Long-distance signals positively regulate the expression of iron uptake genes in tobacco roots

Long-distance signals generated in shoots are thought to be associated with the regulation of iron uptake from roots; however, the signaling mechanism is still unknown. To elucidate whether the signal regulates iron uptake genes in roots positively or negatively, we analyzed the expressions of two representative iron uptake genes: NtIRT1 and NtFRO1 in tobacco (Nicotiana tabacum L.) roots, after shoots were manipulated in vitro. When iron-deficient leaves were treated with Fe(II)-EDTA, the expressions of both genes were significantly reduced; nevertheless iron concentration in the roots maintained a similar level to that in roots grown under iron-deficient conditions. Next, all leaves from tobacco plants grown under the iron-deficient condition were excised. The expression of two genes were quickly reduced below half within 2 h after the leaf excision and gradually disappeared by the end of a 24-h period. The NtIRT1 expression was compared among the plants whose leaves were cut off in various patterns. The expression increased in proportion to the dry weight of iron-deficient leaves, although no relation was observed between the gene expression and the position of excised leaves. Interestingly, the NtIRT1 expression in hairy roots increased under the iron-deficient condition, suggesting that roots also have the signaling mechanism of iron status as well as shoots. Taken together, these results indicate that the long-distance signal generated in iron-deficient tissues including roots is a major factor in positive regulation of the expression of NtIRT1 and NtFRO1 in roots, and that the strength of the signal depends on the size of plants.     

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.     

Salt-avoidance tropism in Arabidopsis thaliana

The orientation of plant root growth is modulated by developmental as well as environmental cues. Among the environmental factors, gravity has been extensively studied because of its overpowering effects in modulating root growth direction. However, our knowledge of the effects of other abiotic signals that influence root growth direction is largely unknown. Recently, we have shown that high salinity can modify root growth direction by inducing rapid amyloplast degradation in root columella cells of Arabidopsis thaliana. By exploiting salt overly sensitive (sos) mutants and PIN2 expression analyses, we have shown that the altered root growth direction in response to salt is mediated by ion disequilibrium and is correlated with PIN2 mRNA abundance and expression and localization of the protein. Our study demonstrates that the SOS pathway may mediate this process. Here we discuss our data from broader perspectives. We propose that salt-induced modification of root growth direction is a salt-avoidance behavior, which is an active adaptive mechanism for plants grown under saline conditions. Furthermore, high salinity also stimulates alteration of gravitropic growth of shoots. These findings illustrate that plants have a fine and sophisticated sensory and communication system that enable plants to dynamically and efficiently cope with rapidly changing environment.Key words: abidopsis, adaptation, avoidance, root, salt stress, tropic growthOwing to their sessile nature, plant roots are constantly bombarded with various environmental stimuli from the soil, such as gravity, physical obstacles and imbalanced distribution of water and/or nutrients and high salinity. Where to grow is an important developmental decision in the life cycle of a plant that is crucial for its adaptation and the subsequent reproductive success. The proper orientation of root growth is shaped by both the developmental inputs and external signals.^1,2 The overwhelming environmental factor that modulates root growth direction is gravity, and plant primary roots grow downward toward the gravity vector. This directed growth of root in response to gravity is named as tropic growth to gravity or gravitropism. Studies of gravity perception and signaling pathway of the root cap at the primary root of Arabidopsis strongly support the starch statolith hypothesis.³ In this hypothesis, the columella cells in the root cap, which contain sedimentable amyloplasts, are the gravity-perceptive site in roots. The inner columella cells of the second tier have been proposed as making the greatest contribution to root gravitropism.⁴ Upon gravity stimulation, cytosolic ions such as Ca²⁺ and rapid cytoplasmic alkalization may be involved in gravity signal transduction.^5–7 Asymmetric distribution of auxin in roots caused by basipetal transport mainly through the auxin efflux carrier PIN-FORMED2 (PIN2), which is distributed asymmetrically within the cells, results in gravitropic root response of the root elongation zone.^8,9In contrast to our understanding of gravitropism of root, our knowledge of tropistic responses of root to other major environmental stimuli, such as water availability, imbalanced nutrient distribution and high salinity, and the interplay between these stimuli in determining the directional growth of root remains enigmatic. Recent studies have confirmed the existence of hydrotropism and the molecular genetic basis of the tropistic growth of root to water in determining the final direction of root growth starts to be deciphered.^10–12 Hydrotropic growth of roots is an important trait for plants to actively find water and to optimize their fitness under drought condition. Salinity is another major constraint to root system development, and limits the productivity of agricultural crops and the distribution of plant species.^13–15 It is known that salt stress-induced disturbed balance of ions is the primary cause for inhibition of plant growth and subsequent yield reduction. How does root minimize entrance of harmful ions and subsequently avoid salt injury? Does plant have capacity to sense salt signal, and prevent potentially harmful ions reaching root and shoot?Previous studies have shown that plant use different strategies to avoid salt injury at various levels. After Na⁺ enters the root cells, the Casparian strip can restrict the movements of the harmful ion into the xylem.¹⁶ Root cells also avoid salt injury by extruding Na⁺ actively back to the outside solution. This energy-dependent ion efflux from cytosol across the plasma membrane is mediated by SOS1 gene, a Na⁺-H⁺ antiporter, which is regulated by at least other genes, SOS3 (calcium binding protein) and SOS2 (serine/threonine kinase). This is the well characterized SOS (Salt Overly Sensitive) signaling pathway.^17,18 Another way for plant root cells to avoid ion injury is to accumulate Na⁺ into vacuole. Vacuolar compartmentation of Na⁺ is also in part regulated by Na⁺-H⁺ antiporters, such as AtNHX1.¹⁹ These findings reveal mechanisms of how plants avoid Na⁺ injury after passive entrance of sodium ions into root cells. We questioned whether a plant is capable of actively preventing the harmful ions from reaching root cells or escape from high salinity in the environment, and how plant roots respond to changing salt conditions, because salt distribution is unbalanced under natural saline conditions, especially after rain and irrigation. With a new assay that allows us to specifically address how plant roots respond to changing salt levels, we discovered an alternative adaptive mechanism for plant root to avoid salt injury.²⁰We set up a two-layer medium assay in which a sodium ion gradient would be generated. A normal nutrient agar medium is at the top of the growth bottle and an agar with salt-stressed medium is in the bottom of the bottle. This simple assay allows us to monitor root growth and orientation. The roots of the wild type seedlings penetrated the interface of the layers and grew straight downwards exhibiting gravitropism, when both layers were MS media. In contrast, when the bottom medium contained NaCl, roots of seedlings grew downward first, and then curved and grew upward toward the lower levels of salt. Roots started to bend upward at an early stage even before contacting high-salt medium (250 mM NaCl) on the bottom. The results indicate that roots can sense ion gradients in the growing environment and transduce the signal, combine with internal signals to make decisions that enable roots to stay away from high salt.^21,22 Here, we would like to propose this salt-induced tropic growth as a salt-avoidance tropism, which is an important adaptive behavior for plant roots to avoid salt injury and direct them toward their goal of optimal fitness.²³ Because salt stress inhibits root elongation, we tested impact of salt-induced negative gravitropism on the root growth. The results showed that inhibitory effect of salt on root growth was largely alleviated with this tropic curve (Fig. 1), further verifying our hypothesis that the salt-induced developmental plasticity is a salt-avoidance behavior (Fig. 2).Open in a separate windowFigure 1Effects of salt on root elongation of Arabidopsis thaliana seedlings from different salt treatments. The inhibitory effect of salt stress on root growth was greatly alleviated in the wild type (Col-0) when root growth of the seedlings was analyzed using a two-layer medium assay (black bars). The MS nutrient medium is on the top, and NaCl concentrations in the media on the bottom are 0, 150 and 250 mM. More severe inhibition of root growth of the seedlings by various levels of NaCl in a root bending assay (white bars) was observed. Data represents means of measurements from >40 individuals from three independent experiments. Bars represent standard error.Open in a separate windowFigure 2An illustrative model of the sensing and response by the plant root when grown under different saline conditions. This model proposes two major mechanisms of salt responses by plants, where salt tolerance is the ability to function while stressed; Salt avoidance is the capacity to stay away from salt stress when growing in changing saline conditions.Another important point that we would like to bring out based on our observation in this work is that salinity also stimulated shoot positive gravitropism or negative phototropism. The observation implicates long-distance communication from root to shoot during plant salt response in the stressed plants. The exact biological function of shoot tropic growth, the signals in this long-distance communication, and underlying molecular mechanism still remains unknown.In conclusion, our study has revealed a novel complex adaptive mechanism that provides plants a capacity for avoiding injury from salt. The hypothesis we have proposed here should provide novel insights into plant stress avoidance. Further analysis using a combinatorial approach, mutant analysis and genomics, is required to decipher the molecular network underlying this salt-avoidance behavior.     

Linking protein kinase CK2 and auxin transport

Studies performed in different organisms have highlighted the importance of protein kinase CK2 in cell growth and cell viability. However, the plant signaling pathways in which CK2 is involved are largely unknown. We have reported that a dominant-negative mutant of CK2 in Arabidopsis thaliana shows phenotypic traits that are typically linked to alterations in auxin-dependent processes. We demonstrated that auxin transport is, indeed, impaired in these mutant plants, and that this correlates with misexpression and mislocalization of PIN efflux transporters and of PINOID. Our data establishes a link between CK2 activity and the regulation of auxin homeostasis in plants, strongly suggesting that CK2 might be required at multiple points of the pathways regulating auxin fluxes.Key words: protein kinase CK2, root development, auxin, PIN, PINOIDThe plant hormone auxin plays critical roles in plant growth and development.¹ The most abundant natural auxin is the indol-3-acetic acid (IAA), which is synthesized in young apical tissues and then transported to the growing zones of the stem and root. The major route for long distance IAA movement is via the vascular tissue, but, additionally, a slower transport via cell-to-cell (called polar transport) is critical to generate auxin gradients within tissues. Formation of correct auxin gradients is thought to be essential for many plant developmental processes.² In recent years, the IAA transporters have been identified, establishing the molecular basis to understand how auxin transport is regulated. In particular, the identification of the family of plasma-resident PIN proteins, the members of which function as IAA efflux carriers, and the knowledge of their polar localization in the plasma membrane (PM), contributed to generate models predicting the direction of IAA fluxes.^3,4The factors that govern PIN targeting to a particular membrane domain are still not understood. It is known that PIN proteins constitutively undergo cycles of exocytosis and endocytosis to and from the PM, using distinct sorting and recycling endosome trafficking pathways.^5–7 Phosphorylation/dephosphorylation by the Ser/Thr kinase PINOID (PID) and the protein phosphatase 2A, respectively, controls PIN proteins apical/basal localization at the PM, via the GNOM-mediated vesicle trafficking system.⁸ Interestingly, PID is a member of the plant AGC kinases, and, as it happens with its mammals AGC counterparts, is activated by a membrane-associated 3-phosphoinositide-dependent kinase (PDK1).⁹ Moreover, a functional similarity between PIN polar localization in response to auxin and glucose receptor (GLUT4) asymmetrical distribution in response to insulin, has been pointed out.¹⁰ In both cases, cargo proteins (GLUT4 and PIN, respectively) are transported from endosomal vesicles to PM and the process is mediated by PDK1-activated AGC kinases.Protein kinase CK2 is a Ser/Thr kinase evolutionary conserved in eukaryotes, which plays key roles in cell survival, cell division and other cellular processes. A loss-of-function mutant of CK2 in Arabidopsis, obtained by overexpression of a CK2α-inactive subunit, confirmed the essential role of this protein kinase for plant viability.¹¹ Moreover, CK2mut plants showed a dramatic decrease of lateral root formation, inhibition of root growth and overproliferation of root hairs. We have further demonstrated that auxin transport is impaired in this plants, which is concomitant with missexpression of most of the PM-resident PIN proteins, and of PID.¹² In addition, PIN proteins accumulated in endosomal vesicles and auxin gradients were disturbed, both in roots and shoots of CK2mut plants. In particular, root columella cells were depleted of auxin, although the maximum at the quiescent center was unchanged. Starch granule staining with lugol revealed that columella cells retained their fate, although their organization and/or cell shape were clearly affected (Fig. 1).Open in a separate windowFigure 1Lugol-stained starch granules in uninduced (−Dex) and Dex-induced (+Dex) CK2mut roots. In the central part of the figure, a sketch of the main morphogenetic characteristics of mutant roots (right plantlet) as compared to wild-type roots (left plantlet) is shown. Note the shorter roots, wavy phenotype, absence of lateral roots and overproliferation of root hairs in mutant plants.Our results strongly suggest that CK2 is a regulator of auxin-dependent responses, most likely by participating in the regulation of auxin transport. Strikingly, depletion of CK2 activity inhibits some auxin-dependent physiological responses whereas it enhances others. For instance, whereas shoot phototropism was completely absent, root gravitropism was enhanced.¹² Figure 2 shows a time-course of DR5rev::GFP-derived signal after changing the gravity vector, in mutant and control Arabidopsis roots. The progressive auxin translocation to the lower side of the root after gravistimulation is more rapid and sustained in mutant than in control roots, which is likely responsible for the enhanced response to gravity found in mutant roots. Based on these results, we postulate that CK2 might act at different points of the auxin-induced regulatory pathway. As far as is known, the core module that regulates auxin transport is constituted by the protein kinase PID and a protein of the NPH3-domain family. NPH3-containing proteins play important roles in phototropic and gravitropic responses, and regulate polarity and endocytosis of PIN proteins.¹³ As has been proposed by other authors, the participation of one AGC kinase and one NPH3-like protein upstream of an ARF factor might be a common theme in response to different stimulus that are signaled by auxin.¹⁴ We propose that one of the functions of CK2 is the regulation of the activity of core proteins (Fig. 3). Mammalian AGC kinases are well known substrates of CK2 and CK2-dependent phosphorylation is critical for a full display of their activity. The PID and the NPH3-containing protein sequences contain numerous acidic-based motifs that are predicted CK2 phosphorylation sites. Moreover, according to Arabidopsis phosphoproteome databases, several members of the NPH3-containing protein family are predicted to be phosphorylated.¹⁵ In addition, we do not discard the possibility that other proteins involved in PIN transport might also be regulated by CK2-dependent phosphorylation. Experiments are in progress in our laboratory to assess the regulatory role of CK2 in auxin transport.Open in a separate windowFigure 2Time course of auxin relocation during root gravitropic response, as visualized by DR5rev::GFP fluorescence. Root pictures were taken at the indicated times after changing the direction of the gravity vector. Translocation of auxin to the lower part of the root is more rapid in Dex-induced CK2mut plants. Arrows indicate asymmetrical DR5::GFP fluorescence.Open in a separate windowFigure 3Proposed model for the role of CK2 in regulating auxin transport. The core module that regulates auxin transport (shown here as a black box) is constituted by the protein kinase PID and a protein of the NPH3-domain family. PID regulates apical-basal targeting of PIN proteins, by phosphorylating conserved Ser residues present in PIN hydrophilic loops.¹⁶ On the other hand, the family of NPH3-containing proteins regulates polarity and endocytosis of PIN proteins.¹³ There is also a functional similarity between the intracellular transport of PIN proteins and that of the glucose receptor (GLUT4),¹⁰ two processes that are signaled by AGC kinases. We propose that CK2 might be a regulator of the activity of the core proteins, by phosphorylating either the AGC kinase and/or the NPH3-containing protein. Mammalian CK2 is a known regulator of the activity of AGC kinases and other proteins participating in signaling pathways, such as in the Wnt/β-catenin signaling pathway.¹⁷     

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.     

Efflux of hydraulically lifted water from mycorrhizal fungal hyphae during imposed drought

Apart from improving plant and soil water status during drought, it has been suggested that hydraulic lift (HL) could enhance plant nutrient capture through the flow of mineral nutrients directly from the soil to plant roots, or by maintaining the functioning of mycorrhizal fungi. We evaluated the extent to which the diel cycle of water availability created by HL covaries with the efflux of HL water from the tips of extramatrical (external) mycorrhizal hyphae, and the possible effects on biogeochemical processes. Phenotypic mycorrhizal fungal variables, such as total and live hyphal lengths, were positively correlated with HL efflux from hyphae, soil water potential (dawn), and plant response variables (foliar ¹⁵N). The efflux of HL water from hyphae was also correlated with bacterial abundance and soil enzyme activity (P), and the moistening of soil organic matter. Such findings indicate that the efflux of HL water from the external mycorrhizal mycelia may be a complementary explanation for plant nutrient acquisition and survival during drought.Key words: hydraulic lift, nitrogen, phosphorus, microbial abundance, mycorrhizal hyphae, QuercusIn environments that experience seasonal or extended drought, plant productivity, resource partitioning, and competition are limited by the availability of water and mineral nutrients. One mechanism that is important to whole plant water balance in these environments is hydraulic lift (HL), a passive process driven by gradients in water potential among soils layers. Soil water is transported upwards from deep moist soils and released into the nutrient-rich upper soil layers by root systems accessing both deep and shallow soil layers.¹ HL water may improve the lifespan and activity of fine roots in a wide variety of plant life forms.²Hydraulic lift may also have a second ecological function in facilitating plant nutrient acquisition.² It been hypothesized that HL water could enhance the supply of nutrients to roots through mass flow or diffusion,³ or trigger episodes of soil biotic activity such as microbe-mediated nutrient transformations^4,5 that are analogous to the increased inflow of nitrogen (N) into roots and flushes of carbon (C) and N mineralization respectively that follow precipitation events.^4,6 However, few data currently exist with which to test these possibilities.Hydraulically lifted water also sustains mycorrhizal fungi,^7,8 a mutualism that enhances the acquisition of water and mineral nutrients in many terrestrial plant species. Mycorrhizal fungal hyphae provide comprehensive exploration and rapid access to small-scale or temporary nutrient flushes that may not be available to plant roots.⁹ This resource flow has often been assumed to be a unidirectional flux whereby resources are moved from source (soil) into the sink (plant) by the fungal hyphae. However, there is now evidence to suggest that the physiological plasticity of the peripheral extramatrical hyphae, and in particular the hyphal tips, permits the exudation, and subsequent reabsorption, of water and solutes.^10,11 Laboratory experiments using pure cultures have demonstrated that water may be exuded from the hyphal tips, especially in fungal species with hydrophobic hyphae, along with a variety of organic molecules, such as free amino acids.^10–13 At the same time, water, mobile minerals, amino acids and other low-molecular weight metabolites may be selectively and actively reabsorbed by mycorrhizal fungal hyphae.¹¹ However, quantitative data on the environmental impact of hyphal exudation and reabsorption is still largely lacking.We ask: could the diel cycle of water availability created by HL produce a water efflux from hyphal tips and if so, would this be sufficient to impact biogeochemical processes? Is there also an opposite rhythm driven by plant transpiration so that any resultant soil solution is pulled towards hyphal tips and consequently, the host plant? By imposing drought on seedlings of Quercus agrifolia Nee (coast live oak; Fagaceae) grown in mesocosms (Fig. 1), we identified a composite of feedbacks that could influence nutrient capture with HL (Fig. 2). Our analyses provide support for the key predictions of the HL-nutrient cycling scenario including the efflux of HL water from the extramatrical hyphae (Fig. 3), moistening of soil organic matter (Figs. 3 and ​and4),4), and the maintenance of soil microbial activity and nutrient capture (N, P; Open in a separate windowFigure 1Quercus mesocosms demonstrating the plant, root, and hyphal compartments. Details of soil conditions, plant inoculation protocol, mycorrhizal fungi and dye injection methods are detailed in previous work (ref. 7) Point 1 (tap root compartment) denotes the region in which fluorescent tracer dyes were injected into the mesocosm at dusk to track the path of HL water. Point 2 (hyphal chamber) denotes spots adjacent to or distant from the mesh screen into which a small volume (200 µl) of fluorescent and ¹⁵N tracers (99% as ¹⁵NH₄¹⁵NO₃) were injected at dawn to measure water and nutrient uptake by the external hyphae.Open in a separate windowFigure 2Path analysis of the influence of different soil and mycorrhizal factors on nutrient capture with HL, and resultant model showing the significant path coefficients among variables in the Q. agrifolia mesocosms. Lines with a single arrow denote possible cause-effect relationships. The partial correlation coefficients adjacent to each line indicate the strength of the association between the individual factors. Thick lines are statistically significant (p < 0.05) whereas thin lines indicate no significant relationship between parameters (p > 0.05) and only significant coefficients are given (p < 0.05).Open in a separate windowFigure 3Fluorescently-labeled structures recovered from the hyphal chamber of Quercus microcosms following 80 days of soil drying and with nocturnal hydraulic lift. Yellow-green fluorescence indicates samples labeled with Lucifer yellow CH (LYCH), blue fluorescence denotes samples labeled with Cascade blue (CB) hydrazide. (A) CB-labeled leaf litter from the soil and (B) soil particle; (C) LYCH-labeled root fragment in the soil mixture with adherent extramatrical hyphae; (D) LYCH tracer dye fluorescence in labeled extramatrical hyphae and in efflux (arrow) from the hyphal tip onto organic matter; (E and F) external hyphae filled with LYCH (influx; arrow) and (G) background fluorescence in non-labeled extramatrical hyphae.Open in a separate windowFigure 4Measurements of hyphal efflux and influx based on the quantitative analysis of LYCH fluorescence intensity in soil solution. Fluorescent intensity values were converted to LYCH concentration using a standard curve generated for the dye since fluorescent intensity correlates with the number of fluorescent molecules in solution. Influx is the uptake of LYCH by hyphae as driven by plant transpiration demands (day), and measured efflux is the passive loss of LYCH from hyphae into the surrounding soil during HL (night). Vertical bars indicate the standard error of the means.

Table 1

Summary of soil, microbial, mycorrhizal and plant parameters in plant or hyphal compartments

A systems biology approach to dissect the contribution of brassinosteroid and auxin hormones to vascular patterning in the shoot of Arabidopsis thaliana

Systems biology can foster our understanding of hormonal regulation of plant vasculature. One such example is our recent study on the role of plant hormones brassinosteroids (BRs) and auxin in vascular patterning of Arabidopsis thaliana (Arabidopsis) shoots. By using a combined approach of mathematical modelling and molecular genetics, we have reported that auxin and BRs have complementary effects in the formation of the shoot vascular pattern. We proposed that auxin maxima, driven by auxin polar transport, position vascular bundles in the stem. BRs in turn modulate the number of vascular bundles, potentially by controlling cell division dynamics that enhance the number of provascular cells. Future interdisciplinary studies connecting vascular initiation at the shoot apex with the established vascular pattern in the basal part of the plant stem are now required to understand how and when the shoot vascular pattern emerges in the plant.Key words: Arabidopsis, vascular, auxin, brassinosteroids, mathematical model, computer simulationsThe plant vascular system is responsible for the long-distance transport of water, solutes and molecules throughout the plant, being essential for plant growth and development. It is formed by two different functional tissues: the xylem, which transports water from roots to aerial organs, and the phloem, through which nutrients and photosynthetic products and signaling molecules are transported.During embryogenesis, the vasculature is characterized as an undifferentiated procambial tissue in the innermost part of the plant embryo.¹ Later in development, the procambium (i.e., a group of pluripotent stem cells²) begins to divide and differentiate into xylem and phloem tissues through oriented cell divisions. In the shoot, procambium generates xylem tissue centripetally and phloem tissue centrifugally, driving the formation of collateral vascular bundles around it.^3,4 In the inflorescence stem of the model plant Arabidopsis, the radial pattern of the vasculature exhibits a periodic organization made by the alternation of vascular bundles and interfascicular fibers, which altogether form the vascular ring (Fig. 1A).Open in a separate windowFigure 1Vascular patterning in Arabidopsis shoot inflorescence stem. (A) Radial section of DR5::GUS expression at the base of the inflorescence stem in Arabidopsis Col-0 plants. (B) Computer simulation result for auxin concentration ([Auxin]) in arbitrary units (a.u.) along a ring of cells; x and y stand for spatial coordinates. Auxin is distributed in maxima which, according to the model hypothesis, position vascular bundles. (C) Longitudinal section of Arabidopsis Col-0 wild-type plant at the most apical zone, immediately below the shoot apical meristem. Arrows point to xylem strains coming from the lateral organs.Previous studies have documented the importance of plant hormones such as auxin and BRs in vascular cell differentiation and patterning.⁵ Defective polar auxin transport distorts shoot vascular patterning^6,7 and BR loss-of-function mutants exhibit few vascular bundles.^8,9 But how do these hormones control shoot vascular patterning? In order to answer this question, we used both quantitative measurements of vascular phenotypes and computational modeling.¹⁰     

Focus Issue on Roots: Branching Out in Roots: Uncovering Form,Function, and Regulation

Root branching is critical for plants to secure anchorage and ensure the supply of water, minerals, and nutrients. To date, research on root branching has focused on lateral root development in young seedlings. However, many other programs of postembryonic root organogenesis exist in angiosperms. In cereal crops, the majority of the mature root system is composed of several classes of adventitious roots that include crown roots and brace roots. In this Update, we initially describe the diversity of postembryonic root forms. Next, we review recent advances in our understanding of the genes, signals, and mechanisms regulating lateral root and adventitious root branching in the plant models Arabidopsis (Arabidopsis thaliana), maize (Zea mays), and rice (Oryza sativa). While many common signals, regulatory components, and mechanisms have been identified that control the initiation, morphogenesis, and emergence of new lateral and adventitious root organs, much more remains to be done. We conclude by discussing the challenges and opportunities facing root branching research.Branching through lateral and adventitious root formation represents an important element of the adaptability of the root system to its environment. Both are regulated by nutrient and hormonal signals (Bellini et al., 2014; Giehl and von Wirén, 2014), which act locally to induce or inhibit root branching. The net effect of these adaptive responses is to increase the surface area of the plant root system in the most important region of the soil matrix for resource capture (e.g. surface layers for phosphorus uptake and deeper layers for nitrate uptake) or to secure anchorage. Different species use different combinations of lateral or adventitious roots to achieve this, with lateral roots dominating the root system of eudicots while adventitious (crown and brace) roots dominate the root system of monocots, including in cereal crops.Our understanding of the mechanisms controlling lateral and adventitious root development has accelerated in recent years, primarily through research on model plants. The simple anatomy and the wealth of genetic resources in Arabidopsis (Arabidopsis thaliana) have greatly aided embryonic and postembryonic root developmental studies (De Smet et al., 2007; Péret et al., 2009a; Fig. 1, A and E). Nevertheless, impressive recent progress has been made studying root branching in other crop species, notably cereals such as maize (Zea mays) and rice (Oryza sativa).Open in a separate windowFigure 1.A to D, Schematics showing diversity in root system architecture at both seedling (left) and mature (right) stages in eudicots (A and C) and monocots (B and D). A, Arabidopsis root system. B, Maize root system. C, Tomato root system (for clarity, stem-derived adventitious roots are only shown in the labeled region). D, Wheat root system. E and F, Cross sections of emerging lateral root primordia in Arabidopsis (E) and rice (F). E and F are adapted from Péret et al. (2009b).In this Update, we initially describe the diversity of postembryonic root forms in eudicots and monocots (Fig. 1). Next, we highlight recent advances in our understanding of the genes, signals, and mechanisms regulating lateral root and adventitious root branching in Arabidopsis, rice, and maize. Due to space limits, we cannot provide an exhaustive review of this subject area, focusing instead on recent research advances. However, we direct readers to several recent in-depth reviews on lateral root (Lavenus et al., 2013; Orman-Ligeza et al., 2013) and adventitious root development (Bellini et al., 2014).     

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.     

Negative effects of desiccation on the protein sorting and post-translational modification

Bryophytes as the first land plants are believed to have colonized the land from a fresh water origin, requiring adaptive mechanisms that survival of dehydration. Physcomitrella patens is such a non-vascular bryophyte and shows rare desiccation tolerance in its vegetative tissues. Previous studies showed that during the course of dehydration, several related processes are set in motion: plasmolysis, chloroplast remodeling and microtubule depolymerization. And proteomic alteration supported the cellular structural changes in respond to desiccation stress.¹ In this addendum, we report that Golgi bodies are absent and adaptor protein complex AP-1 large subunit is downregulated during the course of dehydration. Those phenomena may be adverse in protein posttranslational modification, protein sorting and cell walls synthesis under the desiccation condition.Key words: AP-1 protein, cell ultrastructure, desiccation, golgi bodies, physcomitrella, proteomeThe plant Golgi apparatus is composed of many small stacks of cisternae, sometimes known as dictyosomes. The Golgi is a complex polarized organelle consisting of both a cis and trans side, containing compartments with functionally different capacities for directing cellular components. The plant Golgi apparatus synthesis a wide range of cell wall polysaccharides and proteoglycans, and also carries out O-linked glycosylation and N-linked glycan processing.^2–5 Moreover, the Golgi is involved in returning escaped proteins back to the endoplasmic reticulum, sorting of proteins and polysaccharides to the cell wall or vacuoles, and in organizing the compartmentation of its own enzymes by retention or retrieval mechanisms.⁶ In conclusion, The Golgi apparatus is central to the growth and division of the plant cell through its roles in protein glycosylation, protein sorting and cell wall synthesis.The transit of proteins and lipids from the trans-Golgi network (TGN) and the plasma membrane to endosomes within eucaryotic cells occurs via the budding and fusion of clathrin-coated vesicles (CCVs).^7,8 At the TGN, this process is mediated by the heterotetrameric AP-1 adaptor complex, which consists of two large subunits, β and γ1; a medium subunit, µ1; and a small σ1 subunit. Recruitment of AP-1 to the TGN membrane is regulated by a small GTPase, ADP-ribosylation factor 1 (ARF1), which cycles between an inactive GDP-bound form in cytosol and an active GTP-bound form that associates with the membrane like other small GTPase.⁹ There is also evidence that phosphorylation/dephosphorylation events are involved in the regulation of the function of AP-1. Ghosh and Kornfeld demostrated that AP-1 recruitment onto the membrane is associated with protein phosphatase 2A (PP2A)-mediated dephosphorylation of its β1 subunit, which enables clathrin assembly. This Golgi-associated isoform of PP2A exhibits specificity for phosphorylated β1 compared with phosphorylated µ1. Once on the membrane, the µ1 subunit undergoes phosphorylation, which results in a conformation change. This conformational change is associated with increased binding to sorting signals on the cytoplasmic tails of cargo molecules. Dephosphorylation of µ1 (and µ2) by another PP2A-like phosphatase reversed the effect and resulted in adaptor release from CCVs. Cyclical phosphorylation/dephosphorylation of the subunits of AP-1 regulate its function from membrane recruitment until its release into cytosol.¹⁰Plants experience desiccation stress either as part of a developmental programme, such as during seed maturation, or because of reductions in air humid and water availability in the soil. Underlying the ability of bryophytes to withstand periods of desiccation are morphological and biochemical adaptations. Plants respond to stress as individual cells and synergistically as a whole organism. Scanning electron microscopy observation showed that the P. patens gametophore cells were shrunk upon the treatment of desiccation, and the shrinking started from the edge of the leaves (Fig. 1). We could clearly observe some dark granula in the untreated cells, but these granula disappeared post-desiccation treatment (Fig. 1). Transmission electron microscopy also revealed that the large stacks of Golgi bodies and numerous coated vesicles are typically visible in the hydrated cells (Fig. 2), but these are absent in the desiccative cells (data not shown). The plant Golgi apparatus plays an important role in protein glycosylation and sorting. Therefore, this event means that the protein sorting and the cargo transporting are disrupted by desiccation stress. During desiccation, the absentness of Golgi bodies reduce the leaf activities of cell, and this is expected to similar to plant dormancy which is a phenomenon in resurrection plants and some drought-tolerant plants. In addition, through two-dimensional gel electrophoresis (2-DE) and LC-MS/MS analysis, AP-1 large subunit was identified as downregulated protein during the course of dehydration (Fig. 3). AP-1 is ubiquitously expressed and participates in the budding of clathrin-coated vesicles from the trans-Golgi network (TGN) and endosomes. AP-1 also recognizes sorting motifs in cargo molecules. Our results suggested that desiccation led to a marked disrupt in protein posttranslational modification, protein sorting and cell walls synthesis.Open in a separate windowFigure 1Scanning Electron microscopy images of normal and dehydrated P. patens gametophores. (A) the fresh leaf; (B) enlargement of the rectangle area of (A); (C) dehydrated gametophores of P. patens. Bar = 5 µm.Open in a separate windowFigure 2Transmission electron microscopy images of cell in fresh game-tophores. The arrows indicate Golgi body, Bar = 2 µm.Open in a separate windowFigure 3Part protein profile of the control and desiccation plants. The arrows indicate the AP-1 large subunit.     

FZR2/CCS52A1 mediated endoreduplication in Arabidopsis development

Plant organogenesis generally involves three basic processes: cell division, cell expansion and cell differentiation. Endoreduplication, a process of genome replication without intervening mitosis, often occurs during cell expansion and cell differentiation. The switch from the mitotic cell cycle to the endocycle, however, is still poorly understood in plants. We have recently demonstrated that FIZZY-RELATED2 (FZR2) is a factor controlling endoreduplication in Arabidopsis. fzr2 mutants lacked gross morphological defects but showed a general decrease of endoploidy level in trichomes and other leaf cells, while expression of FZR2 under constitutive or tissue specific promoters induced extra or ectopic endoreduplication in all tissues examined. We also showed that decrease of leaf cell size in fzr2 mutants could be compensated by increased cell proliferation. In this addendum, we discuss additional phenotypes of FZR2 misexpression, including apparent mosaic leaf sectors in which local cell overexpansion due to 35S::FZR2 appears to be compensated by reduced cell expansion in neighboring tissues.Key words: Arabidopsis, fizzy-related, CCS52A1, endoreduplication, embryo, mosaic analysis, compensation mechanismPlants begin vegetative development as single zygotes and reach final sizes million-fold bigger, with complex tissues and cell types. During development, plants undergo three basic processes: cell division, cell expansion and cell differentiation. While cell division is dependent on mitotic cell cycle, cell expansion and cell differentiation are often coupled with a modified cell cycle called endoreduplication.¹ Endoreduplication enables a cell to increase its ploidy by replicating its genome without subsequent chromosomal and cellular division. This endocycle, widespread in eukaryotes but especially common in plants, may provide individual cells with the gene-expression capacity to reach larger sizes.² In the well-studied dicotyledon model Arabidopsis thaliana, endoreduplication occurs in most of the differentiated cell types, such as trichoblasts and trichomes, or cells with very high metabolic activity, such as the endosperm.³ The switch from mitotic cycles to endocycles requires cells to start another round of DNA replication without intervening mitosis. Therefore, a cell must induce re-entry into S-phase after G₁-phase while inhibiting M-phase. The regulatory mechanisms mediating the G₁ to S transition in endocycles share components of the mitotic pathways,⁴ with the CDK/CYCLIN B complex influencing DNA replication.⁵ Much evidence in yeast, fly and plants has pointed to the involvement of a WD-40 protein, Fizzy-Related/Ccs52, in triggering the switch to endoreduplication by controlling the degradation of Cyclin B.^6,7Using reverse genetics in Arabidopsis, we investigated FIZZY-RELATED2 loss-of-function mutants.⁸ fzr2 plants showed reduced endoreduplication and cell size in both pavement cells and trichomes.⁸ When FZR2 was misexpressed with the CMV 35S promoter, transgenic plants showed a range of phenotypes such as retarded growth, supernumerary trichome branches and distorted roots, with ectopic endoreduplication induced in all examined tissues. When expressed under control of the petal- and stamen-specific APETELA3 promoter, FZR2 caused great increases in the cell and nuclear sizes of petal and stamen cells, which normally endocycle little or not at all in Arabidopsis.⁸Since AP3 also drives gene expression in pollen, and pollen mother cells undergo two rounds of meiosis to generate haploid sperm cells,⁹ the effects of FZR2 expression on male gametogenesis seemed particularly interesting. Microscopic analysis showed larger pollen grains in AP3::FZR2 plants relative to wildtype, whereas DAPI staining revealed a concomitant increase in sperm cell nuclear size (Fig. 1A–D). These results suggested that endoreduplication had been induced in these pollen grains. Although these polyploid sperm cells proceeded through double fertilization, the corresponding embryos failed to complete development. Examination of cleared embryos with Nomarski microscopy showed that about half of them stopped growth at the torpedo stage (Fig. 1G and H), possibly due to abnormal endosperm development. When endosperm cellularization was completed in wildtype seeds (Fig. 1E), there were only 2 to 3 bubble-like structures at the chalazal poles of developing AP3::FZR2 seeds (Fig. 1F). This phenotype was similar to that of developing seeds derived from fertilization of a diploid plant with pollen from an hexaploid plant,¹⁰ further supporting the conclusion that AP3::FZR2 sperm cells underwent endoreduplication.Open in a separate windowFigure 1Comparisons of pollen grain sizes, nuclear sizes and embryo development among wildtype (WT, left: A, C, E and G) and AP3::FZR2 lines (right: B, D, F and H). (A and B) Micrographs of representative pollen grains. (C and D) DAPI staining of representative pollen grain. Arrowheads in (C and D) indicate the enlarged nuclei of sperm cells. (E and F) Micrographs of heart-stage embryos. (G and H) Micrographs of torpedo-stage embryos. Arrowheads in (F and H) indicate the abnormal endosperms. In (E–H), seeds were cleared with Hoyer solution and viewed using Nomarski optics. Scale bars represent 10 µm in (A–D), and 100 µm in (E–H).Another interesting result of this study was the different manner in which stamens and petals were altered by AP3::FZR2 expression. While petal cells showed extreme increases in size and decreases in numbers, the organs became disrupted, losing their characteristic laminar shape. Conversely, AP3::FZR2 stamens maintained their cylindrical shape, despite becoming wider at the organ level and composed of larger cells.⁸ This discrepancy in the severity of petal and stamen organ-level phenotypes may be because the two tissues respond differently to FZR2 misexpression, or because the shapes of these two organs place unique constraints on the effects of cell overgrowth. Like these stamens, roots and stems of 35S::FZR2 plants also retained normal shape despite severe distortion of internal tissue architecture.⁸ Perhaps a cylindrical organ is maintained more easily due to the dynamics of biophysical forces. It is also possible that the morphogenesis of a filamentous structure makes more use of intercellular communication than a laminar structure, so the cell proliferation and cell expansion are more strictly regulated by non-cell autonomous signals such as protein movement via plasmodesmata to provide additional positional information.¹¹ The regulatory contribution of these additional signals may override the effects of FZR2 ectopic expression.Finally, the most intriguing phenotype found in fzr2-1 mutant was that the overall leaf size showed no significant difference compared with wildtype, although the average cell was smaller. This suggests that proliferation is enhanced to generate more cells in response to the decreased average cell size. A mechanism called compensation is postulated to coordinate cell proliferation and cell expansion to attain proper organ size.¹² For example, mutations or transgenes that cause decreases in leaf cell proliferation can be compensated by extra leaf cell expansion, such that the organ approaches normal size.¹³ Little is known, however, about how organs and cells respond to local perturbations of cell sizes. In a subset of 35S::FZR2 transgenic plants, the expression of FZR2 was silenced at the whole plant level, but some groups of cells escaped silencing. These sectors showed FZR2 overexpression phenotypes such as over-branched trichomes and giant pavement cells, whereas nearby sections of the same leaf contained normal-sized pavement cells and 3- or 4-branch trichomes. These mosaic sectors provided an opportunity to observe how compensation works even within an organ. Inside the sectors were overgrown pavement cells typical of some FZR2 overexpression lines (Fig. 2A). Away from the sectors, the pavement cells were wildtype in appearance (Fig. 2C and D). At the sector boundary, however, a strip of very small cells formed (Fig. 2B). The smaller cell size at the border may have came about to compensate for the abnormally large cells within the sector, although it is unclear whether this decrease in cell size was followed reduced endoreduplication or simple space limitation.Open in a separate windowFigure 2Comparisons of cell sizes inside and outside of a mosaic sector. (A) Scanning electron microscope graphs of epidermal cells from a mosaic sector of 35S::FZR2. (B) Epidermal cells at boundary region between mosaic sector (OE) and surrounding normal cells (N). Black lines highlight the band of smaller cells. (C) Normal epidermal cells outside the mosaic sector in the same leaf (N). (D) Epidermal cells from wildtype plants (WT). Scale bar represents 100 µm.By studying fzr2 mutants and misexpression lines, we showed that FZR2 is necessary and sufficient to induce endoreduplication in various cell types. Our observation that cells increase proliferation to compensate the decreased cell size in fzr2 mutants provides important evidence that cell proliferation and cell expansion are closely interconnected to regulate organ development in Arabidopsis. Further experiments such as mosaic analysis are needed to further elucidate the compensation mechanism.     

Plant small monomeric G-proteins (RAC/ROPs) of barley are common elements of susceptibility to fungal leaf pathogens,cell expansion and stomata development

Small monomeric RAC/ROP GTPases act as molecular switches in signal transduction processes of plant development and stress responses. They emerged as crucial players in plant-pathogen interactions either by supporting susceptibility or resistance. In a recent publication, we showed that constitutively activated (CA) mutants of different barley (Hordeum vulgare) RAC/ROPs regulate susceptibility to barley fungal leaf pathogens of different life style in a contrasting way. This illustrates the distinctive signalling roles of RAC/ROPs for different plant-pathogen combinations. We also reported the involvement of RAC/ROPs in plant epidermis development in a monocotyledonous plant. Here we further discuss a failure of CA HvRAC/ROP-expressing barley to normally develop stomata.Key words: Hordeum vulgare, G-proteins, RAC, ROP, cell expansion, stomata, transpirationMembers of the RHO family of small G-proteins in plants (RAC/ROPs) regulate signal transduction processes at the plasma membrane.¹ They act as multifunctional signalling switches in plant development and a variety of stress responses. RAC/ROP GTPases play regulatory roles in polar growth and cell morphogenesis in several cell systems including pollen tubes, developing root hairs and leaf pavement cells.²In a recent publication,³ we showed that constitutively activated (CA) mutants of different barley (Hordeum vulgare) RAC/ROPs support susceptibility to the barley powdery mildew fungus Blumeria graminis f.sp. hordei (Bgh). CA HvRAC1 supported susceptibility to biotrophic Bgh but resistance to hemibiotrophic Magnaporthe oryzae in barley at the penetration level in both cases. Additionally, CA HvRAC1 supported local callose deposition at sites of attack from Bgh and a secondary H₂O₂ burst in whole non-penetrated epidermal cells. This supports a regulatory function of RAC/ROPs in plant defence¹ and the potential corruption of defence pathways in susceptibility to Bgh. Because the rice ortholog of HvRAC1, OsRAC1, can regulate an H₂O₂ burst via activation of the plasma membrane NADPH oxidase OsRBOHB,⁴ one can speculate that the secondary H₂O₂ burst in CA HvRAC1 barley could also be caused by over-activation of an NADPH oxidase. However, CA HvRAC1 barley was also more susceptible to fungal penetration, and penetrated cells did not show an H₂O₂ burst. Hence, CA HvRAC1 did not contribute to penetration resistance, and the H₂O₂ burst might have been suppressed by Bgh after successful penetration. Interestingly, Bgh secretes a catalase during interaction with the plant.⁵The involvement of RAC/ROPs in plant development has been widely studied in the dicots Arabidopsis and tobacco. In Arabidopsis, CA AtRAC/ROPs disturb root hair tip growth and epidermal cell morphogenesis.^6,7 We showed similar developmental aberrations as a result of CA HvRAC/ROP expression in monocotyledonous barley. Root hair polarity disruption and enhanced leaf epidermal cell expansion was observed in CA HvRAC/ROP expressing barley. Here, we further report on reduced or abnormal development of stomata as an effect of CA HvRAC/ROP expression.In barley, stomata and short epidermal cells alternate in a row of leaf epidermal cells (Fig. 1A). The number of stomata number was significantly reduced in three CA HvRAC/ROP (CA HvRACB, CAHvRAC3, CA HvRAC1) expressing barley genotypes when compared to azygous controls (barley siblings that lost the transgene due to segregation) (Fig. 1E). In part, this could be explained by enhanced length of epidermal cells intercalated between stomata (Fig. 1B). The presence of longer epidermal cells in all CA HvRAC/ROP-barleys further supports that RAC/ROPs are operating in epidermal cell expansion.³Open in a separate windowFigure 1Stomatal abnormalities observed in CA HvROPexpressing transgenic barley leaves. (A) Wild type leaf adaxial epidermis with alternating stomata complexes (arrows) and short epidermal cells (asterisk). (B) Presence of more than one short epidermal cell in between two stomata. Arrows point the stomata. Double headed arrows highlight intercalated cells with enhanced cell length (C) Two stomata lacking an intercalated short epidermal cell. (D) Stoma failed to develop and left an abnormal blank cell. (E) Average number of stomata present in 5 cm of a stomatal row in transgenic plants expressing distinct CA barley CA HvRAC/ROPs. For all samples, stomatal rows present on either side of the mid rib were counted in the leaf upper epidermis. Fully expanded leaves of 3-weeks-old barley plants were used for counting stomata. Error bars show 95% confidence intervals. Repetition of the experiment led to similar results. Scale bars = 50 µm.Previously, we carried out porometry experiments to measure stomata conductivity in CA HvRACB expressing barley leaves.⁸ The CA HvRACB leaves showed up to 50% less transpiration than azygous controls without any treatment. Additionally, CA HvRACB leaves were less responsive to abscisic acid (ABA) and subsequently they could not effectively reduce transpiration when treated with ABA or when cut-off from water supply.⁸ Our data on numbers of stomata per leaf segment could now explain the lower rates of transpiration in non-stressed CA HvRACB barley when compared to wild type.Apart from the stomata number, developmental abnormalities were observed in the arrangement of epidermal cells. Generally, the shape of epidermal cells was less regular in CA HvRAC/ROP barley.³ We also observed the presence of more than one short epidermal cell in between two stomata (Fig. 1B) or two stomata lacking an intercalated short epidermal cell (Fig. 1C), or stomata failed to develop, which ended up in an abnormally short epidermal cell (Fig. 1D). Although such abnormalities were also rarely observed in wild type plants, all three CA HvRAC/ROP-barley leaves exhibited a clearly higher frequency of abnormalities in a given length of a stomata row. Together, CA HvRAC/ROPs had an effect on both the number and development of stomata. These observations suggest that RAC/ROPs are not only operating in cell expansion but also in barley cell differentiation for stomata development.