Thigmomorphogenesis in Solanum lycopersicum: Morphological and biochemical responses in stem after mechanical stimulation

The activation of the phenylpropanoid pathway in plants by environmental stimuli is one of the most universal biochemical stress responses known. In tomato plant, rubbing applied to a young internode inhibit elongation of the rubbed internode and his neighboring one. These morphological changes were correlated with an increase in lignification enzyme activities, phenylalanine ammonia-lyase (PAL), cinnamyl alcohol dehydrogenase (CAD) and peroxidases (POD), 24 hours after rubbing of the forth internode. Furthermore, a decrease in indole-3-acetic acid (IAA) content was detected in the rubbed internode and the upper one. Taken together, our results suggest that decrease in rubbed internode length is a consequence of IAA oxidation, increases in enzyme activities (PAL, CAD and POD), and cell wall rigidification associated with induction of lignification process.Key words: Mechanical stimulation, PAL, CAD, POD, IAAIn their environment, plants are constantly submitted to several stimuli such as wind, rain and wounding. The growth response of plants to such stimuli was termed thigmomorphogenesis and was observed in a wide range of plants.^1–3 The most common thigmomorphogenetic response is a retardation of tissue elongation accompanied by an increase in thickness.⁴ The plant response to mechanical perturbation is mainly restricted to the young developing internode, since no influence can be detected when the internode has reached its final length.^5,6 These plant growth modifications, which characterize thigmomorphogenesis, are related to biochemical events associated with lignification process⁷ and ethylene production.^8,9In tomato plant the length of internodes 4 (N4) and 5 (N5) was measured 14 days after rubbing of the fourth internode. Results reported in Figure 1 show that rubbing led to a significant reduction of elongation of the stressed internode (N4) (decrease of N4 length from 4.3 cm in the control plant to 2.9 in the rubbed one). This effect was not limited to the rubbed area but affected also the elongation of the neighboring internodes (N5) that were shorter in rubbed plants than in control ones.Open in a separate windowFigure 1Internode lengths of control and rubbed plants measured 14 day after mechanical stress applied to the fourth internode. Standard errors are indicated by vertical bars.Results reported in Figure 2 show an increase in PAL activity in both internodes N4 and N5, 24 hours after mechanical stress application as compared with corresponding controls. CAD activity was also investigated in N4 and N5, 24 h after rubbing of the fourth internode. Results presented in Figure 3 show that mechanical stress application induces a strong increase of CAD activity in the rubbed internode N4 (5.3 nkatal μg-1 protein) with an approximately two-fold increase when compared to control tomato internodes (2.3 nkatal μg-1 protein). Further, CAD activity in N5 was also increased in the rubbed internode (5.538 nkatal μg-1 protein) as compared with the control one (3.256 nkatal μg-1 protein).Open in a separate windowFigure 2PAL activity of internode 4, and 5 in control and rubbed plants 24 h after rubbing of the fourth internode. Standard errors are indicated by vertical bars.Open in a separate windowFigure 3CAD activity of internode 4, and 5 in control and rubbed plants 24 h after rubbing of the fourth internode. Standard errors are indicated by vertical bars.Syringaldazine (S-POD) and gaïacol (G-POD) peroxidase activities were measured in tomato N4 and N5. Results reported in Figure 4 show an increase in soluble peroxidase activity with both substrates in the rubbed internode N4 as compared with control plant. Enhancement in peroxidase activities in N4 was more pronounced with gaïacol (80.7 U) as an electron donor than syringaldazine (33.8 U). Similar results were observed in internode 5 as compared with control one (Fig. 4).Open in a separate windowFigure 4(A) Syringaldazine-POD (Syr-POD) activity of internode 4 and 5 in control and rubbed plants 24 h after rubbing of the fourth internode. Standard errors are indicated by vertical bars. (B) Gaiacol-POD (G-POD) activity of internode 4 and 5 in control and rubbed plants 24 h after rubbing of the fourth internode. Standard errors are indicated by vertical bars.IAA was quantified in control and rubbed plant internodes 24 h after rubbing of the fourth internode. Results reported in figure 5 show that in control sample and as expected, the content of IAA was found to be higher in the younger internode (N5) as compared to the older one (N4). Rubbing led to a significant decrease in IAA levels in N4 (5.06 nmol g⁻¹ MF⁻¹) as compared with corresponding controls (7.27 nmol g⁻¹ MF⁻¹). Similar results were observed in internode 5, where IAA content was reduced from 16.52 nmol g⁻¹ MF⁻¹ in control internode to 12.35 nmol g⁻¹ MF⁻¹ in the rubbed internode (Fig. 5).Open in a separate windowFigure 5IAA Level of internode 4 and 5 in control and rubbed plants 24 h after rubbing of the fourth internode. Standard errors are indicated by vertical bars.The results reported here establish an evident correlation between growth limitation of the rubbed internode and their degree of lignification, the increase in lignification enzymes activities and auxin degradation after mechanical stress application.Auxin seems to be involved in thigmomorphogenesis.¹⁰ It was proposed that MIS (Mechanically-induced stress) has opposite effects on auxin levels in the two species studied to date, Phaseolus vulgaris¹⁰ and Bryonia dioica.^11,12 Auxin level as measured by bioassay, increased in Phaseolus vulgaris following rubbing of the stem.¹⁰ It was proposed that a build up of auxin may result from the reduced polar transport of IAA at the rubbed internode, causing a build up of IAA in the stem tissue. Exogenous IAA did not reverse the MIS inhibition of growth in Phaseolus vulgaris and high levels of IAA retarded growth in non-stressed plants.¹⁰ Thus, retardation of extension growth in Phaseolus vulgaris may have been caused by high levels of endogenous auxin and the increase in stem diameter by increased ethylene production.⁴ However, ethylene increases radial growth only if auxin is present.¹³Boyer¹¹ reported a decrease in auxinlike activity in Bryonia dioica following MIS and this was confirmed in the same species by Hofinger et al.¹² who reported a decrease in IAA using gas chromatography-mass spectrometry. Auxin catabolism was accompanied with changes in both soluble and ionically bound cell wall basic peroxidases¹⁴ and the appearance of an additional peroxidase. This can suggest that in Bryonia, auxin catabolism is hastened by mechanical stimulated peroxidase. In addition, Boyer et al.¹⁵ reported that lithium pre-treatment prevents both thigmomorphogenesis and appearance of specific cathodic isoperoxidase in Bryonia plants subjected to MIS. This is give further credence to the possibility that the peroxidase-auxin system is involved in Bryonia thigmomorphogenesis. In addition, ethylene increases peroxidase activity which reduces the auxin content in the tissue to a level low enough not to support normal growth. We have evidence that decrease of auxin level contribute to mechanism leading to tomato internode inhibition subjected to mechanical stress.Growth inhibition has been suggested to be the result of tissues lignification.⁶ As the initial enzyme in the monolignol biosynthesis pathway, PAL has a direct influence on lignin accumulation.¹⁶ The characteristics of lignin differ among cell wall tissues and plant organs.¹⁷ It comprises polyphenolic polymers derived from the oxidative polymerization of different monolignols, including p-coumaryl, coniferyl and sinapyl alcohols via a side pathway of phenylalanine metabolism leading to lignin synthesis.¹⁸ The increase in lignin content in the rubbed tomato internode could be a response mechanism to mechanical damage caused by rubbing.³ It is known that plants create a natural barrier that includes lignin and suberin synthesis, components directly linked to support systems.^19,20The increase in lignin content of rubbed tomato internode³ is paralleled by a rise in CAD activity and whilst such direct proportionality between CAD activity and lignin accumulation does not always agree with the results in the literature, it clearly is responding in ways similar to those of the other enzymes in the pathway.²¹Mechanical stress-induced membrane depolarization would generate different species of free radicals and peroxides, which in turn initiate lipid peroxidation.²² The degradation of cell membranes is suggested to bring about rapid changes in ionic flux, especially release of K⁺ which would result in an enhanced endogenous Ca/K ratio and in leakage of solutes, among them electron donors such as ascorbic acid and phenolic substances. The increased intracellular relative calcium level activated secretion of basic peroxidases²³ into the free space where, in association with the electron donors and may be with the circulating IAA, they eliminate the peroxides, and facilitated binding of basic peroxidases to membrane structures allowing a role as 1-aminocyclopropane-1-carboxylic acid (ACC)-oxidases. The resulting IAA and ACC oxidase-mediated changes in ethylene production²⁴ would further induce (this time through the protein synthesis machinery) an increase in activity of phenylalanine ammonia-lyase and peroxidases. The resulting lignification and cell wall rigidification determines the growth response of tomato internode to the mechanical stress.     

Dynamic protein trafficking to the cell wall

Recently we have studied the secretion pattern of a pectin methylesterase inhibitor protein (PMEI1) and a polygalacturonase inhibitor protein (PGIP2) in tobacco protoplast using the protein fusions, secGFP-PMEI1 and PGIP2-GFP. Both chimeras reach the cell wall by passing through the endomembrane system but using distinct mechanisms and through a pathway distinguishable from the default sorting of a secreted GFP. After reaching the apoplast, sec-GFP-PMEI1 is stably accumulated in the cell wall, while PGIP2-GFP undergoes endocytic trafficking. Here we describe the final localization of PGIP2-GFP in the vacuole, evidenced by co-localization with the marker Aleu-RFP, and show a graphic elaboration of its sorting pattern. A working model taking into consideration the presence of a regulated apoplast-targeted secretion pathway is proposed.Key words: cell wall trafficking, endocytosis, GPI-anchor, PGIP2, PMEI1, secretion pathway, vacuole fluorescent markerCell wall biogenesis, growth, differentiation and remodeling, as well as wall-related signaling and defense responses depend on the functionality of the secretory pathway. Matrix polysaccharides, synthesized in the Golgi stacks, and cell wall proteins, synthesized in the ER, are packaged into secretory vesicles that fuse with the plasma membrane (PM) releasing their cargo into the cell wall. Also the synthesis and deposition of cellulose itself are driven by the endomembrane system which controls the assembly, within the Golgi, and the export to the plasma membrane of rosette complexes of cellulose synthase.¹ Secretion to the cell wall has always been considered a default pathway² but recent studies have evidenced a complex regulation of wall component trafficking that does not seem to follow the default secretion model. Recent evidence that several cell wall proteins are retained in the Golgi stacks until specific signals at the N-terminal domain are proteolitically removed is a case in point.^3–5 Moreover, it has previously been reported that secretion of exogenous marker proteins (secGFP and secRGUS) and cell wall polysaccharides reach the PM through different pathways.⁶ More recently, we have reported that cell wall protein trafficking also occurs through mechanisms distinguishable from that of a secreted GFP suggesting that more complex events than the mechanisms of bulk flow control cell wall growth and differentiation.⁷ To follow cell wall protein trafficking we used a Phaseolus vulgaris polygalacturonase inhibitor protein (PGIP2) and an Arabidopsis pectin methylesterase inhibitor protein (PMEI1) fused to GFP (PGIP2-GFP and secGFP-PMEI1). Both apoplastic proteins are involved in the remodeling of pectin network with different mechanisms. PGIP2 specifically inhibits exogenous fungal polygalacturonases (PGs) and is involved in the plant defense mechanisms against pathogenic fungi.^8,9 PMEI1 counteracts endogenous PME and takes part in the physiological synthesis and remodeling of the cell wall during growth and differentiation.^10,11 The specific functions of the two apoplastic proteins seem to be strictly related to the distinct mechanisms that control their secretion and stability in the cell wall. In fact, while secGFP-PMEI1 moves through ER and Golgi stacks linked to a glycosyl phosphatidylinositol (GPI)-anchor, PGIP2-GFP moves as a cargo soluble protein. Furthermore, secGFP-PMEI1 is stably accumulated in the cell wall, while PGIP2-GFP, over the time, is internalized into endosomes and targeted to vacuole, likely for degradation. After reaching the cell wall, the different fate of the two proteins seems to be strictly related to the presence/absence of their physiological counteractors. PMEI regulates the demethylesterification of homogalacturonan by inhibiting pectin methyl esterase (PME) activity through the formation of a reversible 1:1 complex which is stable in the acidic cell wall environment.¹² Stable wall localization of PMEI1 is likely related to its interaction with endogenous PME, always present in the wall. Unlike PMEs, fungal polygalacturonases (PGs), the physiological interactors of PGIP2, are present in the cell wall only during a pathogen attack. The absence of PGs may determine PGIP2 internalization. Internalization events have been already reported for PM proteins,^13–16 while cell wall protein internalization is surely a less well-known event. To date, only internalization of an Arabidopsis pollen-specific PME^4,5,17 and PGIP2 ⁷ has been reported.To further confirm the internalization of PGIP2-GFP and its final localization into the vacuole, we constructed a red fluorescent variant (RFP) of the green fluorescent marker protein that accumulates in lytic or acidic vacuole because of the barley aleurain sorting determinants (Aleu-RFP).¹⁸ The localization of PGIP2-GFP was compared to that of Aleu-RFP by confocal microscopy in tobacco protoplasts transiently expressing both fusions. Sixty hours after transformation, PGIP2-GFP labeled the central vacuole as indicated by complete co-localization with the vacuolar marker (Fig. 1A–D). Instead, at the same time point, secGFP-PMEI1 still labeled the cell wall (Fig. 1E–H) and never reached the vacuolar compartment. To summarize PGIP2-GFP secretion pattern, a graphic elaboration of confocal images is reported describing the sorting of PGIP2GFP in tobacco protoplast (Fig. 1I). The protein transits through the endomembrane system (green) and reaches the cell wall which is rapidly regenerating as evidenced by immunostaining with the red monoclonal antibody JIM7 that binds to methylesterified pectins.¹⁹ PGIP2-GFP is then internalized in endosomes, labeled in yellow because of the co-localization with the styryl dye FM4-64, a red marker of the endocytic pathway.Open in a separate windowFigure 1PGIP2-GFP, but not secGFP-PMEI1, is internalized and reaches the vacuole in tobacco leaf protoplasts. (A) Approximately 60 h after transformation, PGIP2-GFP labeled the central vacuole as indicated by co-localization with the vacuole marker Aleu-RFP (B). (C) Merged image of (A and B). (D) Differential interference contrast (DIC) image of (A–C). On the contrary, secGFP-PMEI1 still labeled cell wall (E). (F) No co-localization is present in the vacuole labeled by Aleu-RFP. (G) Merged image of (E and F). (H) DIC image of (E–G). (I) Graphic elaboration of confocal images describing the sorting of PGIP2. The protein is sorted by the endomembrane system (green) to the cell wall (red) that is regenerated by the protoplast. Lacking the specific ligand, it is then internalized in endosome (yellow). Details are reported in the text.In Figure 2 we propose a model of the mechanism of secGFP-PMEI1 and PGIP2-GFP secretion derived from the different lines of evidence previously reported in reference ⁷. SecGFPPMEI1 (Fig. 2-1), but not PGIP2-GFP (Fig. 2-2), carries a GPI-anchor, required for its secretion to the cell wall. When the anchorage of GPI is inhibited by mannosamine (Fig. 2-a) or by the fusion of GFP to the C-terminus of PMEI1 (Fig. 2-b), the two non-anchored proteins accumulate in the Golgi stacks. Evidence of retention in Golgi stacks has already been reported for other two cell wall proteins.^3–5 Unlike secGFP-PMEI1, PGIP2-GFP is not stably accumulated in the cell wall and undergoes endocytic trafficking (Fig. 2-3). PGIP2-GFP internalization, likely due to the absence of PGs, might also be related with its ability to interact with homogalacturonan and oligogalacturonides,²⁰ which have been reported to internalize^21,22 (Fig. 2-4). Since SYP 121, a Qa-SNARE, is involved in the default secretion of secGFP,²³ but not in secretion of PGIP2-GFP and secGFP-PMEI1, trafficking mechanisms underlying secretion into the apoplast are likely different from those underlying the default route (Figs. 2-5). Taken as a whole, evidence suggests the existence of currently undefined signals that control apoplast-targeted secretion.Open in a separate windowFigure 2Schematic illustration for secGFP-PMEI1 and PGIP2-GFP trafficking. See text for details.     

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.     

High-throughput transient transformation of Arabidopsis roots enables systematic colocalization analysis of GFP-tagged proteins

Determination of the subcellular localization of an unknown protein is a major step towards the elucidation of its function. Lately, the expression of proteins fused to fluorescent markers has been very popular and many approaches have been proposed to express these proteins. Stable transformation using Agrobacterium tumefaciens generates stable lines for downstream experiments, but is time-consuming. If only colocalization is required, transient techniques save time and effort. Several methods for transient assays have been described including protoplast transfection, biolistic bombardment, Agrobacterium tumefaciens cocultivation and infiltration. In general colocalizations are preferentially performed in intact tissues of the same species, resembling the native situation. High transformation rates were described for cotyledons of Arabidopsis, but never for roots. Here we report that it is possible to transform Arabidopsis root epidermal cells with an efficiency that is sufficient for colocalization purposes.Key words: Arabidopsis, GFP-fusions, protein localization, root, transient transformationSince the release of the Arabidopsis thaliana genome sequence plant biologists set the goal to elucidate the functions of all coded genes. Apart from the spatio-temporal expression patterns of genes, the subcellular localization of gene products can play an essential role in deciphering their function. Classical immunological approaches to localize proteins can be hindered by cross-reactivity, time-consuming generation of antibodies and the low temporal resolution. Expression of tagged proteins forms a suitable alternative. Lately, fusions with fluorescent proteins in combination with confocal (CLSM)¹ or spinning disc microscopy² allow real time protein localization and even subcellular trafficking at high resolution. An overview of fluorescent tagging approaches can be found elsewhere.³Currently several techniques to introduce the coding region for a tagged protein in a plant are available. The generation of stable lines transformed by Agrobacterium tumefaciens offers a continuous source of plant material, but it is time-consuming especially when only colocalization experiments are required. Transient assays, on the other hand, offer the advantage of being fast and amenable to high throughput strategies. Each of these techniques, however, has some limitations and drawbacks. Particle bombardment (biolistics) ^4–6 for example circumvents the host specificity of Agrobacterium strains, but requires expensive equipment. Moreover, it is rather disruptive and imposes a significant stress upon the plants, possibly influencing the results. Protoplasts lack a cell wall and protoplast transformation^7,8 is therefore not suitable for certain experiments related to cell wall proteins or when interactions between cells on tissue level might be important.⁹ Moreover, protoplasts have lost their identity which might be critical for the correct functioning of certain transgenic constructs. Agrobacterium infiltration of tobacco leaves¹⁰ is regularly used and represents an efficient, fast and relatively easy transformation technique. However, tobacco leaves easily show autofluoresence due to tissue damage as a result of experimental manipulations. As it has been reported that some protein fusions expressed in an heterologous system localize to different subcellular localizations11 it is advisable not to use tobacco when localizing Arabidopsis proteins. Leaf infiltrations have been performed in Arabidopsis,¹² but apparently their leaves are much more prone to mechanical damage and the leaf developmental stage is critical, complicating this technique. Cocultivation of Agrobacterium with seedlings offers a rapid and efficient approach applicable to many mono and dicot species. It was reported to work efficiently in Arabidopsis cotyledons, but not in roots.⁹ As an alternative method, Agrobacterium infiltration of Arabidopsis seedlings¹¹ seems an efficient technique for transient expression. However, expression in root cells could not be obtained. Colocalizations are required in the native cells or tissue for the correct localization of an unknown protein or proteins that need interaction partners. As a consequence this technique can not be reliably used when root expressed gene products are studied. Here we show evidence that it is possible to use the described technique¹¹ to induce transient expression in Arabidopsis roots.We used the Agrobacterium infiltration of Arabidopsis seedlings technique¹¹ to colocalize several C-terminal (S65T)-sGFP fusions generated in the plant binary vector pGWB6.¹³ Each construct was transformed into Agrobacterium tumefaciens (C59C1Rif^R) containing the helper plasmid pMP90. Subsequently different stable marker lines, wild type Arabidopsis (Col-0) bearing mCherry fusion constructs,¹⁴ were transiently transformed.¹¹ After 2 or 3 days seedlings were studied using CLSM. Besides being expressed in cotyledons fusion proteins were clearly observed in root epidermis and root cap cells (Fig. 1A and B). As reported¹¹ the transformation efficiency in cotyledons was considerably higher than in root cells. However, in each experiment we obtained a considerable amount of transformed root epidermal cells which was more than sufficient for colocalization studies (Fig. 2). It was remarkable that transformation was repeatedly successful in groups of cells, adjacent or close to each other.Open in a separate windowFigure 1Transient transformation of Arabidopsis root cells. Expression of the protein-GFP fusion product can be seen in the epidermal (A) and root cap cells (B) on fluorescence/transmission merged images. As seen in (A) high efficiencies of root transformation can be reached.Open in a separate windowFigure 2Colocalization of mCherry and GFP constructs. Confocal image of the mCherry fluorescence (A), the GFP signal (B) and the merged image (C).In contrast to what was reported earlier we show here that the Agrobacterium infiltration technique¹¹ is perfectly capable of transiently transforming Arabidopsis root epidermal cells. It allows the transient production and study of proteins in their native environment, considerably increasing the reliability of such experiments. Additionaly the use of RFP marker constructs in colocalisation studies in the root is free of interference by the red background autofluorescence of chlorophyll.     

A new function for odorant receptors: MOR23 is necessary for normal tissue repair in skeletal muscle

Myofibers with an abnormal branching cytoarchitecture are commonly found in various neuromuscular diseases as well as after severe muscle injury. These aberrant myofibers are fragile and muscles containing a high percentage of these myofibers are weaker and more prone to injury. To date the mechanisms and molecules regulating myofiber branching have been obscure. Recent work analyzing the role of mouse odorant receptor 23 (MOR23) in muscle regeneration revealed that MOR23 is necessary for proper skeletal muscle regeneration in mice as loss of MOR23 leads to increased myofiber branching. Further studies demonstrated that MOR23 expression is induced when muscle cells were extensively fusing and plays an important role in controlling cell migration and adhesion. These data demonstrate a novel role for an odorant receptor in tissue repair and identify the first molecule with a functional role in myofiber branching.Key words: muscle regeneration, odorant receptor, olfactory receptor, MOR23, myofiber splitting, myofiber branching, myoblast, fusion, myotube, olfr16Skeletal muscle is characterized by an extensive ability to regenerate after injury due to trauma or disease. Muscle regeneration results from a finely orchestrated series of steps that are spatially and temporally regulated, many of which are not understood. Elucidation of the mechanisms that regulate muscle regeneration may be beneficial for enhancing the rate or extent of muscle regeneration in injury, disease or age.Skeletal muscle is composed of myofibers, which are long cylindrical cells containing hundreds of myonuclei in a common cytoplasm (Fig. 1). Each myofiber is surrounded by a basal lamina sheath; between the myofiber and the basal lamina lie myogenic stem cells called satellite cells. When muscle is injured, myofibers degenerate and satellite cells proliferate to give rise to progeny myoblasts. Myoblasts differentiate and undergo migration, adhesion and fusion to form regenerated myofibers and normal tissue architecture is restored. In many neuromuscular diseases muscle regeneration is aberrant and various abnormalities, such as variation in myofiber size, decreased myofiber number, fibrosis and branched myofibers, are observed. In the clinical literature, branched myofibers are more commonly referred to as “split myofibers.”Open in a separate windowFigure 1Myofiber growth during normal muscle regeneration. (A) Myofibers contain many myonuclei within a common cytoplasm and are surrounded by a basal lamina sheath. Underneath the basal lamina lie satellite cells, myogenic stem cells responsible for muscle regeneration. (B) Myofiber degeneration leads to activation of quiescent satellite cells and their reentry into the cell cycle. Their progeny myoblasts proliferate to yield a pool of progenitor cells. (C) Myoblasts differentiate and undergo migration, adhesion and fusion to form nascent myofibers within the original basal lamina sheath. Additional myoblasts fuse with these newly formed myofibers and the myofiber will continue to grow in size. (D) At later time points regenerated myofibers are similar in size to undamaged myofibers but contain centrally located myonuclei, a hallmark of a regenerated myofiber.Branched myofibers are malformed cells which, instead of having a normal cylindrical shape, contain one or more offshoots of small daughter myotubes contiguous with the parent myofiber (Fig. 2). Branched myofibers can be simple with only one branch (Fig. 2A) or complex with many anastomosing branches resembling a gnarled tree root (Fig. 2B).¹ In myofibers with complex cytoarchitecture, individual branches can persist up to hundreds of microns and then eventually recombine with the parent myofiber. Each daughter branch is enclosed in its own basal lamina, which is contiguous with the basal lamina of the parent myofiber.² The frequency of branched myofibers in rodent muscle under normal conditions is low, on the order of 0.003%.³ However, the frequency in both rodent and human muscle is increased in response to hypertrophy^4,5 as well as regeneration due to induced injury,^6–8 muscle transplantation,^9,10 or muscular dystrophy.^1,11–16 In mdx mice, a model of Duchenne Muscular Dystrophy, up to 65–90% of myofibers by 7 months and older display this abnormal branched morphology compared to 6–17% of myofibers at 1–3 months of age.^12–14 Branched myofibers display functional abnormalities such as alterations in myofiber calcium signaling.¹³ Additionally, isolated branched myofibers are more prone to rupture at branches during stimulation¹⁷ and mdx muscles containing a high percentage of branched myofibers are more vulnerable to contraction-induced injury.^12,14 Thus, decreasing the number of branched myofibers would likely be beneficial to muscle function.Open in a separate windowFigure 2Myofiber branching during aberrant muscle regeneration. (A) Phase contrast microscopy of a normal (left) and a branched (right) myofiber. The branched myofiber contains one branch at the end of the myofiber. (B) Schematic diagrams of myofibers with more complex patterns of branching than depicted in (A).Although branched myofibers have been reported in literature for over 100 years, the mechanism by which they arise is unknown and no causative molecules have been identified. The aberrant cytoarchitecture of branched myofibers likely arises from incomplete fusion of small myotubes during muscle regeneration⁸ though direct proof is lacking. Evidence in favor of these malformed myofibers arising from abortive regenerative processes includes the expression of neonatal myosin, a marker of early muscle regeneration, in the small branches.¹³ That these branches are newly formed is further suggested by the presence of centrally located nuclei,^13,18 a hallmark of regenerated myofibers. The observation that during muscle regeneration multiple small myotubes can form within the old basal lamina sheath^8,16,19 leads credence to the idea that aberrant fusion of such small myotubes underlies generation of branched myofibers. Indeed, electron microscopic studies support the ability of myotubes to fuse with one another in vivo;²⁰ myotubes readily fuse with one another in vitro.²¹ Recent studies of odorant receptor function during muscle regeneration in mice¹⁸ have identified the first molecule with a functional role in controlling myofiber branching and suggest that defects in muscle cell migration and/or adhesion may underlie the genesis of these branches.     

The Twisted Dwarf's ABC: How Immunophilins Regulate Auxin Transport

There is increasing evidence that immunophilins function as key regulators of plant development. One of the best investigated members, the multi-domain FKBP TWISTED DWARF1 (TWD1)/FKBP42, has been shown to reside on both the vacuolar and plasma membranes where it interacts in mirror image with two pairs of ABC transporters, MRP1/ MRP2 and PGP1/PGP19(MDR1), respectively. Twisted dwarf1 and pgp1/pgp19 mutants display strongly overlapping phenotypes, including reduction and disorientation of growth, suggesting functional interaction.In a recent work using plant and heterologous expression systems, TWD1 has been demonstrated to modulate PGP-mediated export of the plant hormone auxin, which controls virtually all plant developmental processes. Here we summarize recent molecular models on TWD1 function in plant development and PGP-mediated auxin tranport and discuss open questions.Key Words: Twisted Dwarf1, plant development, auxin, immunophilin, P-glycoprotein, ABC transporterFK506-binding Proteins (FKBPs), together with unrelated cyclophilins, belong to the immunophilins, an ancient and ubiquitous protein family.^1,4,5 They were first described as receptors for immunosuppressive drugs in animal and human cells, FK506 and cyclosporin A, respectively.¹ All FKBP-type immunophilins share a characteristic peptidyl-prolyl cis-trans isomerase domain (PPIase domain or FKBD, Fig. 2A) making protein folding a key feature among immunophilins.² The best investigated example, the human cytosolic single-domain FKBP12, modulates Ca²⁺ release channels^6,7 and associates with the cell cycle regulator TGF-β.⁸ Furthermore, the human FKBP12/FK506 complex is known to bind and inhibit calcineurin activity,⁹ leading to immune response inhibition. However, not all single- and multiple-domain FKBPs own folding activity and, interestingly, many form distinct protein complexes with diverse functions.^3–5Open in a separate windowFigure 2Model of TWISTED DWARF 1 interacting proteins. (A) Domain structure of TWD1 and putative interacting proteins. FKBD, FK506-binding domain: TPR, tetratricopeptide repeat; CaM(-BD, calmodulin-binding domain; MA, membrane anchor. For details, see text. (B) Functional TWD1-ABC transporter complexes on both the vacuolar and plasma membrane. While for TWD1/PGP pairs, the positive regulatory role on auxin transport was demonstrated,¹⁸ the modulation of MRP-mediated vacuolar import of glutathion conjugates (GS-X) was established using mammalian test substrates¹⁷ because the in vivo substrates are unknown. Note that C-terminal nucleotide binding folds of MRP- and PGP-like ABC transporters interact with distinct functional domains of TWD1, the TPR and FKBD, respectively. The native auxin, IAAH, gets trapped by deprotonization upon uptake into the cell. Export is catalyzed by secondary active export via PIN-like efflux carriers¹⁵ and/or by primary active, ATP-driven P-glycoproteins (PGPs, right panel); loss-of TWD1 function abolishes PGP-mediated auxin export (left panel).     

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.     

Membrane localization diversity of TPK channels and their physiological role

Potassium (K) is one of the major nutrients that is essential for plant growth and development. The majority of cellular K⁺ resides in the vacuole and tonoplast K⁺ channels of the TPK (Two Pore K) family are main players in cellular K⁺ homeostasis. All TPK channels were previously reported to be expressed in the tonoplast of the large central lytic vacuole (LV) except for one isoform in Arabidopsis that resides in the plasma membrane. However, plant cells often contain more than one type of vacuole that coexist in the same cell. We recently showed that two TPK isoforms (OsTPKa and OsTPKb) from Oryza sativa localize to different vacuoles with OsTPKa predominantly found in the LV tonoplast and OsTPKb primarily in smaller compartments that resemble small vacuoles (SVs). Our study further revealed that it is the C-terminal domain that determines differential targeting of OsTPKa and OsTPKb. Three C-terminal amino acids were particularly relevant for targeting TPKs to their respective endomembranes. In this addendum we further evaluate how the different localization of TPKa and TPKb impact on their physiological role and how TPKs provide a potential tool to study the physiology of different types of vacuole.Key words: TPK channels, small vacuoles, vacuolar targeting, potassiumThe roles of plant vacuolar K⁺ channels are diverse and include potassium homeostasis, turgor regulation and responses to abiotic stress. Vacuolar K⁺-selective channels belong to two-pore K⁺ (TPK) channel families which have been found in genomes of many plant species such as Arabidopsis, poplar, Physcomitrella, Eucalyptus, barley, potato, rice and tobacco (Fig. 1). TPKs have structural similarity to mammalian “tandem P domain” channels with a secondary structure that contains four transmembrane domains and two pore regions (Fig. 2).^1–5 TPK channels have pore regions with a GYGD signature that endows K⁺ selectivity and a variable number of Ca²⁺ binding EF domains in the C terminus.^3–8 One of the best characterized members of the TPK family is AtTPK1 from Arabidopsis thaliana. AtTPK1 activity is voltage independent but sensitive to cytosolic Ca²⁺, cytosolic pH and N-terminal phosphorylation by 14-3-3 proteins.^5,6,8,9 In Arabidopsis, AtTPK1 expresses in the large lytic vacuole (LV) and plays roles in cellular K⁺ homeostasis, K⁺-release during stomatal closure and seed germination.^4,5 Other members of the Arabidopsis TPK family (AtTPK2, AtTPK3, AtTPK5) have been shown to localize to the LV but also showed some expression in smaller, vesicle-like, compartments.⁴ However, none of these isoforms appears to form functional channels in planta although our experiments with heterologous expression of AtTPK3 and AtTPK5 in the K⁺ uptake deficient E. coli LB2003 demonstrates complementation of bacterial growth phenotype (Isayenkov S, et al. unpublished results). Equally intriguing, is the plasma membrane localization of the Arabidopsis TPK4 isoform, in spite of its sequence being very similar to that of other TPKs.¹⁰Open in a separate windowFigure 1Phylogenetic tree of plant TPKs. The three main clusters of TPKs comprise: Cluster 1 with AtTPK1-like channels; Cluster 2 with AtTPK3/TPK5-like channels; Cluster 3 with barley HvTPKb. Bootstrap analysis was performed using ‘Molecular Evolutionary Genetics Analysis, MEGA4’ software available at www.megasoftware.net/mega4/megaOpen in a separate windowFigure 2Two-pore potassium channel secondary structure. TPK channels comprise four transmembrane domains (1–4) and two pore regions (P) per subunit. Functional channels are formed from two subunits. In most TPKs, both P regions contain a K⁺ selectivity signature, GYGD. However, the tobacco NtTPKa isoform has different motifs in the second P domain. In the N terminal region, TPKs have a 14-3-3 binding domain that impact on channel activity, with the binding of 14-3-3 protein leading to channel activation. C-termini of TPKs show a varying number of putative Ca²⁺ binding “EF hands” which may vary from zero to two.     

Auxin distribution and lenticel formation in determinate nodule of Lotus japonicus

Legumes can establish a symbiosis with rhizobia and form root nodules that function as an apparatus for nitrogen fixation. Nodule development is regulated by several phytohormones including auxin. Although accumulation of auxin is necessary to initiate the nodulation of indeterminate nodules, the functions of auxin on the nodulation of determinate nodules have been less characterized. In this study, the functions of auxin in nodule development in Lotus japonicus have been demonstrated using an auxin responsive promoter and auxin inhibitors. We found that the lenticel formation on the nodule surface was sensitive to the auxin defect. Further analysis indicated that failure in the development of the vascular bundle of the determinate nodule, which was regulated by auxin, was the cause of the disappearance of lenticels.Key words: auxin, lenticel, Lotus japonicus, nodulation, symbiotic nitrogen fixationLegumes (Fabaceae) constitute the third largest plant family with around 700 genera and 20,000 species.¹ Legume plants form root nodules through symbiosis with a soil microbe called rhizobia. This plant-microbe symbiosis in nodules mediates an harmonized exchange of chemical signals between host plants and rhizobia.² Nodules are biologically divided into two different groups, i.e., indeterminate nodules and determinate nodules. Indeterminate nodules, represented by Trifolium repens (white clover) and Medicago truncatula, are initiated from the inner cortex to form a persistent nodule meristem, which allows continuous growth, and leads to the formation of elongated nodules, whereas in determinate legumes, nodules are mostly developed from outer cortical cells and form spherical nodules.³Auxin is one of the most important regulators for nodule development. Since the possible involvement of auxin in nodule formation was first reported by Thimann,⁴ auxin distribution during nodulation has been studied in particular with indeterminate nodules.⁵ However, little is known about auxin involvement in determinate nodule formation. To evaluate auxin functions in the determinate nodulation of legume plants, we performed an auxin-responsive promoter analysis in detail. Using GH3:GUS transformed Lotus japonicus (a kind gift from Dr. Herman P. Spaink, Leiden State University, Netherlands),⁶ we detected auxin signals throughout the nodulation process, e.g., at the basal and front part of the nodule primordia, circumjacent to the infection zone of the young developing nodules (Fig. 1), and at the nodule vascular bundle in mature nodules. We also investigated the effect of several auxin inhibitors, including newly synthesized auxin antagonist PEO-IAA (kindly provided by Dr. Hayashi, Okayama University of Science, Japan),⁷ on the nodulation of L. japonicus, and revealed that auxin was required for forming a nodule vascular bundle and lenticels (Fig. 2).⁸Open in a separate windowFigure 1GH3:GUS expression in determinate nodule at 6 dpi. (A) GUS staining was observed in the central cylinder of the root vascular bundle and in the nodule. (B) Cross section of (A). GUS expression was observed around the infection zone of the nodule. Bars = 100 µm.Open in a separate windowFigure 2The effect of auxin inhibitor on nodule surface. (A) Typical mature nodule of L. japonicus at 21 dpi. Lenticels are pointed out by yellow arrowheads. (B) The treatment of auxin inhibitor (NPA 100 µM) inhibited lenticel formation on the nodule surface. Bars = 500 µm.In indeterminate legumes, auxin is accumulated at the site of rhizobia inoculation.⁹ This is caused by the inhibition of polar auxin transport by accumulation of flavonoids around the infection site, which are known as regulators of auxin transport. When flavonoid biosynthesis is reduced by the gene silencing of chalcone synthase, which catalyzes the first step of flavonoid synthesis, M. truncatula was unable to inhibit polar auxin transport and resulted in reduced nodule number.^10,11 A similar phenotype was observed when the auxin transporter gene was silenced.¹² In addition, treatment of polar auxin transport inhibitors such as NPA and TIBA induce pseudonodule formation,⁹ suggesting that auxin accumulation is required for nodulation of indeterminate legumes. In contrast, the treatment of polar auxin transport inhibitors in determinate nodules did not induce a nodule-like structure, suggesting a different function of auxin between indeterminate and determinate nodules. It is, however, of interest to investigate the involvement of flavonoids in determinate nodule formation, because several genes in the flavonoid biosynthesis pathway are upregulated at 2 dpi (days post inoculation) in L. japonicus.¹³Lenticels regulate gas permeability of nodules.¹⁴ Under low oxygen or water-logged conditions, they develop more extensively, whereas they collapse, or develop very little during insufficient water conditions, or under high oxygen pressure.^14,15 Because lenticel development on the nodule surface is accompanied with the nodule vascular bundle, growth regulators supplied from the vascular system likely facilitate lenticel development.¹⁵ Our data suggests that auxin is necessary to form the nodule vascular bundle, and in fact, auxin itself is one of the candidates of growth substances that control lenticel formation. It is necessary to analyze mutants, which lack in lenticel formation, but can form a nodule vascular bundle, for clarification of further mechanisms of lenticel development.     

Multiple signaling pathways control nitrogen-mediated root elongation in maize

Response of root system architecture to nutrient availability is an essential way for plants to adapt to soil environments. Nitrogen can affect root development either as a result of changes in the external concentration, or through changes in the internal nutrient status of the plant. Low soil N stimulates root elongation in maize. Recent evidence suggests that plant hormones auxin and cytokinin, as well as NO signaling pathway, are involved in the regulation of root elongation by low nitrogen nutrition.Key words: nitrogen, root growth, auxin, cytokinin, NONitrogen acquisition is determined by N demand for plant growth. At low N stress, N demand for maximum plant growth rate is not matched by plant N uptake. To acquire adequate N, plants may increase root length density to explore a larger soil volume and/or increase N uptake activity. High root density is also an important root trait for competition with soil microorganisms.¹ Since nitrate is a highly mobile, non-adsorbing ion, theoretic analysis predicts that its uptake is not limited by transport through soil, and a small root system is sufficient for nitrate acquisition.^2–4 In field conditions, however, genotypes that are efficient in N acquisition generally had a larger root system and higher root length density.^5,6 Under conditions of insufficient N supply, N mass flow to roots may not be adequate to meet the N demand for plant growth. Even in N-sufficient soils, various soil constraints (low water content, etc) may reduce the N mass flow rate. In these cases, large root size and high density will be very important for the utilization of the spatially distributed N, especially newly mineralized N, and the competition for organic N with soil microorganisms.^7,8The development of lateral roots in Arabidopsis in response to nitrate supply has been widely studied.⁹ Less attention has been paid to primary root growth in response to N, possibly because root elongtion is insensitive to increased N supply in Arabidopsis.^10,11 In maize, however, root elongation was sigificantly promoted by suboptimal N supply, and inhibited by overdose supply of N (Fig. 1).^12,13 Until recently less is known about the underlying physiological mechansms. It is well documented that cytokinin is a root-to-shoot signal communicating N availability in addition to nitrate itself.¹⁴ Exogenous cytokinin application suppresses the elongation of primary roots.¹⁵ Recent work in Arabidopsis overexpressing cytokinin synthase (IPT) demonstrate that long-term CK overproduction inhibited primary root elongation by reducing quantitative parameters of primary root meristem.¹⁶ By comparing two maize inbred lines whose root elongation had a differential response to low N stress, it was found that the change of cytokinin content in roots was closely related to low-N induced root elongation.¹³ In the N-sensitive genotype 478, cytokinin (Zeatin + Zeatin riboside) content was significantly lower at low N condition. While in N-insensitive genotype Wu312, cytokinin content was hardly affected at various N supplies. Higher N supply shortened the distance from root apex to the first visible lateral roots, a phenomenen similar to that caused by exogenous cytokinins. Furthermore, exogenous cytokinin 6-benzylaminopurine (6-BA) completely reversed the stimulatory effect of low nitrate on root elongation. All the data suggests that the inhibitory effect of high concentration of nitrate on root elongation is, at least in part, mediated by increased cytokinin level in roots.Open in a separate windowFigure 1Root elongation is inhibited at high nitrate supply.Auxin regulates many cellular responses crucial for plant development. Auxin plays a key role in establishing and elaborating patterns in root meristems.^17,18 Root elongation of Arabidopsis is enhanced by exogenous auxin at low concentrations, but is inhibited at high concentrations.¹⁹ In an earlier report, a high external nitrate supply (8 mM) did cause a 70% decrease in the auxin concentration of the root in soybean.²⁰ In maize, inhibition of root growth by high nitrate was found closely related to the reduction of IAA levels in roots and exogenous NAA and IAA restored primary root growth in high nitrate concentrations.²¹ Interesting, it was found that auxin concentrations in phloem exudates were reduced by a greater nitrate supply, suggesting that shoot-to-root auxin transport may be inhibited by high N supply. Considering the antagonism between auxin and cytokinin.²² it was possible that, by increasing the cytokinin level and decreasing the auxin level, high nitrate supply may have negative influences on root apex activity so that root apical dominance is weakened and, therefore, root elongation is suppressed and lateral roots grow closer to the root apex.Nitric oxide (NO) is emerging as an important messenger molecule associated with many biochemical and physiological processes in plants. The involvement of NO in IAA-induced adventitious root development has also been reported.²³ Given that nitrate is a substrate for NR-catalysed NO production, and root development and growth are closely related to NO, it is expected that NO may play a role in nitrate-dependent root growth. Surprisingly, endogenous levels of NO in the root apices of maize seedlings grown in high nitrate solution were much lower than those in apices grown in low nitrate. The nitrate-induced inhibition of root elongation in maize was markedly reversed by treatments of the roots with a NO donor (SNP) and IAA.²⁴ These data suggest that the arrest of root elongation by high levels of external nitrate concentrations may result from an alteration of endogenous NO levels in root apical cells. NR mediated NO production is unlikely to be involved in the nitrate-dependent NO production and root elongation because NR activity is lower at low N supply. A NO synthase (NOS) inhibitor reduced root elongation in maize plants grown in the low-nitrate medium, suggest that NOS activity may be inhibited in plants grown in high-nitrate solution, thus leading to a reduction of the endogenous NO levels.Taken together, high nitrogen supply increases cytokinin level, but decreases auxin and NO levels in roots of maize. Besides, it was well documented ethylene has a negative effect on root elongation of various plants.^25–27 Exogenous supply of cytokinin increase ethylene production (Stenlid 1982; Bertell et al., 1990). Recently, it was demonstrated in Arabidopsis that auxin transport from the root apex via the lateral root cap is required for ethylene-mediated inhibition of root growth.²⁸ Therefore, a complex multiple siganlling pathways may be involved in N-mediated root elongation (Fig. 2). Further study is required to understand how these pathways interact with each other to reduce root elongation in response to high nitrate supply.Open in a separate windowFigure 2A simplified model explaining nitogen-mediated root elongation in maize.     

Induction as well as suppression: How aphid saliva may exert opposite effects on plant defense

Aphids ingest from the sieve tubes and by doing so they are confronted with sieve-tube occlusion mechanisms, which are part of the plant defense system. Because aphids are able to feed over longer periods, they must be able to prevent occlusion of the sieve plates induced by stylet penetration. Occlusion probably depends upon Ca²⁺-influx into the sieve element (SE) lumen. Aphid behavior, biochemical tests and in vitro experiments demonstrated that aphid''s watery saliva, injected during initial phase of a stylet penetration into the SE lumen, contains proteins that are able to bind calcium and prevent calcium-induced SE occlusion. In this addendum, we speculate on the consequences of saliva secretion for plant resistance. (a) The release of elicitors (e.g., oligogalacturonides) due to cell wall digestion by gel saliva enzymes may increase the resistance of cortex, phloem parenchyma cells and companion cells (CC) around the puncture site. (b) Ca²⁺-binding by aphid watery saliva may suppress the local defense responses in the SEs. (c) Signaling cascades triggered in CCs may lead to systemic resistance.Key words: aphid saliva, calcium binding, elicitor, oligogalacturonides, local plant defense, systemic plant defense, phloem translocation, aphid/plant-interactionAfter having penetrated the sieve-element (SE) plasma membrane, aphids encounter unspecific wound-induced occlusion reactions to prevent sap leakage.^1–4 Occlusion mechanisms by callose, structural P-proteins and forisomes are likely induced by a sudden calcium influx into the sieve-tube lumen.⁵ Calcium possibly enters the sieve-tube lumen through the stylet wounding-site in the plasma membrane and/or stretch-activated calcium-channels.^6–8 After SE penetration, aphids secrete watery saliva that contains calcium-binding proteins presumed to sabotage sieve-plate occlusion.^9,10We demonstrated that Megoura viciae (Buckton) is most likely able to prevent or reduce sieve-tube occlusion in Vicia faba by secretion of watery saliva. By in vitro confrontation of isolated forisomes, protein bodies responsible for sieve-tube occlusion in Fabaceaen,⁵ and watery saliva concentrate, we were able to show that salivary proteins convey forisomes from a dispersed (+Ca²⁺) into a condensed (−Ca²⁺) state.¹⁰ The dispersed forisome functions in vivo as a plug, leading to stoppage of mass flow.⁵This in vitro evidence was corroborated by aphid behavior in response to leaf tip burning, which triggers an electrical potential wave (EPW) along the sieve tubes. Such an EPW induces Ca²⁺-influx and corresponding SE occlusion along the pathway.¹¹ The passage of the EPW is associated with a prolonged secretion of watery saliva of aphids. This is interpreted as an attempt to unplug the SEs by calcium binding.¹⁰ Similar behavioral changes in response to leaf-tip burning were observed in an extended set of aphid/plant species combinations, indicating that attempted sabotage of sieve-tube occlusion by aphid saliva is a widespread phenomenon (unpublished).Aphid feeding was reported to induce local (on the same leaf) and systemic (in distant leaves) reactions of the host plant. The local response led to enhanced feeding,^12–14 while the systemic response showed reduced ingestion and extended periods of watery saliva secretion in sieve tubes distant from previous feeding sites.^12–14 These contrasting observations were described to be independent of the aphid species.¹³ The question arises how aphids induce these seemingly opposite plant responses?The aphid stylet pushing forward through cortical and vascular tissue is surrounded by a sheath of gel saliva, secreted into the apoplast.^15,16 Gel saliva contains cellulase and pectinase that amongst others produce oligogalacturonides (OGs) along the stylet sheath by digestion of cell wall material.^17,18 Usually, OGs act as elicitors, triggering a variety of plant responses against pathogens and insects in which the activation of calcium channels is involved.^19,20 This seems to conflict with a suppression of resistance as observed for the impact of watery saliva in SEs.¹⁰ We will make an attempt to explain this paradoxon.OG induced defense responses may be triggered in all cell types adjacent to the salivary sheath (Fig. 1). Because watery saliva is only secreted briefly into these cells, which are punctured for orientation purposes (Hewer et al., unpublished), it seems unlikely that OG induced defense is suppressed here by saliva-mediated calcium binding.¹⁵ The diffusion range of OGs may be restricted to the close vicinity of the stylet sheath leading to an enhanced regional defense with a limited sphere of action (Fig. 1). Because the settling distance of aphids is restricted by their body size (1–10 mm),²¹ aphids feeding on the same leaf are probably hardly confronted with the regional defense induced by another aphid (Fig. 1). Otherwise, they would show an increased number of test probes before first phloem activity, as described for volatile mediated plant defense in cortex cells.¹³ Circumstantial support in favor of our hypothesis is provided by production of hydrogen peroxide in the apoplast,²² which is most likely associated with the action of OGs.²² Observations of hydrogen peroxide production during aphid (Macrosiphum euphorbiae) infestation of tomato in a limited area along the leaf veins, the preferred feeding sites of this species, indicate a locally restricted defense response (Fig. 1 and ​and22).⁴ The question arises why the cell signals are not spread via plasmodesmata to adjacent cells to induce resistance in a more extended leaf area? Dissemination of the signals may be prevented by closure of plasmodesmata (Fig. 1) through callose deposition,^23,24 which is most likely directly coupled with calcium influx induced by OGs,²⁵ by apoplastic hydrogen peroxide and to a minor extent by stylet puncture (Fig. 2).^7,26Open in a separate windowFigure 1Hypothetical model on how stylet penetration induces and suppresses plant defense. Sheath saliva (light blue) that envelopes the stylet during propagation through the apoplast contains cellulase and pectinase,^17,18 enzymes producing elicitors (e.g., oligogalacturonides (oGs)) by local cell wall digestion.¹⁹ Parenchyma cells adjacent to the sheath may develop a defense response owing to signaling cascades triggered by oG-mediated Ca²⁺-influx.¹⁹ Together with a Ca²⁺-dependent transient closure of plasmodesmata by callose (black crosses),^23,24 the focused production of oGs may cause a defense response with a limited sphere of action (red—strong, brown—light, green—none). This restricted domain of defense may not be perceived by other aphids, since the settling distance is limited by the aphid body size. Nearby aphids do not show any sign of defense perception in their probing and feeding behavior.¹⁴ Signaling cascade compounds may be channeled from parenchyma cells to CCs (dashed yellow arrows), where they are subsequently released into the SEs. There they may act as long-distance systemic defense components (grey arrows). In contrast to the parenchyma domain (where only minor amounts of watery saliva are secreted), Ca²⁺-mediated reactions such as defense cascades and sieve-plate (SP) occlusion are suppressed in SEs by large amounts of watery saliva. The left aphid penetrates an SE and injects watery saliva (red cloud; ws) that inhibits local sieve-plate occlusion and,¹⁰ most likely, is transported by mass flow (black arrow) to adjacent SEs,²⁷ where occlusion is impeded as well. A short-distance systemic spread over a few centimeters may explain local suppression of plant defense resulting in a higher rate of colonization. Salivary proteins or their degradation products may serve as systemic defense signals as well (grey arrows), but may also diffuse via the PPUs into CCs where additional systemic signals are induced (yellow arrows).Open in a separate windowFigure 2Hypothetical involvement of Ca²⁺-channels in aphid-induced cell defense (detail). During probing with its stylet the aphid secretes gel saliva as a lubrication substance (light blue) into the apoplast.¹⁵ on the way to the sieve tubes, aphids briefly puncture most non-phloem cells (red) after which the puncturing sites are sealed with gel saliva.^7,16 Gel saliva also most likely prevents the influx of apoplastic calcium into pierced sieve elements (green) by sealing the penetration site.⁷ Watery saliva (red cloud), injected into the SE lumen,⁹ contains proteins which bind calcium ions (marked by X) that enter the SE via e.g., mechano sensitive Ca²⁺-channels activated by stylet penetration (blue tons).¹⁰ In this way, aphids suppress SE occlusion and activation of local defense cascades. In the parenchyma cells around the gel saliva sheath, a small cylindrical zone of defense may be induced by oligogalacturonides (oGs; brown triangles) produced by cell wall (grey) digestion.^17–19 Perceived by unknown receptor proteins (R; e.g., a receptor like protein kinase)³⁴ and kinase mediation (black dotted and dashed arrows), oGs lead to a Ca²⁺-influx through kinase activated calcium channels (orange tons).²⁵ Around the probing site, aphids apparently induce the production of superoxide by Ca²⁺-induced activation of the NADPH oxidase (violet box) and its following conversion to hydrogen peroxide (red spots) is mediated by superoxide dismutase (SoD).⁴ Hydrogen peroxide activates Ca²⁺-channels (violet tons) and diffuses through plasma membrane (curled arrows) therefore potentially acting as a intracellular signal.²⁶By contrast, Ca²⁺-influx into SEs, induced by presence of OGs or stylet insertion (Fig. 2), is not expected to trigger local defense given the abundant excretion of Ca²⁺-binding watery saliva.^7,10,25 Watery saliva may spread to down-stream and adjacent SEs through transverse and lateral sieve plates (Fig. 1).^7,27 Aphids puncturing nearby SEs may therefore encounter less severe sieve-plate occlusion which results in facilitated settling and thus in increased population growth. Aggregation of feeding aphids would self-amplify population growth until a certain density is attained. Farther from the colonization site, this effect may be lost due to dilution. Stimulation of aphid feeding by aphid infestation was observed locally on potato by Myzus persicae and M. euphorbiae, respectively, 96 h after infestation.¹³ However, a similar effect was not observed for M. persicae on Arabidopsis thaliana where aphids induced premature leaf senescence and resistance 12 h after infestation,²⁸ possibly induced by OGs.¹⁹As a speculation, OG induced Ca²⁺-influx into parenchyma cells adjacent to the salivary sheath activate Ca²⁺-induced signaling cascades via CaM,^26,29 CDPKs,^30,31 MAPKinases and reactive oxygen species (Fig. 2).³² Systemic resistance, induced by aphid infestation,^12–14 is mediated by unknown compounds such as, e.g., salivary proteins, their degradation products, signal cascade products or volatiles.¹³ Compounds produced in CCs first have to pass the PPUs, while SE signaling elements can be directly transported via mass flow (Fig. 1).The question arises if aphids profit from induced resistance on local (cortex and parenchyma cells) and systemic (distant plant organs) levels as holds for suppression of defense in SEs. Possibly settling and subsequent spread of competing pathogens/herbivores (e.g., fungi or other piercing-sucking insects) are suppressed by induced defense. In this context it is intriguing to understand how aphids cope with the self-induced systemic resistance, which probably lasts over weeks.³³