Co-regulation of root hair tip growth by ROP GTPases and nitrogen source modulated pH fluctuations

Growth of plant cells involves tight regulation of the cytoskeleton and vesicle trafficking by processes including the action of the ROP small G proteins together with pH-modulated cell wall modifications. Yet, little is known on how these systems are coordinated. In a paper recently published in Plant Cell and Environment¹ we show that ROPs/RACs function synergistically with NH₄NO₃-modulated pH fluctuations to regulate root hair growth. Root hairs expand exclusively at their apical end in a strictly polarized manner by a process known as tip growth. The highly polarized secretion at the apex is maintained by a complex network of factors including the spatial organization of the actin cytoskeleton, tip-focused ion gradients and by small G proteins. Expression of constitutively active ROP mutants disrupts polar growth, inducing the formation of swollen root hairs. Root hairs are also known to elongate in an oscillating manner, which is correlated with oscillatory H⁺ fluxes at the tip. Our analysis shows that root hair elongation in wild type plants and swelling in transgenic plants expressing a constitutively active ROP11 (rop11^CA) is sensitive to the presence of NH₄⁺ at concentrations higher than 1 mM and on NO₃⁻. The NH₄⁺ and NO₃⁻ ions did not affect the localization of ROP in the membrane but modulated pH fluctuations at the root hair tip. Actin organization and reactive oxygen species distribution were abnormal in rop11^CA root hairs but were similar to wild-type root hairs when seedlings were grown on medium lacking NH₄⁺ and/or NO₃⁻. These observations suggest that the nitrogen source-modulated pH fluctuations may function synergistically with ROP regulated signaling during root hair tip growth. Interestingly, under certain growth conditions, expression of rop11^CA suppressed ammonium toxicity, similar to auxin resistant mutants. In this short review we discuss these findings and their implications.Key words: ROP, RAC, nitrogen, root hair, cell polarity, ammoniumIn Arabidopsis, root hairs grow out at the basal, rootward region (closer to root tip) of specialized root epidermal cells and expand exclusively at their apical end in a strictly polarized manner by a process known as tip growth. Tip growth is facilitated by Rho of Plants (ROP)-regulated processes such as maintenance of longitudinally-oriented actin cables in the shank of the root hair that are required for myosin-mediated organelle transport through the cytoplasm. ROPs also play a role in sustaining fine F-actin structures at the root hair tip, which promote the transport of secretory vesicles to sites of their fusion with the plasma membrane.^2,3 In addition, the polar growth of root hairs involves an oscillatory tip-focused Ca²⁺ gradient⁴ and tip-localized reactive oxygen species (ROS).⁵ Tip growth is also associated with oscillatory fluxes of H⁺ at the apex that correlate with the periodicity of growth.^6,7 These oscillations in extracellular pH and ROS have been shown to modulate tip growth and are predicted to act in a coordinated and complementary mode to regulate root hair elongation. Growth accelerates following reduction of apoplastic pH and slows upon apoplastic ROS increase and a coincident pH increase.⁷ROPs are small G proteins that localize to the plasma membrane at the apex of growing root hairs, where they activate a range of downstream pathways required for tip growth.^8,9 ROP activity is regulated by its cycling between a GTP-bound, active and GDP-bound, inactive state. Ectopic expression of constitutively active mutants of ROPs (dominant mutations in conserved residues that abolish the GTPase activity) depolarizes the growth of root hairs.^8–10 Downstream pathways activated by such ROP GTPases include the regulation of cytoskeletal dynamics and vesicular trafficking, production of ROS, maintenance of intracellular Ca²⁺ gradients and accumulation of signaling lipids, features all related to the regulation of apical growth.^11,12 For example, ectopic expression of constitutively active ROP11 (Atrop11^CA) depolarizes root hair growth, leading to the formation of swollen root hairs. This bulging root hair phenotype was associated with altered actin organization and inhibition of endocytosis.¹⁰It is well known that root hair development is highly plastic and regulated by environmental signals.^13,14 Yet, despite the known function of ROP GTPases and their regulatory proteins in root hair growth there is no data in the literature describing the relationship between ROP signaling and environmental factors in this process. Our results¹ show that induction of root hair swelling by rop11^CA occurs only under specific growth conditions, indicating that there is an interplay between ROP activity and the external environment, particularly nitrogen supply. We demonstrated that high external concentrations of ammonium are essential for the induction of depolarized root hair growth and activation of downstream pathways by rop11^CA. Depletion of ammonium did not affect the membrane localization and expression of GFP-rop11^CA, implying that NH₄⁺ was required in addition to ROP activity to cause root hair swelling. In agreement with this idea, normal actin organization and ROS localization were detected in rop11^CA root hairs when NH₄⁺ was depleted, suggesting that ammonium functions downstream of, or in parallel to ROP signaling (Fig. 1).Open in a separate windowFigure 1A model for regulation of root hair tip growth by ROP GTPases and pH oscillations dependent on nitrogen supply. GTP bound ROPs activate downstream effectors which directly affect actin organization, vesicular trafficking and localized ROS production as well as indirectly affecting the localization of membrane proteins involved in ion/proton fluxes. High concentrations of nitrogen ions in the growth medium increase pH oscillations at the apex of growing root hairs. In turn downstream ROP effectors sense the changes in pH and adjust their function accordingly. pH oscillations affect tip growth independent of ROPs via changes of wall pH and possibly through additional unknown factors. Dashed lines indicate that these effects were not confirmed experimentally.Plants can absorb and use various forms of nitrogen from soils, primarily the inorganic ions ammonium and nitrate. The concentrations of these ions are highly heterogeneous around the plant and can vary across several orders of magnitude among different soils and as a result of seasonal changes.¹⁵ Thus, plants would be expected to display highly plastic, N-regulated developmental responses and to employ a range of nitrogen uptake transport systems to optimize exploitation of local N resources. Transport systems that mediate NH₄ fluxes across the plasma membrane of root cells are divided into two categories: high affinity transport systems (HATS) that mediate uptake from relatively dilute solutions at relatively low rates and low affinity transport systems (LATS) that operate at high rates and higher external concentrations.¹⁶ The HATS are plasma membrane localized NH₄⁺-specific transporters (AMTs) that are most likely proton-coupled and their expression and function are repressed at external ammonium concentrations of 1 mM or higher.^17–19 In contrast, ammonium uptake by LATS is believed to take place through non-specific cation channels.^17,20 The NH₄⁺ concentration in the 0.5× Murashige Skoog (MS) medium is 10.3 mM, exceeding by an order of magnitude the concentration at which the high affinity NH₄⁺ uptake system is repressed. The root hair swelling in Atrop11_CA plants and inhibition of root hair elongation in wild type plants occurred primarily at external ammonium concentrations greater than 1 mM, and thus is most likely associated with uptake by the LATS.As noted above, root hair elongation is associated with oscillations of cytoplasmic and apoplastic pH that have been linked to growth control. Simultaneous fluorescence ratio imaging of internal and external pH revealed that application of 10 mM NH₄NO₃ enhanced the amplitude of these pH oscillations at the extreme apex of wild type root hairs1 and Figure 2. These oscillations are thought to modulate tip growth through altering the extensibility of the wall.⁴ Additional measurements (Fig. 2) show that similar to the effects of NH₄NO₃, addition of NH₄Cl induced increase in the apoplastic pH fluctuations and reduced the pH. However, the effects of NH₄Cl on cytoplasmic pH fluctuations seem subtler compared to the effects of NH₄NO₃. Thus, one possible explanation for the observed swelling of the root hair apex in rop11_CA expressing plants in media containing NH₄NO₃ is that rop11_CA root hairs are affected in their ability to re-establish the normal proton gradient across the plasma membrane in response to ammonium transport. The altered proton gradient would then prevent the normal localized oscillatory changes in pH-dependent wall properties required to restrict expansion to the very tip of the elongating root hair.Open in a separate windowFigure 2Changes in apoplastic and cytoplasmic pH fluctuations, following application of NH₄NO₃, NH₄Cl or KNO₃. (A) Apolplastic pH (pH_ex) following treatments with either NH₄NO₃, NH₄Cl or KNO₃. Note the increase pH fluctuations induced by either NH₄NO₃ and NH₄Cl but not by KNO₃. (B) Cytoplasmic pH (pH_cyt) following treatments as above. Note the changes in pH fluctuations induced by NH₄NO₃ and the subtler effects of NH₄Cl.Concurrent absorption of NH₄⁺ and NO₃⁻- maintains the cation-anion balance within both the rooting medium and the root, and thus potentially has an important function in maintaining intracellular and extracellular pH.^21,22 In agreement, application of these ions affected the amplitude of pH oscillations¹ and Figure 2. Interestingly, treatments of WT seedlings with 10 mM NH₄NO₃ causes increase in root hair pH oscillations and often tip bursting. Yet, prolonged exposure of WT root hairs to NH₄NO₃ is accompanied by adaptation (our unpublished data). This adaptation does not occur in rop11_CA mutants, suggesting that cycling of ROPs between active and inactive states maybe important in adaptation to changing environment. These data strongly suggest that NH₄⁺-dependent root hair swelling in the plants expressing activated ROP resulted from physiological changes in ion balance rather than a direct effect of ammonium on enzymatic activities required for root hair growth (Fig. 1). Application of NH₄⁺ and NO₃⁻, in the absence of other ions, induced formation of additional growth tips, in which the membrane localized GFP-rop11^CA was concentrated. This observation suggests that interplay between the regulation of ROP localization and activity and the regulation of nitrogen fluxes may have an important function in the maintenance of unidirectional growth. As root hair elongation is coupled to spatially distinct regulation of extracellular pH oscillations and ROS production,⁷ it seems likely that there is a mechanism that can adjust the fluxes of nitrogen ions relative to these pH fluxes. This system would then maintain the oscillations in pH such that polarized growth is continued. One possible mechanism for this coordination is through the highly localized ROP cycling between active and inactive states that has an important role in the spatial activation of cell polarization machinery.^23–27 Due to the function of ROP GTPases in vesicle trafficking, actin organization and maintenance of ROS and Ca²⁺ gradients,^{2,8,9,23,24,28–33} expression of activated ROP11 may indirectly influence cell wall properties by altering the localization and/or recycling of cation and anion transporters/channels or plasma membrane H⁺-ATPases delivered to the growing tip of the hair and in this way affect the maintenance of the proton gradients. In agreement with a possible effect of activated ROPs on localization and/or recycling of membrane transporters we discovered that rop11^CA plants were resistant to ammonium toxicity when grown in the presence of NH₄NO₃ and several micronutrients.¹We propose a model (Fig. 1) in which spatial regulation of ROP activity creates a positive feedback loop with pH oscillations around the growing apex of root hairs. According to this model ROP cycling between active and inactive states spatially and temporally activates the downstream signaling cascades essential for the tip-growth of root hairs. At the same time, localization of membrane proteins involved in maintenance of normal nitrogen fluxes across the plasma membrane is indirectly affected by ROP signaling. Alternatively, ROP signaling is modulated to adapt to altered nitrogen fluxes. NH₄⁺ fluxes increase the amplitude of pH oscillations at the root hair apex and in turn affect cell-wall properties. Thus, when the ROP activity is upregulated by dominant mutations, the synergistic effects of pH changes and constant activation of ROP downstream effectors result in the uncontrolled cell expansion seen as root hair bulging. Previous studies have suggested that feedback between oscillatory pH change and ROS distribution is required to support tip growth.⁷ However, the factors that may integrate these processes are unknown. Our results suggest that spatial regulation of ROP activity in response to changing environments is one of the key elements that may coordinate the pH and ROS oscillations during the root hair tip growth.It will be interesting to examine whether ROP function is coordinated with apoplastic pH fluctuation in other cell types. Recently, it has been suggested that the effects of auxin on pavement cell structure in leaf epidermis require Auxin Binding Protein 1 (ABP1) dependent ROP activation.³⁴ It is well known that auxin induces changes in apoplastic pH. Possibly, like nitrogen source in root hairs, auxin dependent apolplastic pH fluctuations in the leaf epidermis may function coordinately with ROP in the regulation of cell growth. Consistent with this idea, it has been shown that auxin inhibits clathrin-dependent endocytosis through ABP1 reinforcing a possible role in modulating membrane flux/membrane properties.³⁵ Some auxin resistant mutants also display resistance to ammonium toxicity³⁶ further suggesting a link between auxin and membrane transport. Hence, auxin and ROPs may indeed function synergistically to modulate plasma membrane properties, in turn affecting ion balance in the apoplast and so modulating cell wall properties and growth.     

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

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

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.     

ARF-GTPase as a Molecular Switch for Polar Auxin Transport Mediated by Vesicle Trafficking in Root Development

Polar auxin transport (PAT), which is controlled precisely by both auxin efflux and influx facilitators and mediated by the cell trafficking system, modulates organogenesis, development and root gravitropism. ADP-ribosylation factor (ARF)-GTPase protein is catalyzed to switch to the GTP-bound type by a guanine nucleotide exchange factor (GEF) and promoted for hybridization to the GDP-bound type by a GTPase-activating protein (GAP). Previous studies showed that auxin efflux facilitators such as PIN1 are regulated by GNOM, an ARF-GEF, in Arabidopsis. In the November issue of The Plant Journal, we reported that the auxin influx facilitator AUX1 was regulated by ARF-GAP via the vesicle trafficking system.¹ In this addendum, we report that overexpression of OsAGAP leads to enhanced root gravitropism and propose a new model of PAT regulation: a loop mechanism between ARF-GAP and GEF mediated by vesicle trafficking to regulate PAT at influx and efflux facilitators, thus controlling root development in plants.Key Words: ADP-ribosylation factor (ARF), ARF-GAP, ARF-GEF, auxin, GNOM, polar transport of auxinPolar auxin transport (PAT) is a unique process in plants. It results in alteration of auxin level, which controls organogenesis and development and a series of physiological processes, such as vascular differentiation, apical dominance, and tropic growth.² Genetic and physiological studies identified that PAT depends on efflux facilitators such as PIN family proteins and influx facilitators such as AUX1 in Arabidopsis.Eight PIN family proteins, AtPIN1 to AtPIN8, exist in Arabidopsis. AtPIN1 is located at the basal side of the plasma membrane in vascular tissues but is weak in cortical tissues, which supports the hypothesis of chemical pervasion.³ AtPIN2 is localized at the apical side of epidermal cells and basally in cortical cells.^1,4 GNOM, an ARF GEF, modulates the localization of PIN1 and vesicle trafficking and affects root development.^5,6 The PIN auxin-efflux facilitator network controls root growth and patterning in Arabidopsis.⁴ As well, asymmetric localization of AUX1 occurs in the root cells of Arabidopsis plants,⁷ and overexpression of OsAGAP interferes with localization of AUX1.¹ Our data support that ARF-GAP mediates auxin influx and auxin-dependent root growth and patterning, which involves vesicle trafficking.¹ Here we show that OsAGAP overexpression leads to enhanced gravitropic response in transgenic rice plants. We propose a model whereby ARF GTPase is a molecular switch to control PAT and root growth and development.Overexpression of OsAGAP led to reduced growth in primary or adventitious roots of rice as compared with wild-type rice.¹ Gravitropism assay revealed transgenic rice overxpressing OsAGAP with a faster response to gravity than the wild type during 24-h treatment. However, 1-naphthyl acetic acid (NAA) treatment promoted the gravitropic response of the wild type, with no difference in response between the OsAGAP transgenic plants and the wild type plants (Fig. 1). The phenotype of enhanced gravitropic response in the transgenic plants was similar to that in the mutants atmdr1-100 and atmdr1-100/atpgp1-100 related to Arabidopsis ABC (ATP-binding cassette) transporter and defective in PAT.⁸ The physiological data, as well as data on localization of auxin transport facilitators, support ARF-GAP modulating PAT via regulating the location of the auxin influx facilitator AUX1.¹ So the alteration in gravitropic response in the OsAGAP transgenic plants was explained by a defect in PAT.Open in a separate windowFigure 1Gravitropism of OsAGAP overexpressing transgenic rice roots and response to 1-naphthyl acetic acid (NAA). (A) Gravitropism phenotype of wild type (WT) and OsAGAP overexpressing roots at 6 hr gravi-stimulation (top panel) and 0 hr as a treatment control (bottom panel). (B) Time course of gravitropic response in transgenic roots. (C and D) results correspond to those in (A and B), except for treatment with NAA (5 × 10⁻⁷ M).The polarity of auxin transport is controlled by the asymmetric distribution of auxin transport proteins, efflux facilitators and influx carriers. ARF GTPase is a key member in vesicle trafficking system and modulates cell polarity and PAT in plants. Thus, ARF-GDP or GTP bound with GEF or GAP determines the ARF function on auxin efflux facilitators (such as PIN1) or influx ones (such as AUX1).ARF1, targeting ROP2 and PIN2, affects epidermal cell polarity.⁹ GNOM is involved in the regulation of PIN1 asymmetric localization in cells and its related function in organogenesis and development.⁶ Although VAN3, an ARF-GAP in Arabidopsis, is located in a subpopulation of the trans-Golgi transport network (TGN), which is involved in leaf vascular network formation, it does not affect PAT.¹⁰ OsAGAP possesses an ARF GTPase-activating function in rice.¹¹ Specifically, our evidence supports that ARF-GAP bound with ARF-GTP modulates PAT and gravitropism via AUX1, mediated by vesicle trafficking, including the Golgi stack.¹Therefore, we propose a loop mechanism between ARF-GAP and GEF mediated by the vascular trafficking system in regulating PAT at influx and efflux facilitators, which controls root development and gravitropism in plants (Fig. 2). Here we emphasize that ARF-GEF catalyzes a conversion of ARF-bound GDP to GTP, which is necessary for the efficient delivery of the vesicle to the target membrane.¹² An opposite process of ARF-bound GDP to GTP is promoted by ARF-GTPase-activating protein via binding. A loop status of ARF-GTP and ARF-GDP bound with their appurtenances controls different auxin facilitators and regulates root development and gravitropism.Open in a separate windowFigure 2Model for ARF GTPase as a molecular switch for the polar auxin transport mediated by the vesicle traffic system.     

Tubular actin filaments in tobacco guard cells

The dynamic remodeling of actin filaments in guard cells functions in stomatal movement regulation. In our previous study, we found that the stochastic dynamics of guard cell actin filaments play a role in chloroplast movement during stomatal movement. In our present study, we further found that tubular actin filaments were present in tobacco guard cells that express GFP-mouse talin; approximately 2.3 tubular structures per cell with a diameter and height in the range of 1–3 µm and 3–5 µm, respectively. Most of the tubular structures were found to be localized in the cytoplasm near the inner walls of the guard cells. Moreover, the tubular actin filaments altered their localization slowly in the guard cells of static stoma, but showed obvious remodeling, such as breakdown and re-formation, in moving guard cells. Tubular actin filaments were further found to be colocalized with the chloroplasts in guard cells, but their roles in stomatal movement regulation requires further investigation.Key words: actin dynamics, tubular actin filaments, chloroplast, guard cell, stomatal movementStomatal movement responses to surrounding environment are mediated by guard cell signaling.^1,2 Actin filaments within guard cells are dynamic cytoarchitectures and function in stomatal development and movement.³ Arrays of actin filaments in guard cells that are dependent on different stomatal apertures have also been reported in references ^4–7. For example, the random or longitudinal orientations of actin filaments in closed stomata change to a radial orientation or ring-like array after stomata opening.^5,6,8 The reorganization of the actin architecture during stomatal movement depends on the depolymerization and repolymerization of actin filaments in guard cells. In contrast to the traditional treadmill model of actin dynamic mechanisms, stochastic dynamics of actin have been revealed in plant cells, such as in the epidermal cells of hypocotyl and root, the pavement cells of Arabidopsis cotyledons, and the guard cells of tobacco (Nicotiana tabacum).^9–11 In this alternative system, the short actin fragments generated from severed long filaments can link with each other to form longer filaments by end-joining activity. The actin regulatory proteins, Arp2/3 complex, capping protein and actin depolymerizing factor (ADF)/cofilin, may also be involved in the stochastic dynamics of actin filaments.^12,13Using tobacco GFP-mouse talin expression lines, we have previously analyzed the stochastic dynamics of guard cell actin filaments and their roles in chloroplast displacement during stomatal movement.^6,11 We found from these analyses that another arrangement of actin filaments, i.e., tubular actin filaments, exists in the guard cells of these tobacco lines. We first found the circle-like actin filaments in 82% of the guard cells (counting 320 cells) in tobacco expressing GFPmouse talin when analyzing a single optical section (Fig. 1A). In a previous study of BY-2 cells expressing GFP-Lifeact labeled actin filaments, Smertenko et al. found similar structures, i.e., quoit-like structures or acquosomes in all of the plant tissues examined except growing root hairs.¹⁰ However, in our present analysis of serial sections, we determined that the circle-like actin filaments in the tobacco guard cells were long tubes (Fig. 1A), as the lengths (about 3–5 µm) of these structures were greater than their diameter (about 1–3 µm). Hence, we denoted these structures as tubular actin filaments to distinguish them from the circular conformations of actin filaments observed previously in other plant cell tissues.^10,14–19 About 2.3 of these tubular actin filaments were found per guard cell, which is less than the number of acquosomes reported in BY-2 cells (about 6.7 per cell).¹⁰ Analysis of serial optical sections at the z-axis revealed that the tubular actin filaments localize in the cytoplasm near the inner walls of the guard cells (Fig. 1B), which is similar to the distribution of chloroplasts in guard cells.¹¹ Longitudinal sections further revealed a colocalization of tubular actin filaments and chloroplasts (Fig. 1B).Open in a separate windowFigure 1Tubular actin filaments in the guard cells of a tobacco (Nicotiana tabacum) line expressing GFP-mouse talin. (A) Optical-sections (interval, 1.5 µm) of guard cells in a moving stoma showing tubular actin filaments (arrow heads). Frames (a1) and (a2) are cross sections of 1.5-µm-picture through the yellow and red lines, respectively, revealing the cross section of the circle structures are parallel lines (arrows). (B) Optical-sections of a stoma from the outer periclinal walls to the inner walls of the guard cells (interval, 1 µm). The tubular actin filaments (arrow heads) are localized in the cytoplasm near to the inner periclinal walls of guard cells. Frame (b1) is the guard cell on the right of the frame “4 µm”; (b2) is the cross section of b1 through the red line; and (b3) is a higher magnification image of the area encompassed by the white square in b2. Arrows indicate the colocalization between the tubular actin filaments and the chloroplast (indicated using a red pseudocolor). (C) Time-series imaging showing the movement of tubular actin filaments in the guard cells of static stomata. Frame (c1) comprises three images colored red (0 S), green (40 S) and blue (80 S), that are merged in a single frame to show the translocation of the tubular actin filaments (arrows). (D) Time-series images of the opening stomata showing the breakdown (arrows) and re-formation (arrowheads) of the tubular actin filaments. All images were captured using a Zeiss LSM 510 META confocal laser scanning microscope, as described by Wang et al.¹¹ Bars, 10 µm.We performed time-lapse imaging and found that the translocation of tubular actin filaments is slow in static stomata in which the distance between two tubular actin filaments typically increased from 2.22 to 2.50 µm after 80 sec (Fig. 1C). In moving stomata, however, the tubular actin filaments showed an obvious dynamic reorganization whereby they could be processed into short fragments and also reemerged after they had disintegrated (Fig. 1D). These results indicate that tubular actin filaments have stochastic dynamics that are similar to the long actin filaments of guard cells.¹¹ In our previous study, we found that the stochastic dynamics of actin filaments correlate with light-induced chloroplast movement in guard cells.¹¹ However, whether the dynamics of the tubular actin filaments are also involved in chloroplast movement during stomatal movement remains to be investigated. In cultured mesophyll cells which had been mechanically isolated from Zinnia elegans, Wilsen et al. previously found a close association between fully closed actin rings and chloroplasts.¹⁸ These authors further found that the average percentage of cells with free actin rings increased at the initial culture stage, and then decreased, which indicates that the formation of actin rings might be a response of the actin cytoskeleton to cellular stress or disturbance.¹⁸ The turgor pressure of guard cells is the fundamental basis of stomatal movement leading to changes in the shape, volume, wall structure, and membrane surface of guard cells.^20–24 We speculate from our current data that there is a relationship between tubular actin filaments and the shape changes of guard cells during stomatal movement.     

Alamethicin-induced electrical long distance signaling in plants

Systemic signals induced by wounding and/or pathogen or herbivore attack may be realized by either chemical or mechanical signals. In plants a variety of electrical phenomena have been described and may be considered as signal-transducing events; such as variation potentials (VPs) and action potentials (APs) which propagate over long distances and hence are able to carry information from organ to organ. In addition, we recently described a new type of electrical long-distance signal that propagates systemically, i.e., from leaf to leaf, the “system potential” (SP). This was possible only by establishing a non-invasive method with micro-electrodes positioned in substomatal cavities of open stomata and recording apoplastic responses. Using this technical approach, we investigated the function of the peptaibole alamethicin (ALA), a channel-forming peptide from Trichoderma viride, which is widely used as agent to induce various physiological and defence responses in eukaryotic cells including plants. Although the ability of ALA to initiate changes in membrane potentials in plants has always been postulated it has never been demonstrated. Here we show that both local and long-distance electrical signals, namely depolarization, can be induced by ALA treatment.Key words: alamethicin, long distance electrical signal, depolarization, non-invasive recordingPeptaibols are linear membrane active peptide antibiotics produced by various fungi.^1,2 They are characterized by the presence of an unusual amino acid, α-aminoisobutyric acid, and a C-terminal hydroxylated amino acid and are generated by non-ribosomal peptide synthetases.^2,3 Due to their amphiphilic nature they self-associate into oligomeric ion-channels that span the width of lipid bilayer membranes.³ Peptaibols exhibit antibiotic activity against bacteria and fungi, tissue damage in insect larvae, as well as cytolytic activity towards mammalian cells.² ALA is a voltage-gated ion channel-forming peptide mixture that consists of at least 12 compounds each containing 20 amino acid residues.^1,2 This peptaibole is often used to elicit typical systemic responses in plants, many of which are in the context of indirect defences, such as the induction and accumulation of secondary metabolites in general, volatile compounds, and the phytohormones jasmonic acid and salicylic acid but also tendril coiling is induced.^4–7 Moreover, ALA has been shown to permeabilize mitochondria and the plasma membrane of tobacco suspension culture cells but not the tonoplast for small molecules.⁸ Thus, ALA is a valuable tool in plant science but its primary biological activity in plant tissues has never been shown experimentally.Because of the channel-forming properties of ALA, effects on the membrane potential of cells are very likely. Therefore, to monitor ALA application-elicited electrical signals, we employed the non-invasive method with microelectrodes placed in the apoplasm of the sub-stomatal cavities of open stomata. Whereas electrical signals in plants such as action potentials and variation potentials have been well documented in literature,⁹ this particular approach has been successfully applied for the detection and characterization of the novel system potentials.^10,11 With an intracellular recording the depolarization of a membrane occurs when the cell interior becomes less negative; whereas for the apoplastic recording used here, the inverse argument holds true. To avoid confusion, we follow the convention and call an apoplastic hyperpolarization a depolarization.^10,11ALA (mixture of several isoforms purchased from Sigma-Aldrich, Germany) was tested on the dicotyledonous lima bean (Phaseolus lunatus L.) and the monocotyledonous barley (Hordeum vulgare L.). As shown in Figure 1, upon a 5 nM ALA stimulus a 40 mV depolarization was induced and detected at a distance of 5 cm on the same leaf in lima bean. Systemic electric propagation was tested and demonstrated in barley (Fig. 2). We used barley for these experiments because we learned from earlier studies that electrical signals are not propagated between the primary leaves of lima bean whereas in barley electrical signals pass nodes and internodia more easily. Application of 25 nM ALA on one leaf (S-leaf) resulted in a 45 mV depolarization response at a distant of 25 cm on a different leaf (T-leaf), indicating that ALA induced a systemic electrical response that moved from one to the other leaf. In both plants the velocity of the propagating depolarization signal was calculated to be 2.5 cm min⁻¹. Whereas these results basically demonstrate the ability of ALA to initiate electrical signals in plant tissues both locally and systemically, the molecular interconnections between these electrical phenomena, the induction and synthesis of phytohormones as well as secondary metabolites, and how the ALA signal is transduced mechanistically, remains to be elucidated.Open in a separate windowFigure 1Local response of Phaseolus lunatus to Alamethicin. (A) Experimental setup for the measurement of apoplastic voltage changes using microelectrodes positioned in the sub-stomatal cavities of P. lunatus.¹¹ Stimuli and measurements of apoplastic voltage changes were performed on the same leaf with a distance of approximately 5 cm. The tip of the leaf was submerged in 5 mM KCl with 0.1 mM CaCl₂, pH 5, and the solution was connected to earth with a reference electrode filled with 0.5 M KCl. (B) The voltage response to 5 nM Alamethicin (ALA) (Sigma-Aldrich, Germany) added to a cut injury of the P. lunatus leaf at the indicated time shows a hyperpolarization of the apoplast (negative shift of the apoplastic potential), suggesting depolarization of the symplast. A typical recording of one out of three independent experiments is presented.Open in a separate windowFigure 2Systemic response of Hordeum vulgare to Alamethicin. (A) Experimental setup for the measurement of apoplastic voltage changes using microelectrodes positioned in the sub-stomatal cavities of H. vulgare.¹¹ Stimuli were applied to one leaf (Stimulus leaf, S-leaf), and measurements of apoplastic voltage changes were performed on a second leaf (Target leaf, T-leaf) with a distance of approximately 25 cm. The tip of the T-leaf was submerged in 5 mM KCl with 0.1 mM CaCl₂, pH 5, and the solution was connected to earth with a reference electrode filled with 0.5 M KCl. (B) The voltage response to 25 nM Alamethicin (ALA) (Sigma-Aldrich, Germany) added to a cut injury of the H. vulgare S-leaf at the indicated time shows a hyperpolarization of the apoplast (negative shift of the apoplastic potential), suggesting depolarization of the symplast. A typical recording of one out of three independent experiments is presented.     

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.     

Nitric oxide modulates sensitivity to ABA

Nitric oxide (NO) is a gas with crucial signaling functions in plant defense and development. As demonstrated by generating a triple nia1nia2noa1-2 mutant with extremely low levels of NO (February 2010 issue of Plant Physiology), NO is synthesized in plants through mainly two different pathways involving nitrate reductase (NR/NIA) and NO Associated 1 (AtNOA1) proteins. Depletion of basal NO levels leads to a priming of ABA-triggered responses that causes hypersensitivity to this hormone and results in enhanced seed dormancy and decreased seed germination and seedling establishment in the triple mutant. NO produced under non-stressed conditions represses inhibition of seed developmental transitions by ABA. Moreover, NO plays a positive role in post-germinative vegetative development and also exerts a critical control of ABA-related functions on stomata closure. The triple nia1nia2noa1-2 mutant is hypersensitive to ABA in stomatal closure thus resulting in a extreme phenotype of resistance to drought. In the light of the recent discovery of PYR/PYL/RCAR as a family of potential ABA receptors, regulation of ABA sensitivity by NO may be exerted either directly on ABA receptors or on downstream signalling components; both two aspects that deserve our present and future attention.Key words: nitric oxide, abscisic acid, seed germination, stomata openingPlant development is the result of the succesfull execution of several programs that control the transition between different growth phases. Every developmental transition is regulated through coordinated mechanisms that involved exogenous environmental factors such as light and temperature as well as endogenous cues, including levels of primary and secondary metabolites. Among the latter, hormones such as gibberellins (GA), auxins, citokinins, ethylene and abscisic acid (ABA) participate in the control of most of the developmental transitions.^1,2 During the last years, nitric oxide (NO) has gained an increasing role as an essential player in plant defense responses³ as well as a co-regulator of many developmental processes.⁴ However, studies of NO function as a regulatory molecule in plants have been hampered by the scanty, limited and controversial knowledge on how this gas is synthesized in plants.^5,6 This situation has moved researchers in this area to adopt pharmacological approaches based on chemicals acting as artificial NO donors as well as inhibitors or scavengers of NO action. The lack of specificity and the inherent artificial effects of these chemicals can be overcome by genetic approaches based on the use of mutants with endogenous low levels of NO. In February 2010 issue of Plant Physiology, we report the generation and further characterization of a triple nia1nia2noa1-2 mutant that contains extremely low levels of NO due to the impairment of two NO biosynthetic pathways involving nitrate reductase (NIA/NR) or NO Associated 1 (AtNOA1) proteins.⁷ These findings support that NO is mainly produced through those pathways in Arabidopsis. However, the possible existence of a minor still uncharacterized pathways involved in the residual production of NO can not be ruled out at this time.Further functional characterization of nia1nia2noa1-2 mutant in terms of development has pointed to NO as an overall positive regulator of plant growth, affecting to almost every developmental stage from seed germination to reproductive development. Accordingly, triple mutant plants display a delayed growth resulting in small shoot and root size and they also produce low amounts of viable seeds.⁷Dormancy and seed germination are developmental programs largely regulated by the combined action of GA and ABA.¹ GA promote breaking of dormancy and promote germination whereas ABA acts as a brake in those processes, thus ensuring a timely seed germination. Our data from the characterization of dormancy and seed germination in the nia1nia2noa1-2 mutant suggest that NO’s role in the control of those processes may be exerted through modulation of the sensitivity to ABA (Fig. 1A). Seeds from NO deficient plants have increased dormancy and lower seed germination and seedling establishment rates than wild type seeds due to the enhanced ABA inhibitory action. These effects can be reversed by exogenous application of NO to nia1nia2noa1-2, suggesting that the sensitivity to ABA is actually controlled by the endogenous levels of NO. The recent identification of PYR/PYL/RCAR family of ABA receptors,^8,9 and the further characterization of the essential ABA regulatory module including receptor, protein phosphatases of the 2C class and kinases of the SnRK2 family¹⁰ point to these components as potential targets of NO in regulating sensitivity to ABA (Fig. 1B). This work is in progress in our lab but we already know that some of the PYR/PYL/RCAR receptors and SnRKs are actually regulated by NO and also that this regulation may be exerted at different levels (Lozano-Juste J and León J, unpublished data).Open in a separate windowFigure 1Interactions between NO and ABA results in modulated sensitivity to ABA throughout development. (A) NO synthesized through nitrate reductase (NR/NIA) and NO associatedI (AtNOA1) protein regulate germinative and post-germinative development as well as stomata movements through modulation of the sensitivity to ABA. Arrows and bars represent positive and negative effects, and the thickness of lines are proportional to the magnitud of regulatory effects. (B) Scheme of a minimal ABA signalling module and the potential targets of NO. Dashed lines represent effects still to be demonstrated. (C) ABA signalling in stomata guard cells through Ca²⁺-dependent and -independent pathways and the potential interactions with NO as represented by dashed lines.The enhanced sensitivity to ABA observed in germinative and post-germinative development of nia1nia2noa1-2, is extended throughout plant life cycle and it is actually the cause of the very strong resistance of nia1nia2noa1-2 plants to water deficit conditions.⁷ Stomatal aperture is a fine-tuned process controlled mainly through a balance between the light-promoted opening and the ABA-mediated promotion of closure and inhibition of opening¹¹ (Fig. 1A). It has been previously reported that ABA function on stomata movements involve the participation of NO as well as Ca²⁺ in such a way that Ca²⁺ chelators and NO scavengers block ABA action on stomata movements.¹² Stomata of nia1nia2noa1-2 leaves, despite of being depleted of NO, are not impaired for ABA inhibition of stomata opening but, in turn, they seem to be primed for a more efficient ABA response (Fig. 1A). Contrary to the Ca²⁺ requirements for ABA action on wild type stomata movements, this process is not affected by Ca²⁺ chelators in nia1nia2noa1-2 stomata, and it thus seems to be independent of Ca²⁺ in NO-deficient backgrounds (Fig. 1C). As mentioned above, NO might regulate sensitivity to ABA by acting on ABA receptors or on SnRKs, some of which are Ca²⁺-independent kinases. Both receptors and Ca^2+--independent kinases are likely targets of NO in the modulation of stomata sensitivity to ABA thus explaining the more efficient stomata closure in nia1nia2noa1-2 leaves, and the consequent low rates of evapotranspiration that leads to the extreme resistance of triple mutant plants to drought.The future characterization of the interactions between NO and key components of ABA signaling will be the basis for a better knowledge of the functional interactions between different hormones in plant development and defense, but it will also open up new possibilities of identifying new targets and strategies leading to improved drought resistance.     

Assessing fungal root colonization for plant improvement

Fungal endophytes display a broad range of symbiotic interactions with their host plants. Current studies on their biology, diversity and benefits are unravelling their high relevance on plant adaptation to environmental stresses. Implementation of such properties may open new perspectives in agriculture and forestry. We aim to exploit the endophytic capacities of the fungal species Fusarium equiseti, a naturally occurring root endophyte which has shown antagonism to plant pathogens, and Pochonia chlamydosporia, a nematophagous fungus with putative endophytic behavior, for plant protection and adaptation to biotic and abiotic stress. A real-time PCR protocol for quantification of the fungal population, together with Agrobacterium-mediated genetic transformation with the GFP gene for confocal microscopy analyses, were designed and applied to assess endophytic development of both these fungal species. Although quantification of both F. equiseti and P. chlamydosporia yielded similar degrees of root colonization, microscopical observations demonstrated differences in infection and development patterns. Furthermore, we found evidences of plant response against endophyte colonization, supporting a balanced antagonism between the endophyte virulence and the plant defenses. Optimization and application of the methodologies presented herein will allow elucidation of beneficial interactions among these endophytes and their host plants.Key words: endophytes, Fusarium equiseti, fungal detection, mutualism, Pochonia chlamydosporia, root colonizationTissues of nearly all plants in natural ecosystems appear to be colonized by endophytic fungi, whose importance in plant development and distribution seem to be crucial, but not yet clearly understood. Fungal endophytes may assist their host plants for adaptation to habitat, protection against biotic or abiotic stresses, plant growth promotion or soil nutrients uptake.^1–5 Recently, biology and function of fungal endophytes in nature has been extensively reviewed,⁶ and these have been grouped into four classes according to their differential interactions with host plants, which may range from mere neutralism to an active mutualism.Exploitation of beneficial properties of endophytes is of great relevance at an applied level, either to increase production yields of agricultural crops, control of plant diseases or pests, adapt plants to unsuitable growth conditions, or in reforestation activities. In this sense, we aim to exploit the endophytic capacities of selected fungi from two independent approaches: through the use of natural endophytes, which have shown to confer benefits to their host plants,⁵ or via a putative endophytic colonization of plant tissues by fungi with desirable properties. The latter strategy has been already introduced for several biological control agents such as nematophagous fungi colonizing roots,⁷ or entomopathogenic fungi in above-ground organs.^8,9 As representatives of each category, we selected two fungal species according to their endophytic capacities and antagonism to root pathogens.^10,11 First, Fusarium equiseti represents naturally occurring endophytes from natural vegetation growing under saline stress conditions.¹² The second, Pochonia chlamydosporia, is a nematode egg-parasite which behaves as a putative endophyte of roots.¹³ Both species have promising properties for application as biocontrol agents of either fungal and/or nematode plant parasites. The achievement of these objectives will depend on an exhaustive knowledge on the endophytic behavior and interactions with the host and other existing microbes, which starts with the assessment of the establishment of fungal populations within plant organs (e.g., roots). In many instances this is a complicated task. Fungal growth is characterized by an implicit irregular development due to the filamentous nature of mycelia, whose complexity increases in structured substrates, such as the interior of plant tissues.These difficulties are enhanced due to uneven distribution of nuclei (from zero to several thousands per cell), physiological activity or vitality of hyphae, and influence of competition among species.¹⁴ As a consequence of this, methods applied traditionally to estimate fungal occurrence within plant tissues (e.g., direct visualization, plating on culture media or immunolocalization^4,15,16) are biased or rather laborious and time consuming. Although applications of molecular methods have been directed to settle these problems, these may also present other weaknesses. Combination of quantitative and qualitative data achieved by both molecular and microscopy methods is probably the best choice to monitor fungi in plant roots, since the advantages of each technique may complement the drawbacks of the other.^14,17We recently optimized and applied both real-time PCR and microscopy techniques to exhaustively study the endophytic development of F. equiseti and P. chlamydosporia in barley roots.¹⁸ The first approach consisted in a specific quantification of nucleic acids from either fungus in roots using Molecular beacon probes.¹⁹ This allows an accurate monitoring of the respective amplicon generation over PCR cyclings, which can be correlated with the amount of fungal biomass.²⁰ Fungal populations detected in roots were statistically similar for both fungal species studied, with an overall colonization of roots which ranged from ca. 5 to 11 ng of fungal target DNA per 100 ng of total DNA. The endophytic proportion of these populations (assessed by surface sterilization of roots prior to DNA extraction) was represented by values between ca. 0.5 to 1 ng of fungal target DNA per 100 ng of total DNA.These quantitative data resulted in an increased sensitivity and dynamic range as compared with traditional culturing methods. Nevertheless, though quantification yielded good results under a gnotobiotic system, where only those fungi of interest are present within plant tissues, implementation for non-axenic semi-field or field experimentation should include modifications. These should cover corrections for presence of other colonizing microorganisms under non-axenic growth conditions-which may contribute to the amount of total DNA extracted from roots-, or variability on DNA extraction yields among samples.²¹ Inclusion of internal standards such as the simultaneous detection of fungal and host plant in a single reaction tube²² would settle these interferences. Multiplex PCR amplification of respective plant or fungal loci would permit detection of differential fluorescent signals emitted by Molecular beacons specific for either target DNA. We are currently optimizing primers and probes designed in Maciá-Vicente et al.¹⁸ for multiplexing with primers and Molecular beacons specific for the host (barley) ubiquitin gene (Fig. 1).Open in a separate windowFigure 1Amplification by conventional PCR of DNA from plant (h; barley), endophyte (e; P. chlamydosporia), or endophyte-colonized plants (h + e). For this, single locus detection with primers specific for the fungal alkaline serine protease p1,¹⁸ (Fp) and the plant ubiquitin (Pp) is shown, and multiplex PCR using both primer pairs (Fp + Pp) for one-tube simultaneous detection.In addition to real-time PCR assays, endophytic behavior of F. equiseti and P. chlamydosporia was assessed using live-cell microscopy. Aiming to develop techniques for further studies in semi-field experiments, we generated genetically transformed isolates for both F. equiseti and P. chlamydosporia, with constitutive expression of the fluorescent reporter protein GFP. For this purpose an Agrobacterium tumefaciens-mediated protocol was applied due to its efficiency in fungal transformation.²³ Observation of barley roots inoculated with GFP-tagged isolates under laser scanning confocal microscopy permitted a time-course qualitative monitoring of the infection processes by F. equiseti and P. chlamydosporia, which displayed different endophytic patterns. Loading studies with the endocytotic tracker FM4-64 allowed a discrimination of new traits of fungal colonization of roots. Fungal hyphae appeared tightly fitted in a plant membrane-derived sheath during the first invasive stages, in a similar manner to that which occurs during pathogenesis,²⁴ but also in mutualistic interactions,²⁵ indicating it may be (at least originarly) an unspecific barrier to fungal invasion. However, this membrane was lost with time, as a consequence of hyphal aggressiveness over plant cell infection.Differential wavelength emission between GFP and FM4-64 also allowed detection of a heterogeneous distribution of viable and non-viable hyphae within the root cortex, the latter linked to plant defense responses such as abundant production of papillae or vacuoles. This colonization pattern suggests that endophytic interaction established by both fungi is a “push and pull” balance between hyphal growth and the capacity of the plant to get rid of the invader. Yet we do not know whether remaining undegraded nucleic acids contents of within-cortex dead hyphae may contribute to fungal target DNA detection by real-time PCR. Although this fact may represent a bias for endophytes quantification, combination with microscopical analyses complements the information achieved.We are currently investigating practical and theoretical aspects of the root inoculation of fungi with endophytic capabilities and antagonistic potential to root pathogens (fungi and nematodes). This research also includes evaluating root colonization abilities for different host plants, both monocots and eudicots. As an example, root colonization efficiency by P. chlamydosporia dramatically changes between barley and tomato. In the latter, hyphae within roots are sparse and restricted to epidermal root cells,¹³ with no evident connection between infected cells (Fig. 2A). In spite of this restricted distribution, fungal chlamydospores (the propagules usually found in P. chlamydosporia-harboring soils) may be frequently found on the root surface. Their viability (according to GFP expression in the cytoplasm), irrespective of that of surrounding hyphae (Fig. 2B), could support that root colonization creates a stable source of inoculum to sustain the populations of the microorganism in the rhizospheric soil. Our final goal is to optimize biocontrol performance and crop growth promotion by endophytes under agricultural conditions.Open in a separate windowFigure 2Laser scanning confocal microscopy images of one-month-old tomato roots colonized by GFP-tagged P. chlamydosporia. (A) P. chlamydosporia hypha restricted to a single epidermal root cell. (B) Chlamydospore in the root surface. Note GFP fluorescence within cells (viable) in contrast to lack of fluorescence in peduncle (non-viable). Bars = 20 µm.