidopsis guard-cell-specific expansin.^13,14 Over-expressing AtEXPA1 increases the rate of light-induced stomatal opening,^14,15 while a potential inhibitor of expansin activity, AtEXPA1 antibody, reduces the sensitivity of stomata to stimuli.¹⁴ We showed that the transpiration rate and the photosynthesis rate in plant lines overexpressing AtEXPA1 were nearly two times the rates for wild-type plants (Fig. 1). These in plant data revealed that expansins accelerated stomatal opening under normal physiological conditions. In addition, the increases in the transpiration and photosynthesis rates strongly suggested the possibility of exploiting expansin-regulated stomatal sensitivity to modify plant drought tolerance. Compared with the effect of hydrolytic cell wall enzymes, the destruction of cell wall structures induced by expansins is minimal. In addition, it is very difficult to directly observe the changes in the guard cell wall structure caused by expansins during stomatal movement. Our recent work showed that, in AtEXPA1-overexpressing plants, the volumetric elastic modulus is lower than in wild-type plants,¹⁴ which indicates the wall structure was loosened and that the cell wall was easier to extend. Taken together, our data suggest that expansins participate in the regulation of stomatal movement by modifying the cell walls of guard cells.Open in a separate windowFigure 1Effects of AtEXPA1 overexpression on transpiration rates and photosynthesis rates. The transpiration rate (left) and photosynthesis rate (right) of wild-type and transgenic AtEXPA1 lines were measured at 10:00 AM in the greenhouse after being watered overnight. The illumination intensity was 180 µmol/m²·s. Bars represent the standard error of the mean of at least five plants per line.It is well known that the activation of proton-pumping ATPase (H⁺-ATPase) in the plasma membrane is an early and essential step in stomatal opening.¹⁶ The action of the pump results in an accumulation of H⁺ outside of the cell, increases the inside-negative electrical potential across the plasma membrane and drives potassium uptake through the voltage-gated, inward-rectifying K⁺ channels.^17–19 The main function of the H⁺ pump is well accepted to create an electrochemical gradient across the plasma membrane; however, the other result is the acidification of the guard cell wall, which may also contribute to stomatal opening. A possible mechanism responsible for this effect is as follows. Expansins are in an inactive state when the stomata are in the resting state. Stomatal opening signals induce wall acidification and activate expansins. Then, the expansins move along with cellulose microfibrils and transiently break down hydrogen bonding between hemicellulose and the surface of cellulose microfibrils,^20,21 facilitating the slippage of cell wall polymers under increasing guard cell turgor pressure. The guard cell then swells and the stomata open (Fig. 2).Open in a separate windowFigure 2Model of how guard cell wall expansins regulate stomatal opening. Environmental stimuli, e.g., light, activate guard cell plasma membrane H⁺-ATPases to pump H⁺ into the extracellular wall space. The accumulation H⁺ acidifies the cell wall and induces the activation of expansin. The active expansin disrupts non-covalent bonding between cellulose microfibrils and matrix glucans to enable the slippage of the cell wall. The wall is loosened coincident with guard cell swelling and without substantial breakdown of the structure.Although our results indicate that AtEXPA1 regulates stomatal movement, the biochemical and structural mechanism by which AtEXPA1 loosens the cell wall remains to be discovered. It remains to figure out the existing of other expansins or coordinators involving in this process. In addition, determining the roles of expansins and the guard cell wall in stomatal closing is another main goal of future research.     

Is chloroplast import of photosynthesis proteins facilitated by an actin-TOC-TIC-VIPP1 complex?

Actin filaments are major components of the cytoskeleton that interact with chloroplast envelope membranes to allow chloroplast positioning and movement, stromule mobility and gravitropism perception. We recently reported that Toc159, a component of the TOC complex of the chloroplast protein import apparatus, interacts directly with actin. The interaction of Toc159 and actin was identified by co-immunoprecipitation and co-sedimentation experiments with detergent-solubilised pea chloroplast envelope membranes. In addition, many of the components of the TOC-TIC protein import apparatus and VIPP1 (vesicle-inducing protein in plastids 1) were identified by mass spectroscopy in the material co-immunoprecipitated with antibodies to actin. Toc159 is the receptor for the import of photosynthesis proteins and VIPP1 is involved in thylakoid membrane formation by inducing vesicle formation from the chloroplast inner envelope membrane, suggesting we may have identified an actin-TOCTIC-VIPP1 complex that may provide a means of channeling cytosolic preproteins to the thylakoid membrane. The interaction of Toc159 with actin may facilitate exchange between the putative soluble and membrane forms of Toc159 and promote the interaction of cytosolic preproteins with the TOC complex.Key words: actin, chloroplast, protein import, TOC complex, TIC complex, VIPP1Actin is a ubiquitous protein of eukaryotic cells and a major component of the cytoskeleton as microfilaments. In plant cells, plastids are closely associated with actin microfilaments.^1,2 A direct interaction of plastids with the actin cytoskeleton has been postulated to anchor chloroplasts at appropriate intracellular positions,³ to support chloroplast light-intensitydependent movement,⁴ to facilitate plastid stromule (stroma-filled tubule) mobility^5,6 and to participate in gravity perception.⁷ The known proteins implicated in plastid-actin interaction are CHUP1 (chloroplast unusual positioning 1), a protein exclusively targeted to the chloroplast outer envelope membrane that is essential for chloroplast anchorage to the plasma membrane,⁸ and myosin XI proteins that play a role in stromule movement9 and in gravitropism.^10,11 Recently, we found that Toc159 also interacts with actin.¹²Toc159 is a component of the TOC complex, which is part of the chloroplast protein translocation apparatus. This apparatus consists of two membrane protein complexes that associate to allow translocation of nucleus-encoded proteins from the cytoplasm to the interior stromal compartment (reviewed in ref. ¹³). The translocon at the outer envelope membrane of chloroplasts (TOC complex) mediates the initial recognition of preproteins and their translocation across the outer membrane.¹⁴ The translocon at the inner envelope membrane of chloroplasts (TIC complex) physically associates with the TOC complex and provides the membrane translocation channel for the inner membrane. In addition, the TOC and TIC complexes interact with a set of molecular chaperones (ClpC and Hsp70), which assist the transfer of imported proteins^15–17 (Fig. 1).Open in a separate windowFigure 1Schematic diagram of Toc159-actin interactions and the import of photosynthesis proteins. Toc159, linked to actin by its A-domain, recruits a newly synthesized photosynthesis preprotein by its G-domain. Actin filaments facilitate Toc159 movement to the chloroplast outer envelope membrane for integration into the TOC complex. The core TOC complex is formed by Toc159, Toc34 and Toc75. Tic22 acts to facilitate the passage of preproteins across the intermembrane space and interacts with the TIC complex. The core TIC complex is composed of Tic110, Tic20 and/or Tic21, and Tic40. The Tic110 protein recruits stromal molecular chaperone ClpC. On arrival in the stroma, the transit peptide is cleaved by SPP, and other chaperones (Hsp90 or Hsp70) may assist in the folding. VIPP1 interacts with the chaperones and polymerises, inducing chloroplast inner envelope membrane budding, leading to thylakoid formation.The interaction between actin and Toc159 was identified by co-immunoprecipitation and co-sedimentation experiments with detergent-solubilised pea chloroplast envelope membranes, and confirmed with Toc159 expressed in Escherichia coli. In addition, many other components of the TOC-TIC protein import apparatus were co-immunoprecipitated by antibodies to actin and co-sedimented with added F-actin filaments.¹² Using mass spectrometry, we identified the principal components of the TOC complex (Toc159, Toc75 and Toc34) and three accepted components of the TIC core complex (Tic110, Tic40 and ClpC). The presence of Tic20/21 and Tic22 could not be examined because they migrate in the same position on SDS-PAGE as the light chains of antibody molecules but, since they are involved in linking the TOC and TIC complexes,⁶ they may also be part of the complex with actin.The identification of the region of Toc159 that interacts with actin is an important feature to help establish whether any of the other Toc159 isoforms (such as Toc132 and Toc120) are likely to interact with actin. Toc159 family proteins are composed of three different domains: the A (acidic) domain, the G (GTPase) domain and the M (membrane) domain.¹⁸ The interaction of Toc159 with actin appears most likely to be through the A-domain; the G-domain did not co-sediment with actin filaments¹² and the M-domain is embedded in the chloroplast envelope outer membrane and therefore is unlikely to be accessible to actin. Toc132 and Toc120 have shorter A-domains than Toc159 and this may affect their ability to bind actin. Although all the Toc159 isoforms are implicated in chloroplast protein import, Toc132 and Toc120 are involved in the import of chloroplast housekeeping proteins and Toc159 is specialized for the import of photosynthesis proteins.¹⁸ For import of photosynthesis proteins, two models have been proposed for preprotein recognition by the TOC complex: the ‘targeting model’ where the newly synthesized preprotein is first bound by a free cytosolic form of Toc159, and the ‘motor model’ where the transit peptide is first phosphorylated and then bound to Toc34 associated with the other TOC subunits in the outer envelope membrane.¹³ In support of the first model, Toc159 has been reported to exist in both cytosolic and membrane-bound forms^19,20 and the soluble form of Toc159 is able to bind preproteins.^20,21 Toc159 is proposed to be the major point of contact for preproteins during the early stages of protein import through its A-domain.²² The interaction of Toc159 with actin might provide a means to favor exchange between the putative soluble and membrane forms of Toc159 and potentially facilitate chloroplast photosynthesis protein import (Fig. 1).Several features of this model require additional experimental evidence. The involvement of a soluble form of Toc159 is highly controversial,¹³ and evidence for a physiological role in vivo is required. Experimental evidence for a facilitating role of the actin cytoskeleton in chloroplast protein import is also required. Does the presence of a basket of actin filaments surrounding the chloroplasts² provide a means of concentrating cytosolic Toc159 in the vicinity of the chloroplasts? Or do actin filaments provide a trackway for movement of Toc159 to or from chloroplasts? Myosin, the motor protein for movement along actin filaments, was not detected in the co-immunoprecipitated complex, but this does not necessarily rule out its involvement.VIPP1 was also identified in the complex with actin. VIPP1 is involved in thylakoid membrane formation by vesicle formation from the chloroplast inner envelope membrane²³ and the quantity of thylakoid membrane proteins is closely correlated to the amount of VIPP1 in chloroplasts.²⁴ VIPP1 is also known to interact with Hsp70 and Hsp90 chaperones^25–27 and these chaperones may associate with the stromal face of the TIC complex to support protein folding.¹⁵ This raises the possibility that an actin-TOC-TIC-VIPP1 complex may facilitate thylakoid formation by channeling the import of thylakoid-located photosynthesis proteins through the chloroplast envelope membrane into vesicles directed to the thylakoid membrane (Fig. 1).Our study of actin-binding proteins in the chloroplast envelope membrane may have provided an initial glimpse at previously unrecognized mechanisms facilitating the import of photosynthesis proteins by chloroplasts. The formation of an actin-TOC-TIC-VIPP1 complex may provide a means of channeling cytosolic preproteins to the thylakoid membrane.     

Where and how does phototropin transduce light signals in the cell?

Light plays pivotal roles as an important environmental signal in plant growth and development. In Arabidopsis, phototropin 1 (phot1) and 2 (phot2) are the photoreceptors that mediate phototropism, chloroplast relocation, stomatal opening and leaf flattening, in response to blue light. However, little is known about how phototropins transduce the signals after the light is perceived. Changes induced by blue light in terms of intracellular localization patterns of phot2 in Arabidopsis were examined. Phot2 distributed uniformly in the plasma membrane under dark conditions. Upon irradiation with blue light, some of the phot2 associated with the Golgi apparatus. It was also shown that the kinase domain, but not the photosensory domain, is required for a plasma membrane and Golgi localization. Furthermore a kinase fragment, lacking the photosensory domain, constitutively triggered physiological responses in planta. Thus, the plasma membrane and the Golgi apparatus appear to be the most likely sites for the initial step of phot2 signal transduction. The Golgi apparatus facilitates vesicle trafficking and delivery of membrane proteins to the required locations in the cell. Therefore, this study implicates the regulation of vesicle trafficking by the Golgi apparatus as a mechanism by which phot2 elicits its cellular responses.Key words: Golgi apparatus, kinase, light signal transduction, photoreceptor, phototropin, vesicle traffickingA range of physiological responses in plants is brought about by blue (390–500 nm) and ultraviolet-A (320–390 nm) light. Phototropin, one of major classes of blue light photoreceptors in plants, mediates responses such as phototropism, chloroplast relocation, light-induced stomatal opening and leaf flattening.^1–6 The dicotyledon Arabidopsis, possesses two phototropins, termed phot1 and phot2, which have both overlapping and distinct functions.^5,7 Phototropins consist of two functional domains, a N-terminal photosensory domain, containing two LOV (Light, Oxygen, Voltage) domains (LOV1 and LOV2) and a flavin-mononucleotide (FMN) chromophore and a regulatory serine/threonine kinase domain at the C-terminus.⁸To understand the mechanism of phototropin signal transduction, we expressed phot2 derivatives with translationally-fused green fluorescent protein (GFP) in a phot1phot2 double mutant in a wild type background in Arabidopsis.^9,10 Phototropin is a membrane- associated protein lacking a membrane spanning domain.⁸ Phot1 fused to GFP (P1G) is mainly localized to the plasma membrane, regardless of the light conditions.⁶ This property was retained when phot2 was fused to GFP (P2G).⁹ A part of P2G associates with punctate structures in the cytoplasm in response to blue light. The punctate P2G colocalized with KAM1ΔC:mRFP, a Golgi marker, we therefore conclude that phot2 associated with the Golgi apparatus in a blue light-dependent manner.⁹ This association was observed even in the presence of brefeldin A (BFA), an inhibitor of the vesicle trafficking.⁹To determine which domain of phot2 is responsible for the Golgi association, fragments of phot2 were fused to GFP and expressed in protoplasts.⁹ The N-terminal fragment fused to GFP (P2NG) was distributed uniformly in the cytoplasm. By contrast, the C-terminal fragment fused to GFP (P2CG) localized to both plasma membrane and punctate structures. The latter was shown to be the Golgi apparatus with the aid of the Golgi marker, KAM1ΔC:mRFP.⁹ These observations were corroborated from data using transgenic plants.¹⁰ Hence the C-terminal kinase domain, but not the N-terminal photo-sensory domain, is essential for the association of phot2 with the plasma membrane and the Golgi apparatus.The Golgi network is a key player in vesicle trafficking, to and from ER, vacuoles, trans-Golgi network, endosome and the plasma membrane.¹¹ Membrane spanning proteins are delivered and recycled through the Golgi apparatus. Among the membrane spanning proteins that are especially interesting, with respect to phototropin function, are auxin carriers such as PIN proteins. Phototropic curvature, which is under the control of phototropin, is believed to be caused by an uneven distribution of auxin.¹² The intracellular distribution of PIN proteins is maintained and regulated by vesicle trafficking.¹³ Indeed, factors such as ADP-ribosylation factor1 (ARF1) and guanine-nucleotide exchange factors (GEFs), which are involved in vesicle trafficking, are indispensable for the proper distribution of PIN proteins.^14–17 It is intriguing that a light stimulus alters the distribution pattern of PIN proteins.¹⁸ Hence, a fascinating possibility arises that phot2 alters the intracellular distribution of PIN proteins by regulating vesicle trafficking at the level of the Golgi apparatus.Phototropins are members of the subfamily VIII of AGC kinases.¹⁹ Interestingly, PINOID, another member of the subfamily, is localized at the cell periphery and regulates the apical-basal polar distribution of PIN proteins.^20–22 Accordingly, overexpression of PINOID disturbs the auxin distribution in transgenic plants.^23,24 The kinase fragment of phototropin exhibits constitutive kinase activity in vitro.²⁵ Interestingly, the auxin distribution is disturbed in plants expressing P2CG, as is the case with PINOID.¹⁰ Hence, both PINOID and phot2 might alter the PIN protein distribution in the cell through a common mechanism, in response to distinct stimuli.To date, no authentic substrate has been described for any of the AGC VIII kinases.¹⁹ Considering the localization pattern of phototropins, the substrates are most likely to reside in the plasma membrane and/or the Golgi apparatus. NPH3, RPT2 and PKS1 are downstream factors for phototropic responses,^26–28 all associating with the plasma membrane. Although they interact preferentially with the N-terminal rather than the C-terminal domain of phot1,^26,29 it is also possible that the C-terminal kinase domain interacts transiently with these factors leading to their phosphorylation. However, at present the molecular functions of NPH3, RPT2 and PKS1 remain unclear and await future investigation.Although both phot1 and phot2 are localized to the plasma membrane, punctate structures are yet to be described for P1G. Instead, a part of phot1-GFP is released from the plasma membrane to the cytosol in response to a light stimulus.⁶ We recently reexamined the intracellular localization of P1G. A specific network-like structure in the cytoplasm in addition to intense plasma membrane staining was observed (Fig. 1). A similar pattern was observed for P2G although it is less clear.⁹ Hence, both phot1 and phot2 might be associating with a structure in the cytoplasm that has yet to be described, and which might be another site of phototropin signaling in the cell.Open in a separate windowFigure 1A light-induced network-like distribution pattern of P1G in the cytoplasm. The P1G seedlings grown under dark conditions⁶ were incubated in MS solution (diluted 50%) without (upper panels) or with (lower panels) 100 µM BFA. The cells were inspected with a confocal laser scanning microscope. Images taken before (left) or after (right) blue light illumination at 48 µmol m⁻² sec⁻¹ are shown. Bar = 10 µm.P2CG elicits some phototropin responses without a light stimulus.¹⁰ That is, chloroplasts were in the avoidance position and stomata opened without a blue light stimulus in the P2CG overexpressing plants. It is a fascinating possibility that phototropin elicits those responses through the regulation of vesicle trafficking, although other possibilities exist. Stomata open as the result of phosphorylation of the plasma membrane H⁺-ATPase³⁰ and it is unlikely that the vesicle trafficking is directly involved in this regulatory process. It is possible to conjecture that vesicle trafficking affects chloroplast positioning but how this would work remains to be determined. Overall how a single photoreceptor such as phototoropin might regulate diverse physiological responses awaits future study.     

pH signature for the responses of arbuscular mycorrhizal fungi to external stimuli

Environmental and developmental signals can elicit differential activation of membrane proton (H⁺) fluxes as one of the primary responses of plant and fungal cells. In recent work,¹ we could determine that during the presymbiotic growth of arbuscular mycorrhizal (AM) fungi specific domains of H⁺ flux are activated by clover root factors, namely host root exudates or whole root system. Consequently, activation on hyphal growth and branching were observed and the role of plasma membrane H⁺-ATPase was investigated. The specific inhibitors differentially abolished most of hyphal H⁺ effluxes and fungal growth. As this enzyme can act in signal transduction pathways, we believe that spatial and temporal oscillations of the hyphal H⁺ fluxes could represent a pH signature for both early events of the AM symbiosis and fungal ontogeny.Key words: H⁺-specific vibrating probe, pH signatures, arbuscular mycorrhiza, pH signalling, Gigaspora margaritaThe 450-million-year-old symbiosis between the majority of land plants and arbuscular mycorrhizal (AM) fungi is one of the most ancient, abundant and ecologically important symbiosis on Earth.^2,3The development of AM interaction starts before the physical contact between the host plant roots and the AM fungus. The hyphal growth and branching are induced by the root factors exudated by host plants, followed by the formation of appressorium leading to the hyphal penetration in the root system. These root factors seems to be specifically synthesized by host plants, since exudates from non-host plants are not able to promote neither hyphal differentiation nor appressorium formation.^4,5 Most root exudates contain several host signals or better, active compounds including flavonoids^6,19 and strigolactones,^7,8 however many of them are not yet known.Protons (H⁺) may have an important role on the fungal growth and host signal perception.¹ In plant and fungal cells, H⁺ can be pumped out through two different mechanisms: (1) the activity of the P-type plasma membrane (PM) H⁺-ATPase⁹ and (2) PM redox reactions.¹⁰ The proportional contribution from both mechanisms is not known, but in most plant cells the PM H⁺-ATPase seems to be the major responsible by the H⁺ efflux across plasma membrane. AM Fungal cells also energize their PM using P-type H⁺-pumps quite similar to the plant ones. Indeed, some genes codifying isoforms of P-type H⁺-ATPase have been isolated of AM fungi,^11–13 and AM fungal ATP hydrolysis activity was shown by cytochemistry, localized mainly in the first 70 µm from the germ tube tip.¹⁴ This structural evidence correlates with data obtained by H⁺-specific vibrating probe (Fig. 1A and B), which indicates that the H⁺ efflux in Gigaspora margarita is more intense in the subapical region of the lateral hyphae¹ (Fig. 1A). Furthermore, the correlation between the cytosolic pH profile previously obtained by Jolicoeur et al.,¹⁵ with the H⁺ efflux pattern (erythrosine-dependent), seems to clearly indicate that an active PM H⁺-ATPase takes place at the subapical hyphal region. Using orthovanadate, we could show that those H⁺ effluxes are susceptible mainly in the subapical region, but no effect in the apical was found.¹ Recently, a method to use fluorescent marker expression in an AM fungus driven by arbuscular mycorrhizal promoters was published.³¹ It could be adjusted as an alternative to measure “in vivo” PM H⁺-ATPase expression in AM fungal hyphae and their responses to root factors.³¹Open in a separate windowFigure 1(A) H⁺ flux profile along growing secondary hyphae of G. margarita in the presence (open squares) or absence (closed squares) of erythrosin B and its correlation with cytosolic pH (pHc) data described by Jolicoeur et al.,¹⁵ (dotted line). Dotted area depicts the region with higher susceptibility to erythrosin B. (B) ion-selective electrode near to AM fungal hyphae. (C) Stimulation on hyphal H⁺ efflux after incubation with root factors or whole root system. R, roots; RE, root exudates; CO₂, carbon dioxide; CWP, cell wall proteins; GR24, synthetic strigolactone. The medium pH in all treatment was monitored and remained about 5.7, including with prior CO₂ incubation. Means followed by the same letter are statistically equal by Duncan''s test at p < 5%.The H⁺ electrochemical gradient generated by PM H⁺-ATPases provides not only driving force for nutrient uptake,^9,16 but also can act as an intermediate in signal transduction pathways.¹⁸ The participation of these H⁺ pumps in cell polarity and tip growth of plant cells was recently reported,²⁷ addressing their crucial role on apical growth.²⁸ Naturally, in the absence of root factors the AM fungi have basal metabolic^8,21–23 and respiratory activity.²⁴ However when root signals are recognized and processed by AM fungal cells they might become activated.²² We thus searched for pH signatures that could reflect the alterations on fungal metabolism in response to external stimuli. In fact, preliminary analyses from our group demonstrate that AM fungal hyphae increase their H⁺ efflux in response not only to root exudates recognition, but also to other root factors (Fig. 1C). The incubation for 30 min of AM fungal hyphae with several root factors induces hyphal H⁺ efflux similar to the response to intact root system (5 days of incubation). The major increases were found with 1% CO₂ (750%) followed by root cell wall proteins (221%), root exudates (130%) and synthetic strigolactone (5%) (Fig. 1C). Those stimulations could define the transition from the state without root signals to the presymbiotic developmental stage (Fig. 1C). In the case of CO₂, the incorporation of additional carbon could represent a new source of energy, since CO₂ dark fixation takes place in Glomus intraradices germ tubes.^22,25Interestingly, after the treatment with synthetic strigolactone (10⁻⁵ M GR24), no significant stimulation was found compared to the remaining factors (Fig. 1C). It opens the question if the real effect of strigolactone is restrict to hyphal branching and does not intervene in very fast response pathways. Likewise, strigolactones need additional time to exhibit an effect, as recently discussed by Steinkellner et al.,²⁶ However, at the moment, no comprehensive electrophysiological analyses are presently available separating the effects of strigolactone and some flavonoids in AM fungal hyphae.The next target of our work is the study of ionic responses of single germ tubes or primary hyphae to root factors (Fig. 2). As reported by Ramos et al.,¹ we have been observing that the pattern of ion fluxes at the apical zone of primary hyphae is differentiated from secondary or lateral hyphae. In the primary, two interesting responses were detected in the absence of root factors: (1) a “dormant Ca²⁺ flux” and (2) Cl⁻ or anion fluxes at the same direction of H⁺ ions, suggesting a possible presence of H⁺/Cl⁻ symporters at the apex, similarly to what occurs in root hairs (Fig. 2).³⁰ In the presence of root factors such as root exudates the stimulated influxes of Cl⁻ (anion), H⁺, Na⁺ and effluxes of K⁺ and Ca²⁺ are activated. It can explain why the AM fungi hyphal tips are depolarized^20,29 during the period without root signals—“asymbiosis”—as long as K⁺ efflux and H⁺ influx occur simultaneously. Indeed, H⁺ as well as Ca²⁺ ions may act as second messengers, where extra and intracellular transient pH changes are preconditions for a number of processes, including gravity responses and possibly in plant-microbe interactions.^17,30Open in a separate windowFigure 2Ion dynamics in the apex of primary hyphae of arbuscular mycorrhizal fungi. It represents the Stage 1 described in Ramos et al.¹ After treatment with root factors, an activation of Ca²⁺ efflux is observed at the hyphal apex.Clearly, further data on the mechanism of action of signaling molecules such as strigolactones over the signal transduction and ion dynamics in AM fungi will be very important to improve our understanding of the molecular bases of the mycorrhization process. Future studies are necessary in order to provide basic knowledge of the ion signaling mechanisms and their role on the response of very important molecules playing at the early events of AM symbiosis.